The Dark Side of Light Traps

The Dark Side of Light Traps Abstract Light-baited suction traps are one of the most widely used tools for vector surveillance. Their popularity stems from ease of use even in remote locations, range and abundance of species caught, and low cost. The availability of smaller, portable models, like the CDC miniature light trap, have further increased their ubiquity in entomological field studies. However, when researchers have looked, light trap collections are usually biased in ways that may affect data interpretation for epidemiological studies. If used alone, light traps may fail to collect important or infected vectors, and light traps are inefficient or ineffective when competing ambient light is present. In this article, we discuss these biases and limitations in terms of their effect on collection efficiency, population data, and pathogen detection. While light trap data certainly have a purpose, an over-reliance on light trapping risks drawing false conclusions about vector populations and vector-borne disease epidemiology. These concerns are especially troubling when light trap data are used to inform policy decisions meant to protect human and animal health. Particularly when a species’ response to light is unknown or poorly characterized, light traps should be used in conjunction with supplemental sampling methods. Researchers conducting vector surveillance field studies should carefully consider their study design and objectives when deciding on a trapping method or methods, and specifically endeavor to understand the limitations of their data. Only then can researchers take advantage of the best attributes of light traps while avoiding their dark side. light trap, surveillance, vector, trapping Light traps have been used for almost 130 yr to collect a variety of nocturnal and crepuscular insects. In medical and veterinary entomology, they are most often used to collect mosquitoes (Diptera: Culicidae), biting midges (Diptera: Ceratopogonidae), and sand flies (Diptera: Psychodidae: Phlebotominae), but have also occasionally been used to collect black flies (Diptera: Simuliidae), house flies (Musca domestica L.), horn flies (Haemotobia irritans (L.)) (Diptera: Muscidae), fleas (Siphonaptera), and kissing bugs (Hemiptera: Reduviidae: Triatominae) (Bram 1978). Data stemming from light trap collections, including species composition, population age structure, abundance, and infection rate, are sometimes used to inform epidemiological models for vector-borne diseases, and light traps can be useful in that respect, as we discuss below. However, there also is a growing body of literature describing the ways in which light traps can fail to accurately represent vector populations or pathogen activity in an area. This article discusses some of those issues, in the hopes that researchers may better appreciate caveats regarding the use of light trap data in epidemiological studies, particularly if light traps are used without any supplemental sampling. Light trap limitations range in their severity from relatively minor influences on trap effectiveness (e.g., limited range of attraction), to those that may affect the interpretation of vector populations in an area (e.g., sex biases), all the way to the most serious limitations: failing to collect vector species, or detect pathogens of interest (Table 1). We suggest that the use of light trap data should be tempered by these considerations, and argue that in many cases, alternatives or supplements to light trapping should be sought in order to improve studies on vector biology and vector-borne disease epidemiology. Table 1. Summary of literature discussing the potential biases and drawbacks of light trapping for insect vectors Concern  Vector group  Study  Light source(s) used  I. Limitations of efficiency      Decreased attraction with ambient light  Mosquitoes (Diptera: Culicidae)  Bradley, G.H., and T.E. McNeel. 1935 (J. Econ. Entomol.)  Incandescent      Horsfall, W.R. 1943 (Ann. Entomol. Soc. Am.)  Incandescent      Pratt, H.D. 1948 (J. Natl. Malar. Soc.)  Incandescent      Provost, M.W. 1959 (Ann. Entomol. Soc. Am.)  Incandescent      Barr, A.R., et al. 1963 (J. Econ. Entomol.)  Incandescent      Bidlingmayer, W.L. 1967 (J. Med. Entomol.)  Incandescent      Miller, T.A., et al. 1970 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Bishop, A.L., et al. 2000 (Austr. J. Entomol.)  Incandescent      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV  Limited range of attraction  Mosquitoes (Diptera: Culicidae)  Odetoybino, J.A. 1969 (Bull. Wld. Hlth. Org.)  Incandescent      Constantini, C., et al. 1998 (Bull. Entomol. Res.)  Incandescent, UV    Biting midges (Diptera: Ceratopogonidae)  Venter, G.J., et al. 2012 (Vet. Parasitol.)  UV    Sand flies (Diptera: Psychodidae)  Killick-Kendrick, R., et al. 1985. (Annales de Parasitologie et Humaine Comparee)  Incandescent      Valenta, D.T., et al. 1995. (Proc. II Int. Symp. Phlebot. Sand Flies)  Incandescent      Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified  Attraction to different wavelengths  Mosquitoes (Diptera: Culicidae)  Burkett, D. A., et al. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Bishop, A.L., et al. 2006 (Austr. J. Entomol.)  LED, UV, Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Hope, A., et al. 2015 (Parasites & Vectors)  LED, UV      Gonzalez, M., et al. 2016 (Vet. Parasitol.)  LED, UV, Incandescent    Sand flies (Diptera: Psychodidae)  Hoel, D.F., et al. 2007 (J. Vect. Ecol.)  Incandescent, LED    Kissing bugs (Hemiptera: Reduviidae)  Minoli, S.A., and Lazzari, C.R. 2006 (Acta Tropica)  Incandescent, UV      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  LED, UV, Incandescent    General  Cohnstaedt, L.W., et al. 2008 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED  Small catch size  Mosquitoes (Diptera: Culicidae)  Newhouse, V.F., et al. 1966 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Harrup, L.E., et al. 2012 (J. Med. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified      Andrade, A.J., et al. 2008 (Mem. Inst. Oswaldo Cruz)  Incandescent      Kasap, O.E., et al. 2009 (ACTA Vet. BRNO)  Incandescent    Black flies (Diptera: Simuliidae)  Snoddy, E.L., and K.L. Hays. 1966 (J. Econ. Entomol.)  Incandescent  II. Limitations for population data        Sex biases  Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent  Parity biases  Mosquitoes (Diptera: Culicidae)  Reisen, W.K., and A.R. Pfuntner. 1987 (J. Amer. Mosq. Contr. Assoc.)  Incandescent      Githeko, A.K., et al. 1994 (Bull. Entomol. Res.)  Incandescent      Williams, G.M., and J.B. Gingrich. 2007 (J. Vect. Ecol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Anderson, J.R., and A.X. Linhares. 1989 (J. Amer. Mosq. Cont. Assoc.)  UV      Bellis, G.A., and D.J. Reid. 1996 (Austr. J. Entomol.)  Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV  III. Limitations for pathogen detection      Species composition  Mosquitoes (Diptera: Culicidae)  Morris, C.D., and G.R. Defoliart. 1969 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent      Govella, N.J., et al. 2011 (Parasites & Vectors)  Incandescent      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Carpenter, S., et al. 2008 (J. App. Ecol.)  UV      Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Rutledge, L.C., et al. 1975 (J. Med. Entomol.)  Incandescent      Alexander, B., et al. 1992. (Mem. Inst. Oswaldo Cruz)  Incandescent      Davies, C.R., et al. 1995 (Med. Vet. Entomol.)  Incandescent      Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent      Alten, B., et al. 2015 (Bull. Entomol. Res.)  Not specified  Infection prevalence  Biting midges (Diptera: Ceratopogonidae)  Mayo, C.E., et al. 2012 (Vet. Parasitol.)  UV      McDermott, E.G. et al. 2015 (Parasites & Vectors)  UV    Kissing bugs (Hemiptera: Reduviidae)  Walter, A.I., et al. 2005 (Cademos de Saude Publica)  Not specified      Barghini, A., and B.A.S. de Medeiros. 2010 (Environ. Health Prosp.)  Not specified      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  Incandescent  Concern  Vector group  Study  Light source(s) used  I. Limitations of efficiency      Decreased attraction with ambient light  Mosquitoes (Diptera: Culicidae)  Bradley, G.H., and T.E. McNeel. 1935 (J. Econ. Entomol.)  Incandescent      Horsfall, W.R. 1943 (Ann. Entomol. Soc. Am.)  Incandescent      Pratt, H.D. 1948 (J. Natl. Malar. Soc.)  Incandescent      Provost, M.W. 1959 (Ann. Entomol. Soc. Am.)  Incandescent      Barr, A.R., et al. 1963 (J. Econ. Entomol.)  Incandescent      Bidlingmayer, W.L. 1967 (J. Med. Entomol.)  Incandescent      Miller, T.A., et al. 1970 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Bishop, A.L., et al. 2000 (Austr. J. Entomol.)  Incandescent      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV  Limited range of attraction  Mosquitoes (Diptera: Culicidae)  Odetoybino, J.A. 1969 (Bull. Wld. Hlth. Org.)  Incandescent      Constantini, C., et al. 1998 (Bull. Entomol. Res.)  Incandescent, UV    Biting midges (Diptera: Ceratopogonidae)  Venter, G.J., et al. 2012 (Vet. Parasitol.)  UV    Sand flies (Diptera: Psychodidae)  Killick-Kendrick, R., et al. 1985. (Annales de Parasitologie et Humaine Comparee)  Incandescent      Valenta, D.T., et al. 1995. (Proc. II Int. Symp. Phlebot. Sand Flies)  Incandescent      Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified  Attraction to different wavelengths  Mosquitoes (Diptera: Culicidae)  Burkett, D. A., et al. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Bishop, A.L., et al. 2006 (Austr. J. Entomol.)  LED, UV, Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Hope, A., et al. 2015 (Parasites & Vectors)  LED, UV      Gonzalez, M., et al. 2016 (Vet. Parasitol.)  LED, UV, Incandescent    Sand flies (Diptera: Psychodidae)  Hoel, D.F., et al. 2007 (J. Vect. Ecol.)  Incandescent, LED    Kissing bugs (Hemiptera: Reduviidae)  Minoli, S.A., and Lazzari, C.R. 2006 (Acta Tropica)  Incandescent, UV      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  LED, UV, Incandescent    General  Cohnstaedt, L.W., et al. 2008 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED  Small catch size  Mosquitoes (Diptera: Culicidae)  Newhouse, V.F., et al. 1966 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Harrup, L.E., et al. 2012 (J. Med. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified      Andrade, A.J., et al. 2008 (Mem. Inst. Oswaldo Cruz)  Incandescent      Kasap, O.E., et al. 2009 (ACTA Vet. BRNO)  Incandescent    Black flies (Diptera: Simuliidae)  Snoddy, E.L., and K.L. Hays. 1966 (J. Econ. Entomol.)  Incandescent  II. Limitations for population data        Sex biases  Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent  Parity biases  Mosquitoes (Diptera: Culicidae)  Reisen, W.K., and A.R. Pfuntner. 1987 (J. Amer. Mosq. Contr. Assoc.)  Incandescent      Githeko, A.K., et al. 1994 (Bull. Entomol. Res.)  Incandescent      Williams, G.M., and J.B. Gingrich. 2007 (J. Vect. Ecol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Anderson, J.R., and A.X. Linhares. 1989 (J. Amer. Mosq. Cont. Assoc.)  UV      Bellis, G.A., and D.J. Reid. 1996 (Austr. J. Entomol.)  Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV  III. Limitations for pathogen detection      Species composition  Mosquitoes (Diptera: Culicidae)  Morris, C.D., and G.R. Defoliart. 1969 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent      Govella, N.J., et al. 2011 (Parasites & Vectors)  Incandescent      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Carpenter, S., et al. 2008 (J. App. Ecol.)  UV      Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Rutledge, L.C., et al. 1975 (J. Med. Entomol.)  Incandescent      Alexander, B., et al. 1992. (Mem. Inst. Oswaldo Cruz)  Incandescent      Davies, C.R., et al. 1995 (Med. Vet. Entomol.)  Incandescent      Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent      Alten, B., et al. 2015 (Bull. Entomol. Res.)  Not specified  Infection prevalence  Biting midges (Diptera: Ceratopogonidae)  Mayo, C.E., et al. 2012 (Vet. Parasitol.)  UV      McDermott, E.G. et al. 2015 (Parasites & Vectors)  UV    Kissing bugs (Hemiptera: Reduviidae)  Walter, A.I., et al. 2005 (Cademos de Saude Publica)  Not specified      Barghini, A., and B.A.S. de Medeiros. 2010 (Environ. Health Prosp.)  Not specified      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  Incandescent  Studies are grouped by concern and vector group, and the light source used for each study is noted. View Large Historical Basis of Light Trapping One can imagine some of the earliest entomologists taking advantage of the attraction of nocturnal insects to primitive light sources like firelight or kerosene lamps, such as the attraction of a moth to a flame (Farb 1962). Indeed, even now flame can be used as an insect attractant, as a flickering candle is for fleas (Roucher et al. 2012), although interpreting the role of light from a flame is admittedly complicated by alternative cues such as heat or variable light intensities. This attraction to light, especially ultraviolet light, is likely due to nocturnal insects’ natural behavior to orient their movement based on the polarization patterns of moon and starlight (Warrant and Dacke 2011). The presence of artificial lights may disorient nocturnal insects, and therefore can be used to draw them into collection traps. Based on a Web of Science search using the key terms ‘light trap’, ‘insect’, and ‘trapping’, the first description of an electric light-baited insect trap we found was published in 1889 in The American Naturalist by Jerome McNeill. It was designed so that the dedicated ‘insect-hunter’ could continue to collect when he was ‘forced to go home to steal a few hours for sleep’. McNeill’s trap consisted of a metal funnel soldered to a tin pail, inside of which was a layer of plaster of Paris and potassium cyanide crystals, to kill any captured insects. An electric light was positioned above the mouth of the cone. Insects attracted to the light would fall down the cone, coated with varnish to prevent escape, and into the pail, where they would become ‘engaged in their desperate death struggles’. McNeill stated that his trap was especially good at collecting Hemipterans, Neuropterans, Dipterans, and Hymenopterans. It seems it was less suitable for collecting Lepidopterans, because beetles, less affected by the cyanide fumes, were presumably eating any moths that found their way into the trap (McNeill 1889). Since 1889, numerous versions of light traps designed to collect especially hemtaophagous insects of medical importance have been designed and patented. One of the most recognizable, the New Jersey light trap, was designed in 1932 (Headlee 1932), but its weight and dependence on heavy batteries or direct connection to an AC outlet for power made it impractical for large-scale field surveillance. The New Jersey light trap was nevertheless used, mainly for mosquitoes, for several decades. New Jersey traps are still sometimes used today, particularly to compare contemporary and historical collection data. The CDC miniature light trap, a much lighter and more convenient alternative for vector sampling and surveillance, has become a widely used alternative. The CDC light trap was designed by Drs. Dan Sudia and Roy Chamberlain at the Centers for Disease Control in the early 1960s (Sudia and Chamberlain 1962), and was inspired by an injury resulting from trying to carry the batteries required for the New Jersey trap (CDC 2015). CDC light traps rely on a small light (incandescent, UV, or LED) above a fan that creates a downdraft, sucking attracted insects into a hanging collection container below. Today, CDC traps, and variations on them, remain some of the most widely used traps for collecting crepuscular and nocturnal insects. The original light traps utilized incandescent bulbs, which give off much of their energy as heat and red or infrared light. The majority of nocturnal insects perceive wavelengths in the green-to-ultraviolet (UV) range, and cannot see red or infrared light (Briscoe and Chittka 2001), limiting the effectiveness of incandescent bulbs as attractants. To address this problem, entomologists began outfitting suction traps in the 1950s and 1960s with UV bulbs, which dramatically increased the number of insects collected (Pfrimmer 1955, Tashiro and Tuttle 1959, Belton and Pucat 1967, Rowley and Jorgensen 1967). More recently, there has been interest in the use of light-emitting diodes (LEDs) for trapping. LEDs have the added benefits of being available in a wide range of colors, and emitting very specific wavelengths, which certain genera, species, or physiological states may respond to differently (Hoel et al. 2007, Cohnstaedt et al. 2008, Snyder et al. 2016). By the early 1950s, entomologists had also begun to supplement light traps with carbon dioxide (CO2), which increased the number of hematophagous insects collected compared to light alone (Brown 1951, Reeves 1951, Newhouse et al. 1966, Snoddy and Hays 1966). Since then, other attractive odor cues (usually, but not always host-related) have been identified and used (Gibson and Torr 1999, Takken and Knols 2010), including 1-octen-3-ol (Hall et al. 1984, Takken and Kline 1989, Kline et al. 1994), the BG lure (Biogents AG, Regensburg, Germany), and lactic acid (Dekker et al. 2002), among others. Advantages of Light Traps There are advantages for the widespread and common use of light traps for vector surveillance, including 1) small size and weight, 2) reasonably low cost, 3) ability to run remotely (e.g., on small, portable batteries and without 120–220 V household current) for fairly long periods or to be automatically controlled by light-sensitive switches and fan breeze-operated gates, and 4) ability to collect a large variety of species, including both sexes. As an example, the model 512 CDC trap (J.D. Hock Co., Gainesville, FL) is only 14 cm tall and 8 cm in diameter. Apart from batteries or collecting container, it weighs a mere 325 g. The size and weight translate into the ability of a researcher to carry several such traps a number of kilometers into very remote field locations (e.g., Mullens and Dada 1992). Such traps are small enough to be inconspicuous, and they can often be placed in areas of fairly high human activity without great risk of vandalism. Their small size also allows for placement into confined places a larger trap won’t fit (e.g., into small rock recesses or upper tree canopies). Advances in light bulb technology, specifically the small but bright LEDs and automated photo-switches, also reduce the electrical requirements, and potentially extend trap run life into periods of several days. The ability of such traps to run overnight or longer without human monitoring is of course a very powerful and practical advantage. Another significant advantage of light traps is that they do in fact collect a lot of insects, both in terms of numbers and diversity. Together with the above advantages, this can make light traps the tool of choice for initial general surveys. Perhaps no other single vector surveillance tool gathers a large variety of species so easily. While some biases are known with regard to physiological condition (see below), the light traps may collect at least some insects that may not be receptive to host cues, such as gravid or blood-fed female flies. Simultaneously, since the main targets for vector sampling are nematocerous Diptera, males do not feed on blood and may not respond to host chemical cues like CO2. Light traps may be the only easy way researchers can collect males for studies of that sex or for purposes of identification of vector species where the females are difficult to recognize morphologically. Despite these advantages, certain limitations of light traps merit caution when interpreting collection data. Limitations of Light Trap Efficiency Decreased Attraction With Ambient Light Since the 1930s, the reduced ability of light-baited suction traps to collect hematophagous Diptera during nights on or near the full moon has been documented repeatedly (Bradley and McNeel 1935, Horsfall 1943, Pratt 1948, Provost 1959, Barr et al. 1963, Bidlingmayer 1967, Miller et al. 1970, Stryker and Young 1970, Bishop et al. 2000, Meiswinkel and Elbers 2016). This may be true even though some nocturnal biting flies fly more abundantly under full moon conditions, as was shown in flight interception studies of the bluetongue virus (BTV) vector Culicoides sonorensis Wirth and Jones (Diptera: Ceratopogonidae) (Nelson and Bellamy 1971, Akey and Barnard 1983, Linhares and Anderson 1990), as well as for several mosquito species (Bidlingmayer 1964). Perkin et al. (2014) recently showed that substantial artificial lighting (streetlights) can increase general activity of insects, including aquatic Diptera in the area, relative to darker zones. This increased activity does not appear to necessarily translate into larger trap catches. It is thought that ambient light reduces the contrast around the light trap, and so reduces its attractiveness to insects, or that artificial light sources compete with light traps. Even moderate amounts of ambient light may be enough to reduce collections in specific light traps, as those light traps only efficiently collect insects when ambient light intensity measures 0 lux (Meiswinkel and Elbers 2016). Because smaller catches on brighter nights could be misinterpreted as changes in the overall vector abundance in an area, it is important to note that simultaneously deployed, alternative trapping methods, including animal-baited collections (Pratt 1948), truck-traps (Provost 1959), and un-baited suction traps (Bidlingmayer 1967), do not show the same variation. While the reduced ability of light traps to collect insects a few nights a month near the full moon might be a minor problem, global light pollution exacerbates these concerns. It is estimated that light pollution increases by 6% every year, and extends even into historically rural or isolated areas (Gaston et al. 2012). Increased levels of nightly ambient light decrease nocturnal insect collections. As social progress has brought power and a better standard of living to many developing areas of the world, the amount of ambient light in areas that were previously rural and dark has increased dramatically. The ability to detect vector activity using light-baited suction traps in these areas will likely decrease over time, and research on the effect of artificial lighting on vector surveillance in urban areas is needed. Limited Range of Attraction Perhaps related to their reduced effectiveness with ambient light, light traps may vary drastically in their range of attraction. Studies with Culicoides biting midges differ in their conclusions about the range of light traps. Using CDC light traps in Denmark, Kirkeby et al. (2013) estimated their range of attraction to be ~15 m. Rigot and Gilbert (2012) found the range for Onderstepoort traps, which have a more powerful light and fan than CDC traps, to be as great as 30 m. However, other field studies have demonstrated far smaller ranges for Culicoides, even using the same traps. Venter et al. (2012) for example, demonstrated that the range of attraction of Culicoides to an Onderstepoort light trap in South Africa was only 2–4 m. Studies on Phlebotomine sand flies have also demonstrated that light traps have a short attraction range of 2–6 m (Killick-Kendrick et al. 1985, Valenta et al. 1995, Alexander 2000). Odeyoyinbo (1969) found that the range of attraction of Anopheles spp. in Africa to CDC incandescent light traps was ~5 m, while Costantini et al. (1998) found their range of attraction to be even less. These differences in the observed range of attraction are likely due to a number of potential confounding variables, including the type of trap used (and by extension, type of light used), species, ambient light, and the heterogeneity of the surrounding landscape. Because of this variation, comparisons of light trapping data across studies should be made with caution. Wavelength Differences The technological progression from incandescent light bulbs to UV to LEDs in insect trapping was driven by their increasing ability to collect both more species, as well as higher overall numbers of insects (Belton and Pucat 1967, Rowley and Jorgensen 1967, Cohnstaedt et al. 2008, Venter et al. 2009). Burkett et al. (1998) found that several Florida woodland mosquito species showed distinct color preferences, especially towards blue or green, when encountering CDC traps baited with both LEDs and CO2, though in many cases, incandescent light was equally attractive. In South Korea, LED baited traps producing a 365 nm peak were compared to traps using a 4 W UV bulb producing a less intense, but more diffuse mix of short wavelengths (Kim et al. 2017). The LED traps collected twice as many mosquitoes (188,125 vs 92,230) as well as more genera (12 vs 10), and more species (17 vs 14). LEDs have the advantage over traditional black lights of being available in very specific wavelengths, so they can emit not only in the traditional UV range, but in other colors as well. The availability of numerous wavelengths provides researchers with more tools to improve trapping methodology. The use of specific wavelengths could be an important study design consideration as some species are able to differentiate wavelength changes of only 10 nm (Snyder et al. 2016). Green light has been shown to be more attractive to at least some species of Culicoides than blue light is (Bishop et al. 2006, Hope et al. 2015, Gonzalez et al. 2016). In one study the important BTV vector, Culicoides obsoletus (Meigen) (Diptera: Ceratopogonidae) responded equally to most wavelengths, but less so to red light (Hope et al. 2015). In Egypt, phlebotomine sand flies were shown to be more attracted to red light. There, CDC suction traps baited with blue or green LEDs collected only about one-fifth the number of sand flies (Phlebotomus and Sergentomyia spp.) collected by traps baited with red LEDs (Hoel et al. 2007). Triatomine bugs are more attracted to blue lights, rather than green, yellow or red (Pacheco-Tucuch et al. 2012), but actually show a stronger attraction to incandescent light than UV (Minoli and Lazzari 2006). Certain species of insects may respond to different wavelengths of light (Bishop et al. 2006), and even feeding status may influence attraction (Snyder et al. 2016). In general, different wavelengths are often differentially attractive to different vector species. One must test and appreciate this in interpreting field light trap collections epidemiologically, if inter-species comparisons are intended. As data on these differing responses accumulates, LEDs will become increasingly valuable vector surveillance tools. Catch Size Although LED-baited traps sometimes enhance vector catch size over incandescent or UV-light baited traps, light trap collections in general are often smaller than those from traps baited with host cues, or from the host itself (Newhouse et al. 1966, Kline and Mann 1998, Gerry et al. 2009). Even species that can be collected well in light-only traps are often collected in significantly larger numbers in traps baited with semiochemicals, like CO2, lactic acid, or 1-octen-3-ol (Newhouse et al. 1966, Snoddy and Hays 1966, Stryker and Young 1970, Herbert et al. 1972, Alexander 2000, Andrade et al. 2008, Farajollahi et al. 2009, Kasap et al. 2009, Harrup et al. 2012). These chemical baits mimic the natural cues that host-seeking insects use for activation or orientation in the process of seeking a blood meal. As a result, these traps collect larger numbers of active insects of epidemiological interest, such as host-seeking females. Increasing collection sizes is a critical component of trap efficiency. Small collections are difficult, if not impossible, to analyze statistically. In cases where natural pathogen infection rates in vector populations are low (as is often true for arboviruses), small collections are far less likely to include any positive individuals than larger collections. Limitations of Light Trapping for Population Data Sex Biases Because male and female insects may respond differently to light, light trap collections are frequently biased towards one sex. Typically, these biases seem to be towards females (Belton and Pucat 1967, Venter et al. 2009, McDermott et al. 2016). Females of the sand fly, Lutzomyia whitmani (Coutinho and Antunes) (Diptera: Psychodidae) are much more attracted to light than males. In Brazil, suction traps with light were 76% L. whitmani females, whereas females comprised only 22% of catches of that species when the light was missing (Campbell-Lendrum et al. 1999). In some instances, males may make up a large proportion of collections, but even when considering the same species and same attractant, differences in the sex ratio between trapping locations may exist. One study found that C. sonorensis males made up 13.8% of UV trap collections on average at one dairy site in California, but over 45% of UV collections at a second dairy. In comparison, the proportion of males in simultaneously deployed CO2 baited traps was more similar between farms (4.8 vs 9.7%) (McDermott et al. 2016). It can be difficult to ascertain causes of location-specific differences in relative collections of males versus females. Males of a number of blood-feeding Diptera are more likely to swarm or seek mates close to larval developmental sites, however, and some species will orient to host animals or related chemical cues such as CO2 in order to locate females (Yuval 2006). The consequences of a sex-biased collection vary depending on the goals of the study. In the nematocerous Diptera, males are less epidemiologically important, and so their underrepresentation in light trap collections may not be a major concern. However, broader studies of biology or ecology of a given vector species may suffer from a sole reliance on light traps. In studies where the collection of male individuals is vital, such as for species identification or to understand mating patterns, male-targeted (e.g., seeking out swarming sites) or less biased collection methods (e.g., vehicle-mounted nets, or aerial sweeping) should be employed (Sanders et al. 2012). Parity Biases Differences in the proportions of parous and nulliparous females collected in light traps present more of a problem for epidemiological studies. Many of the pathogens of interest for human and animal health are not frequently or ever transovarially transmitted, and so only parous females (which have previously taken a blood meal from a host) are potentially infected. Because of this, and because of the costs associated with testing insect samples for pathogens, testing only parous females increases both economic efficiency, and the chances of detecting pathogens. Culicoides midges are unique in that they can be sorted visually by parity rather quickly and easily based on parous pigment deposited in the abdominal cuticle (Dyce 1969, Akey and Potter 1979). Other groups of hematophagous insects require more involved, subjective, or laborious techniques such as dissection of the ovaries, or inexact indicators such as wing fray, to determine parity or age (Hayes and Wall 1999, Hugo et al. 2008). Parity sorting thus can quickly become cost- and time- prohibitive, although it can provide invaluable population-level information on vector survival, and thus may be worth the effort. Even for large collections of Culicoides, the time required for visual parity sorting is a concern. Traps that preferentially collect parous females could significantly improve the efficiency of detecting pathogens. Nulliparous female C. sonorensis in the United States and Culicoides brevitarsis Kieffer (Diptera: Ceratopogonidae) in Australia seem to respond relatively poorly to light, and therefore may make up a smaller proportion of catches in UV-baited traps than in CO2-baited traps (Bellis and Reid 1996, McDermott et al. 2016). In a recent study, the proportion of parous females (compared to nullipars and males) collected did not differ between those two attractants (McDermott et al. 2016), so there may not be an advantage to using UV baited traps in terms of pathogen detection. Gravid C. sonorensis do seem to be fairly attracted to light (Anderson and Linhares 1989), so UV traps are useful in their collection. Alternatively, light traps have been criticized for their bias towards unfed/nulliparous female mosquitoes (Reisen and Pfuntner 1987, Githeko et al. 1994, Williams and Gingrich 2007). It is especially important to consider parity rates in collections used to estimate infection rates of mosquito-borne viruses as collections are almost always tested as unsorted pools of parous and nulliparous females. This can negatively affect infection rate estimates. For example, Williams and Gingrich (2007) compared West Nile virus (WNV) infection rates of mosquitoes collected using simultaneously deployed gravid traps and CO2-baited CDC light traps in Delaware. They found that gravid trap collections had estimated infection rates 32 times higher than the light traps, despite light traps collecting 57,000 more insects. Even though their collections are smaller, gravid traps have been recommended as a more efficient means of detecting pathogens in collected mosquitoes (Reisen and Pfuntner 1987, Williams and Gingrich 2007) because barring autogeny, all females have taken at least one blood meal, minimizing the need to sort by parity. Another important consequence of parity error is its effect on survivorship estimates, which are especially relevant for pathogens biologically transmitted by hematophagous Diptera. Probability of daily survival determines how well vectors survive the required extrinsic incubation period, and is highly influential in vectorial capacity calculations (Reisen 2009). In a simple but commonly used form, daily survival is derived by raising the proportion of parous females to the nth power, with n being one divided by the number of days required for a gonotrophic cycle (Davidson 1954). Using one 10-wk long California suction trap study on C. sonorensis (Anderson and Linhares 1989) as an example, a trap with CO2 (no light) had 32% parity, a trap with a UV light plus CO2 had 51% parity, and a trap with UV only had 98% parity (mostly gravid flies). The authors did not attempt to calculate survival, recognizing the inherent biases of the trapping, but, extrapolating from their data to make the point (assuming a 4 d gonotrophic cycle length), this would yield rough estimates of daily survival of 75%, 85%, and over 99%, with consequent impacts on calculated vectorial capacity. Certainly the last estimate would be completely unrealistic if used in vectorial capacity, and it would be based on biased parity information that did not accurately reflect either the vector population as a whole or the biting portion of that population. All of these estimates may also differ from a survival estimate calculated from direct animal aspiration collections, an arguably more realistic way to generate a population snapshot. Limitations of Light Trapping for Pathogen Detection Species Composition Perhaps the most critical overall limitation of light traps is their potential to inaccurately reflect the species composition of host-biting insect communities in an area. Many vector species respond poorly or unevenly to light, resulting in species biases in light trap collections. Studies that rely solely on light traps for collections run the risk of missing important species, and drawing false conclusions about transmission risk and putative vectors. The risk of misinterpreting light trap data is most severe in areas where many potential vector species exist, and are of unknown significance for transmission. This occurs regularly in neglected areas, or with emerging pathogens. This problem can even occur in such a well-studied region as Western Europe. Researchers and regulatory agencies there were unprepared for the large, persistent BTV outbreaks during 1999–2009 (Purse et al. 2015) primarily because many countries, without a compelling economic reason, just had not yet studied the main vector genus (Culicoides). Consequently, many locations were at the point of initial exploratory Culicoides community surveys as the outbreaks began, relative to their far better knowledge of mosquitoes or ticks. Studies on both mosquitoes and biting midges have addressed the problem of species diversity in light traps by comparing light-baited trap collections to those from host cue-baited traps, or to concurrent, direct collections from hosts. In the United Kingdom, while UV light-baited suction traps were efficient in collecting large numbers of C. obsoletus and Culicoides scoticus Downes and Kettle (Diptera: Ceratopogonidae), they only collected a small number of another potential BTV vector, Culicoides chiopterus Meigen (Diptera: Ceratopogonidae). Based on light trap data alone, it might have been concluded that C. chiopterus was not an abundant, and therefore important, vector in the United Kingdom. However, concurrent drop-trap collections from live sheep contained large numbers of C. chiopterus (Carpenter et al. 2008). Similarly, in Spain, direct aspiration collections from live sheep resulted in large numbers of the potential BTV vectors, C. obsoletus and Culicoides parroti Kieffer (Diptera: Ceratopogonidae), which were not well represented in collections from light traps deployed at the same time and place as the animal collections were made (Gerry et al. 2009). In France, light traps significantly underestimated the abundance of Culicoides brunnicans Edwards (Diptera: Ceratopogonidae) on sheep (Viennet et al. 2011). In Vietnam, a comparison of CDC traps baited with either CO2, light, or CO2 and light found that of all 23 mosquito species collected, the vast majority of individuals of a given species were collected by CO2/light combination (6.9–100%) or CO2-only (42.2–100%) traps. By comparison, light-only traps collected only 0–13.8% of the individuals of a given species (Herbert et al. 1972). In Florida, CDC traps baited with both CO2 and light collected a total of 17 mosquito species (Aedes, Anopheles, Culex, Culiseta, Psorophora, and Uranotaenia spp.), compared to only seven species in light-only traps. No Psorophora or Uranotaenia spp. were collected in traps without CO2 (Kline and Mann 1998). In Tanzania, CDC light traps were found to underestimate human biting rates of both Anopheles and Culex spp. (Govella et al. 2011), although other studies in Africa have shown light trap collections to be an acceptable measure of attraction to human hosts (Costantini et al. 1998), especially when deployed inside homes. In Korea, a comparison of UV and LED light baited traps for collecting Japanese encephalitis virus (JEV) vectors found that neither was particularly good at collecting Culex tritaeniorhynchus Giles (Diptera: Culicidae), the primary JEV vector in the area. Cx. tritaeniorhynchus females made up only 0.48 and 0.58% of UV and LED trap collections, respectively (Kim et al. 2017). For situations where the vector species are few and well known, such as Anopheles vectors of human Plasmodium in parts of Africa, relating light trap catches to biting and transmission risk becomes somewhat more tractable. Briet et al. (2015) conducted a meta-analysis of 13 studies that used the human landing catch (HLC) together with CDC light trap sampling. Those authors concluded that collections in CDC traps were at least approximately proportional to HLC, although studies were tremendously variable, and reliable generalizations about the relationship were impossible to derive. They argued that such data were still useful despite the imprecision, given the ease of use and safety of light traps, and that it was the very large changes in risk that were most epidemiologically relevant. Another important consideration for the use of light traps to collect mosquito vectors is the growing importance of day biting, highly anthropophilic Aedes spp., especially Ae. albopictus (Skuse; Diptera: Culicidae) and Ae. aegypti (L.; Diptera: Culicidae). In the last 30 yr, several Aedes-transmitted viruses, including dengue virus, chikungunya virus, and Zika virus, have emerged across the globe, causing massive public health problems (Roth et al. 2014). Aedes spp. competent for these viruses are widely established, including in temperate zones (Kraemer et al. 2015). Aedes spp., and Ae. albopictus and Ae. aegypti in particular, are not well collected in light traps (Morris and Defoliart 1969, Krockel et al. 2006, Farajollahi et al. 2009), and even CO2 baited traps are not particularly effective (Krockel et al. 2006). By comparison, human scent-baited suction traps without light, like the BG-Sentinel trap (Biogents AG) have proved to be much more effective collectors of Aedes vector spp. (Krockel et al. 2006, Maciel-de-Freitas et al. 2006, Farajollahi et al. 2009). The BG trap in particular was also shown to collect fewer nullipars than pars or blood-fed females (Ball and Ritchie 2010), making it a potentially more efficient means of sampling day-biting Aedes populations for viruses. As with the highly anthropophilic mosquito species, UV-light baited suction traps may be of little use for collecting anthro- or zoophilic phlebotomine sand flies (Rutledge et al. 1975, Andrade et al. 2008). Host odor cues, including CO2 (Kasap et al. 2009), 1-octen-3-ol, and the BG-lure (Andrade et al. 2008) can increase sand fly collections. For example, Muller et al. (2015) did have success catching large numbers of Phlebotomus papatasi (Scopoli) (Diptera: Psychodidae) using UV or incandescent light with CDC traps, especially if they also were baited with CO2 and other semiochemicals. Similarly, the widespread sole reliance on light traps to collect Culicoides midges has reinforced the common assumption that biting midges are entirely crepuscular and nocturnal, and not active during winter months. While not an unfounded generalization for the genus, this is not strictly true, as there are Culicoides species that are mainly diurnal, such as the suspected Australian BTV vector Culicoides actoni Smith (Diptera: Ceratopogonidae) (Bellis et al. 2004). Light traps on their own could quite possibly miss such species entirely. More importantly, during fall and winter, Culicoides often display a shift in their activity patterns, and a normally crepuscular or nocturnal species in summer can occasionally (or perhaps only) be collected several hours before sunset when temperatures are favorable for flight during cooler months (Lillie et al. 1987, Sanders et al. 2012). A clear demonstration of this occurred in year-round flight interception studies of C. sonorensis activity in Colorado (Barnard and Jones 1980). There, essentially all the adult activity in spring (April) and fall (November) was before sunset, when light traps were expected to be ineffective. In the summer in temperate climates, the BTV vectors C. obsoletus complex and C. chiopterus can be collected using sweep nets throughout the 24-h period, but light traps only collect insects after the sun has completely set and light intensity measures 0 lux (Meiswinkel and Elbers 2016). In fact, Meiswinkel and Elbers (2016) found that an Onderstepoort light trap operating from 2100 to 2200 h (dusk) collected only one C. obsoletus complex individual, while simultaneous aspiration directly off of sentinel cattle during just that 1-h period resulted in collections of 796 C. obsoletus complex and 709 C. chiopterus. C. chiopterus peak activity was found to actually occur more than 2 h before sunset. In Spain, Gerry et al. (2009) caught many female Culicoides before sunset, and numbers of mammal-feeding insects (C. obsoletus and C. parotti) aspirated from sentinel sheep over short (5 min) periods far exceeded the cumulative number taken in UV-baited CDC light traps running continuously nearby. The Gerry et al. (2009) study emphasizes again that one must know something about host feeding patterns when interpreting light trap catches. The most common Culicoides species in the light traps, by far, was Culicoides circumscriptus Kieffer (Diptera: Ceratopogonidae). Without prior knowledge of the species, particularly in the beginning stages of vector investigations, one might assume that it was a major potential vector of a virus like bluetongue. However, this species was entirely lacking from the sheep collections because C. circumscriptus is a bird feeder (Braverman and Linley 1994). With groups like the sand flies, where species tend to be restricted to certain heights within a vertical space, species biases in light trap collections can also occur. Due to the nature of the attractant, these traps have a relatively limited collection range in vertical space. Phlebotomine sand fly species that are active either lower to the ground or higher up are unlikely to be sampled consistently using light-baited suction traps (Alten et al. 2015). The use of host-cues, or hosts, in place of or in combination with light can increase sand fly numbers in collections. Campbell-Lendrum et al. (1999) placed CDC suction traps for Lutzomyia spp. directly above potential bait animals (humans, chickens, dogs), and found that traps with light collected more L. whitmani and Lutzomyia intermedia (Lutz and Neiva) (Diptera: Psychodidae) than did similarly placed traps without light. Compared to L. whitmani, L. intermedia collections were relatively more enhanced by the addition of light, suggesting that these species differ in their attraction to light. Light traps collected larger total numbers of the potential Peruvian Leishmania vector, Lutzomyia peruensis Velez (Diptera: Psychodidae) than HLC collections, but HLC collections contained a higher proportion of this species (45/955 total sand flies in light traps vs 80/3029 in HLC) (Davies et al. 1995). Once again, prior knowledge of the biology of a particular species is necessary for study design in this case. Infection Prevalence Rarely, cases might arise where the use of light traps has a direct negative impact on pathogen detection or transmission due to vector behavior. One compelling field example is the evidence that BTV infection apparently causes C. sonorensis midges to become averse to UV light. Work by Mayo et al. (2012) and McDermott et al. (2015) in California showed that BTV infection rates in pools of parous C. sonorensis collected using UV-light (315–400 nm, peak at 350 nm) baited CDC suction traps, or traps using both CO2 and UV-light, were significantly lower than in pools collected using CO2-only baited traps. UV trap collections underestimated the infection rates seen in CO2 trap collections by as much as 8.5 times (McDermott et al. 2015). UV wavelengths above 320 nm have essentially no microbiocidal properties and are well above the far higher energy, shorter wavelengths (250–260 nm) used for disinfection (Wolfe 1990, Bintsis et al. 2000), making it highly unlikely that the observed lower infection rates were due to UV degradation of BTV. This is suggestive then of behavioral manipulation of C. sonorensis by BTV, causing infected midges to avoid, or possibly be repelled by, UV light. UV traps were particularly poor at collecting infected midges when the overall number of midges in an area was low. Importantly then, there is the risk of missing active, BTV-infected vectors when vector abundance is low, such as periods of perceived overwintering or following the introduction of infected vectors into virus-free areas, inhibiting the ability to enact early eradication measures. Finally, a second rare, but possibly important, consideration when using light traps is the risk of drawing infected vectors into close proximity with hosts. Although the range of attraction to light for many species is limited (as discussed previously), some vector groups are highly phototactic, and artificial lights can influence natural dispersal behaviors, with unintended consequences for pathogen transmission. In historically rural, Chagas disease-endemic regions, urbanization, improved home construction, and aggressive vector control programs have helped to eliminate the traditional domestic Triatomine kissing bug vector species, but the increase in outdoor electric lighting was accompanied by a subsequent increase in orally transmitted Chagas and shift to sylvatic and peridomestic vector species (Barghini and de Medeiros 2010). These typically outdoor associated species are drawn to houses by bright outdoor lights (Pacheco-Tucuch et al. 2012), where they defecate the infectious metacyclic trypomastigote stage of the parasite onto fruits or other food grown or held nearby. The contaminated food is then consumed by people or animals, resulting in disease (Walter et al. 2005, Barghini and de Medeiros 2010). Although specifically discussed here in terms of highly phototactic Triatomine vectors, all attractive traps could potentially pose this risk, and placement of traps near hosts deserves careful consideration by researchers. Conclusions Light traps are one of, if not the most, widely used vector surveillance methods available. They are both practical and portable, and lend themselves to long-term surveillance studies. In some cases, they may be the only readily available option. However, light traps have potentially serious limitations, especially if used as the sole sampling tool. Researchers must understand the pros and cons associated with light traps when making decisions about their study designs. Ease of use should not outweigh the risks of not identifying vector species or misunderstanding pathogen activity in an area. This holds especially true when light trap data are being used to make important public or livestock health decisions, like when to make pesticide applications or transport animals. The efficacy of light traps for vector collection is highly dependent on assumptions about vector behavior. It is critical to take into account the published data that challenge these assumptions when choosing traps or attractants for medical and veterinary entomology field studies. Light trap collections can be biased towards certain species, sexes or physiological states. Competent vector species may not be strongly phototactic, and their low numbers in light trap collections could lead to their dismissal as important for pathogen transmission. Likewise, when light traps preferentially collect nulliparous females or males, the ability to detect pathogens in pooled insect samples drops. Light traps do not efficiently collect day active or highly anthro- or zoophilic species, so light trap collections may lead to a misunderstanding of vector activity patterns. In some cases, pathogen infection may cause vectors to alter their phototactic behavior, resulting in errors in transmission risk estimates, as has been suggested for BTV-infected midges. Because these ‘subtle effects’ of pathogen manipulation (those that have no obvious impact on transmission) are largely unstudied, other vector-pathogen interactions in other disease systems may exist that limit the effectiveness of light traps. For highly phototactic species, the use of light traps may put people and animals at risk by drawing large numbers of vectors into areas near hosts, as has been hypothesized with Chagas disease. In rural areas, researchers may want to consider whether to place light traps away from livestock enclosures and habitations. In areas where light traps are the only logistically realistic option for vector surveillance, the use of LEDs may improve efficiency by allowing for the use of targeted wavelengths. LED baited traps have been shown to collect higher numbers of midges, mosquitoes, and sand flies compared to UV or incandescent lights, and may also collect a higher diversity of insects. LEDs in specific wavelengths can be selected for surveillance programs or field studies in order to effectively target certain vector species of interest. That being said, ambient light, which is increasingly a problem in our developing world, significantly decreases the effectiveness of all light traps. Even small amounts of ambient light have been shown to reduce the number of insects collected in a given trap. The effects of light pollution across the globe have been discussed in any number of contexts, but perhaps it is time medical and veterinary entomologists began to consider more seriously the effect it may have on vector surveillance. There is no such thing as a perfect insect sampling technique, especially when the stakes are as high as they can be with vector borne diseases. All trapping methods will have advantages and disadvantages. That does not mean, though, that critical flaws in a method should be ignored because of convenience. Today, it is clear that medical and veterinary entomology could benefit from innovative alternatives or supplements to the light baited suction trap. The best way to improve the interpretation of light trap collections, and indeed all trap collections, is to continue to collect data on how they compare to other trapping methods and other light trap studies for as many species and in as many situations as possible. When conducting surveillance studies, researchers should consider including multiple attractants (e.g., UV vs CO2 or UV vs LED) for comparison to traditional light traps. When running long-term vector surveillance programs using light traps (such as those conducted by vector control districts), periodic collections using host-cue baited traps, or other non-light methods, to check light trap assumptions should be considered. Light traps will continue to have a place in medical and veterinary entomology, and in some cases may truly be the most appropriate choice. However, more broad and critical discussion about the validity of the epidemiological conclusions drawn from these collections is needed in order for entomologists and epidemiologists to avoid the dark side of light traps. Acknowledgments We would like to thank William Reisen and Christopher Barker for their input and discussion on mosquito surveillance trends and helpful comments on this article, as well as Alec Gerry and Lindsey Garver, who also provided generous feedback to improve the article. The material to be published reflects the views of the authors and should not be construed to represent those of the United States Department of the Army or the United States Department of Defense. References Cited Akey, D., and Potter H.. 1979. Pigmentation associated with oogenesis in the biting fly Culicoides variipennis (Diptera: Ceratopogonidae): determination of parity. J. Med. Entomol . 16: 67– 70. Google Scholar CrossRef Search ADS PubMed  Akey, D. H., and Barnard D. R.. 1983. Parity in airborne populations of the biting gnat Culicoides variipennis (Diptera: Ceratopogonidae) in northeastern Colorado. Environ. Entomol . 12: 91– 95. Google Scholar CrossRef Search ADS   Alexander, B. 2000. 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Published by Oxford University Press on behalf of Entomological Society of America. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Journal of Medical Entomology Oxford University Press

The Dark Side of Light Traps

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Entomological Society of America
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© The Author(s) 2017. Published by Oxford University Press on behalf of Entomological Society of America. All rights reserved. For permissions, please e-mail: journals.permissions@oup.com.
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0022-2585
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1938-2928
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10.1093/jme/tjx207
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Abstract

Abstract Light-baited suction traps are one of the most widely used tools for vector surveillance. Their popularity stems from ease of use even in remote locations, range and abundance of species caught, and low cost. The availability of smaller, portable models, like the CDC miniature light trap, have further increased their ubiquity in entomological field studies. However, when researchers have looked, light trap collections are usually biased in ways that may affect data interpretation for epidemiological studies. If used alone, light traps may fail to collect important or infected vectors, and light traps are inefficient or ineffective when competing ambient light is present. In this article, we discuss these biases and limitations in terms of their effect on collection efficiency, population data, and pathogen detection. While light trap data certainly have a purpose, an over-reliance on light trapping risks drawing false conclusions about vector populations and vector-borne disease epidemiology. These concerns are especially troubling when light trap data are used to inform policy decisions meant to protect human and animal health. Particularly when a species’ response to light is unknown or poorly characterized, light traps should be used in conjunction with supplemental sampling methods. Researchers conducting vector surveillance field studies should carefully consider their study design and objectives when deciding on a trapping method or methods, and specifically endeavor to understand the limitations of their data. Only then can researchers take advantage of the best attributes of light traps while avoiding their dark side. light trap, surveillance, vector, trapping Light traps have been used for almost 130 yr to collect a variety of nocturnal and crepuscular insects. In medical and veterinary entomology, they are most often used to collect mosquitoes (Diptera: Culicidae), biting midges (Diptera: Ceratopogonidae), and sand flies (Diptera: Psychodidae: Phlebotominae), but have also occasionally been used to collect black flies (Diptera: Simuliidae), house flies (Musca domestica L.), horn flies (Haemotobia irritans (L.)) (Diptera: Muscidae), fleas (Siphonaptera), and kissing bugs (Hemiptera: Reduviidae: Triatominae) (Bram 1978). Data stemming from light trap collections, including species composition, population age structure, abundance, and infection rate, are sometimes used to inform epidemiological models for vector-borne diseases, and light traps can be useful in that respect, as we discuss below. However, there also is a growing body of literature describing the ways in which light traps can fail to accurately represent vector populations or pathogen activity in an area. This article discusses some of those issues, in the hopes that researchers may better appreciate caveats regarding the use of light trap data in epidemiological studies, particularly if light traps are used without any supplemental sampling. Light trap limitations range in their severity from relatively minor influences on trap effectiveness (e.g., limited range of attraction), to those that may affect the interpretation of vector populations in an area (e.g., sex biases), all the way to the most serious limitations: failing to collect vector species, or detect pathogens of interest (Table 1). We suggest that the use of light trap data should be tempered by these considerations, and argue that in many cases, alternatives or supplements to light trapping should be sought in order to improve studies on vector biology and vector-borne disease epidemiology. Table 1. Summary of literature discussing the potential biases and drawbacks of light trapping for insect vectors Concern  Vector group  Study  Light source(s) used  I. Limitations of efficiency      Decreased attraction with ambient light  Mosquitoes (Diptera: Culicidae)  Bradley, G.H., and T.E. McNeel. 1935 (J. Econ. Entomol.)  Incandescent      Horsfall, W.R. 1943 (Ann. Entomol. Soc. Am.)  Incandescent      Pratt, H.D. 1948 (J. Natl. Malar. Soc.)  Incandescent      Provost, M.W. 1959 (Ann. Entomol. Soc. Am.)  Incandescent      Barr, A.R., et al. 1963 (J. Econ. Entomol.)  Incandescent      Bidlingmayer, W.L. 1967 (J. Med. Entomol.)  Incandescent      Miller, T.A., et al. 1970 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Bishop, A.L., et al. 2000 (Austr. J. Entomol.)  Incandescent      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV  Limited range of attraction  Mosquitoes (Diptera: Culicidae)  Odetoybino, J.A. 1969 (Bull. Wld. Hlth. Org.)  Incandescent      Constantini, C., et al. 1998 (Bull. Entomol. Res.)  Incandescent, UV    Biting midges (Diptera: Ceratopogonidae)  Venter, G.J., et al. 2012 (Vet. Parasitol.)  UV    Sand flies (Diptera: Psychodidae)  Killick-Kendrick, R., et al. 1985. (Annales de Parasitologie et Humaine Comparee)  Incandescent      Valenta, D.T., et al. 1995. (Proc. II Int. Symp. Phlebot. Sand Flies)  Incandescent      Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified  Attraction to different wavelengths  Mosquitoes (Diptera: Culicidae)  Burkett, D. A., et al. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Bishop, A.L., et al. 2006 (Austr. J. Entomol.)  LED, UV, Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Hope, A., et al. 2015 (Parasites & Vectors)  LED, UV      Gonzalez, M., et al. 2016 (Vet. Parasitol.)  LED, UV, Incandescent    Sand flies (Diptera: Psychodidae)  Hoel, D.F., et al. 2007 (J. Vect. Ecol.)  Incandescent, LED    Kissing bugs (Hemiptera: Reduviidae)  Minoli, S.A., and Lazzari, C.R. 2006 (Acta Tropica)  Incandescent, UV      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  LED, UV, Incandescent    General  Cohnstaedt, L.W., et al. 2008 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED  Small catch size  Mosquitoes (Diptera: Culicidae)  Newhouse, V.F., et al. 1966 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Harrup, L.E., et al. 2012 (J. Med. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified      Andrade, A.J., et al. 2008 (Mem. Inst. Oswaldo Cruz)  Incandescent      Kasap, O.E., et al. 2009 (ACTA Vet. BRNO)  Incandescent    Black flies (Diptera: Simuliidae)  Snoddy, E.L., and K.L. Hays. 1966 (J. Econ. Entomol.)  Incandescent  II. Limitations for population data        Sex biases  Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent  Parity biases  Mosquitoes (Diptera: Culicidae)  Reisen, W.K., and A.R. Pfuntner. 1987 (J. Amer. Mosq. Contr. Assoc.)  Incandescent      Githeko, A.K., et al. 1994 (Bull. Entomol. Res.)  Incandescent      Williams, G.M., and J.B. Gingrich. 2007 (J. Vect. Ecol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Anderson, J.R., and A.X. Linhares. 1989 (J. Amer. Mosq. Cont. Assoc.)  UV      Bellis, G.A., and D.J. Reid. 1996 (Austr. J. Entomol.)  Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV  III. Limitations for pathogen detection      Species composition  Mosquitoes (Diptera: Culicidae)  Morris, C.D., and G.R. Defoliart. 1969 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent      Govella, N.J., et al. 2011 (Parasites & Vectors)  Incandescent      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Carpenter, S., et al. 2008 (J. App. Ecol.)  UV      Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Rutledge, L.C., et al. 1975 (J. Med. Entomol.)  Incandescent      Alexander, B., et al. 1992. (Mem. Inst. Oswaldo Cruz)  Incandescent      Davies, C.R., et al. 1995 (Med. Vet. Entomol.)  Incandescent      Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent      Alten, B., et al. 2015 (Bull. Entomol. Res.)  Not specified  Infection prevalence  Biting midges (Diptera: Ceratopogonidae)  Mayo, C.E., et al. 2012 (Vet. Parasitol.)  UV      McDermott, E.G. et al. 2015 (Parasites & Vectors)  UV    Kissing bugs (Hemiptera: Reduviidae)  Walter, A.I., et al. 2005 (Cademos de Saude Publica)  Not specified      Barghini, A., and B.A.S. de Medeiros. 2010 (Environ. Health Prosp.)  Not specified      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  Incandescent  Concern  Vector group  Study  Light source(s) used  I. Limitations of efficiency      Decreased attraction with ambient light  Mosquitoes (Diptera: Culicidae)  Bradley, G.H., and T.E. McNeel. 1935 (J. Econ. Entomol.)  Incandescent      Horsfall, W.R. 1943 (Ann. Entomol. Soc. Am.)  Incandescent      Pratt, H.D. 1948 (J. Natl. Malar. Soc.)  Incandescent      Provost, M.W. 1959 (Ann. Entomol. Soc. Am.)  Incandescent      Barr, A.R., et al. 1963 (J. Econ. Entomol.)  Incandescent      Bidlingmayer, W.L. 1967 (J. Med. Entomol.)  Incandescent      Miller, T.A., et al. 1970 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Bishop, A.L., et al. 2000 (Austr. J. Entomol.)  Incandescent      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV  Limited range of attraction  Mosquitoes (Diptera: Culicidae)  Odetoybino, J.A. 1969 (Bull. Wld. Hlth. Org.)  Incandescent      Constantini, C., et al. 1998 (Bull. Entomol. Res.)  Incandescent, UV    Biting midges (Diptera: Ceratopogonidae)  Venter, G.J., et al. 2012 (Vet. Parasitol.)  UV    Sand flies (Diptera: Psychodidae)  Killick-Kendrick, R., et al. 1985. (Annales de Parasitologie et Humaine Comparee)  Incandescent      Valenta, D.T., et al. 1995. (Proc. II Int. Symp. Phlebot. Sand Flies)  Incandescent      Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified  Attraction to different wavelengths  Mosquitoes (Diptera: Culicidae)  Burkett, D. A., et al. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Bishop, A.L., et al. 2006 (Austr. J. Entomol.)  LED, UV, Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Hope, A., et al. 2015 (Parasites & Vectors)  LED, UV      Gonzalez, M., et al. 2016 (Vet. Parasitol.)  LED, UV, Incandescent    Sand flies (Diptera: Psychodidae)  Hoel, D.F., et al. 2007 (J. Vect. Ecol.)  Incandescent, LED    Kissing bugs (Hemiptera: Reduviidae)  Minoli, S.A., and Lazzari, C.R. 2006 (Acta Tropica)  Incandescent, UV      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  LED, UV, Incandescent    General  Cohnstaedt, L.W., et al. 2008 (J. Am. Mosq. Contr. Assoc.)  Incandescent, LED  Small catch size  Mosquitoes (Diptera: Culicidae)  Newhouse, V.F., et al. 1966 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Harrup, L.E., et al. 2012 (J. Med. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Alexander, B. 2000 (Med. Vet. Entomol.)  Not specified      Andrade, A.J., et al. 2008 (Mem. Inst. Oswaldo Cruz)  Incandescent      Kasap, O.E., et al. 2009 (ACTA Vet. BRNO)  Incandescent    Black flies (Diptera: Simuliidae)  Snoddy, E.L., and K.L. Hays. 1966 (J. Econ. Entomol.)  Incandescent  II. Limitations for population data        Sex biases  Biting midges (Diptera: Ceratopogonidae)  Belton, P., and A. Pucat. 1967 (Can. Entomol.)  Incandescent, UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent  Parity biases  Mosquitoes (Diptera: Culicidae)  Reisen, W.K., and A.R. Pfuntner. 1987 (J. Amer. Mosq. Contr. Assoc.)  Incandescent      Githeko, A.K., et al. 1994 (Bull. Entomol. Res.)  Incandescent      Williams, G.M., and J.B. Gingrich. 2007 (J. Vect. Ecol.)  Incandescent    Biting midges (Diptera: Ceratopogonidae)  Anderson, J.R., and A.X. Linhares. 1989 (J. Amer. Mosq. Cont. Assoc.)  UV      Bellis, G.A., and D.J. Reid. 1996 (Austr. J. Entomol.)  Incandescent      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      McDermott, E.G., et al. 2016 (Med. Vet. Entomol.)  UV  III. Limitations for pathogen detection      Species composition  Mosquitoes (Diptera: Culicidae)  Morris, C.D., and G.R. Defoliart. 1969 (Mosq. News)  Incandescent      Stryker, R.G., and W.W. Young. 1970 (Mosq. News)  Incandescent      Herbert, E.W., et al. 1972 (Mosq. News)  Incandescent      Kline, D.L., and M.O. Mann. 1998 (J. Am. Mosq. Contr. Assoc.)  Incandescent      Farajollahi, A., et al. 2009 (J. Med. Entomol.)  Incandescent      Govella, N.J., et al. 2011 (Parasites & Vectors)  Incandescent      Kim, H.-C., et al. 2017 (J. Med. Entomol.)  LED, UV    Biting midges (Diptera: Ceratopogonidae)  Rowley, W.A., and N.M. Jorgensen, 1967 (J. Econ. Entomol.)  Incandescent, UV      Carpenter, S., et al. 2008 (J. App. Ecol.)  UV      Gerry, A.C., et al. 2009 (J. Med. Entomol.)  UV      Venter, G.J., et al. 2009 (Vet. Parasitol.)  Incandescent, UV      Meiswinkel, R., and A.R.W. Elbers. 2016 (Med. Vet. Entomol.)  UV    Sand flies (Diptera: Psychodidae)  Rutledge, L.C., et al. 1975 (J. Med. Entomol.)  Incandescent      Alexander, B., et al. 1992. (Mem. Inst. Oswaldo Cruz)  Incandescent      Davies, C.R., et al. 1995 (Med. Vet. Entomol.)  Incandescent      Campbell-Lendrum, D., et al. 1999 (Mem. Inst. Oswaldo Cruz)  Incandescent      Alten, B., et al. 2015 (Bull. Entomol. Res.)  Not specified  Infection prevalence  Biting midges (Diptera: Ceratopogonidae)  Mayo, C.E., et al. 2012 (Vet. Parasitol.)  UV      McDermott, E.G. et al. 2015 (Parasites & Vectors)  UV    Kissing bugs (Hemiptera: Reduviidae)  Walter, A.I., et al. 2005 (Cademos de Saude Publica)  Not specified      Barghini, A., and B.A.S. de Medeiros. 2010 (Environ. Health Prosp.)  Not specified      Pacheco-Tucuch, F.S., et al. 2012 (PLoS One)  Incandescent  Studies are grouped by concern and vector group, and the light source used for each study is noted. View Large Historical Basis of Light Trapping One can imagine some of the earliest entomologists taking advantage of the attraction of nocturnal insects to primitive light sources like firelight or kerosene lamps, such as the attraction of a moth to a flame (Farb 1962). Indeed, even now flame can be used as an insect attractant, as a flickering candle is for fleas (Roucher et al. 2012), although interpreting the role of light from a flame is admittedly complicated by alternative cues such as heat or variable light intensities. This attraction to light, especially ultraviolet light, is likely due to nocturnal insects’ natural behavior to orient their movement based on the polarization patterns of moon and starlight (Warrant and Dacke 2011). The presence of artificial lights may disorient nocturnal insects, and therefore can be used to draw them into collection traps. Based on a Web of Science search using the key terms ‘light trap’, ‘insect’, and ‘trapping’, the first description of an electric light-baited insect trap we found was published in 1889 in The American Naturalist by Jerome McNeill. It was designed so that the dedicated ‘insect-hunter’ could continue to collect when he was ‘forced to go home to steal a few hours for sleep’. McNeill’s trap consisted of a metal funnel soldered to a tin pail, inside of which was a layer of plaster of Paris and potassium cyanide crystals, to kill any captured insects. An electric light was positioned above the mouth of the cone. Insects attracted to the light would fall down the cone, coated with varnish to prevent escape, and into the pail, where they would become ‘engaged in their desperate death struggles’. McNeill stated that his trap was especially good at collecting Hemipterans, Neuropterans, Dipterans, and Hymenopterans. It seems it was less suitable for collecting Lepidopterans, because beetles, less affected by the cyanide fumes, were presumably eating any moths that found their way into the trap (McNeill 1889). Since 1889, numerous versions of light traps designed to collect especially hemtaophagous insects of medical importance have been designed and patented. One of the most recognizable, the New Jersey light trap, was designed in 1932 (Headlee 1932), but its weight and dependence on heavy batteries or direct connection to an AC outlet for power made it impractical for large-scale field surveillance. The New Jersey light trap was nevertheless used, mainly for mosquitoes, for several decades. New Jersey traps are still sometimes used today, particularly to compare contemporary and historical collection data. The CDC miniature light trap, a much lighter and more convenient alternative for vector sampling and surveillance, has become a widely used alternative. The CDC light trap was designed by Drs. Dan Sudia and Roy Chamberlain at the Centers for Disease Control in the early 1960s (Sudia and Chamberlain 1962), and was inspired by an injury resulting from trying to carry the batteries required for the New Jersey trap (CDC 2015). CDC light traps rely on a small light (incandescent, UV, or LED) above a fan that creates a downdraft, sucking attracted insects into a hanging collection container below. Today, CDC traps, and variations on them, remain some of the most widely used traps for collecting crepuscular and nocturnal insects. The original light traps utilized incandescent bulbs, which give off much of their energy as heat and red or infrared light. The majority of nocturnal insects perceive wavelengths in the green-to-ultraviolet (UV) range, and cannot see red or infrared light (Briscoe and Chittka 2001), limiting the effectiveness of incandescent bulbs as attractants. To address this problem, entomologists began outfitting suction traps in the 1950s and 1960s with UV bulbs, which dramatically increased the number of insects collected (Pfrimmer 1955, Tashiro and Tuttle 1959, Belton and Pucat 1967, Rowley and Jorgensen 1967). More recently, there has been interest in the use of light-emitting diodes (LEDs) for trapping. LEDs have the added benefits of being available in a wide range of colors, and emitting very specific wavelengths, which certain genera, species, or physiological states may respond to differently (Hoel et al. 2007, Cohnstaedt et al. 2008, Snyder et al. 2016). By the early 1950s, entomologists had also begun to supplement light traps with carbon dioxide (CO2), which increased the number of hematophagous insects collected compared to light alone (Brown 1951, Reeves 1951, Newhouse et al. 1966, Snoddy and Hays 1966). Since then, other attractive odor cues (usually, but not always host-related) have been identified and used (Gibson and Torr 1999, Takken and Knols 2010), including 1-octen-3-ol (Hall et al. 1984, Takken and Kline 1989, Kline et al. 1994), the BG lure (Biogents AG, Regensburg, Germany), and lactic acid (Dekker et al. 2002), among others. Advantages of Light Traps There are advantages for the widespread and common use of light traps for vector surveillance, including 1) small size and weight, 2) reasonably low cost, 3) ability to run remotely (e.g., on small, portable batteries and without 120–220 V household current) for fairly long periods or to be automatically controlled by light-sensitive switches and fan breeze-operated gates, and 4) ability to collect a large variety of species, including both sexes. As an example, the model 512 CDC trap (J.D. Hock Co., Gainesville, FL) is only 14 cm tall and 8 cm in diameter. Apart from batteries or collecting container, it weighs a mere 325 g. The size and weight translate into the ability of a researcher to carry several such traps a number of kilometers into very remote field locations (e.g., Mullens and Dada 1992). Such traps are small enough to be inconspicuous, and they can often be placed in areas of fairly high human activity without great risk of vandalism. Their small size also allows for placement into confined places a larger trap won’t fit (e.g., into small rock recesses or upper tree canopies). Advances in light bulb technology, specifically the small but bright LEDs and automated photo-switches, also reduce the electrical requirements, and potentially extend trap run life into periods of several days. The ability of such traps to run overnight or longer without human monitoring is of course a very powerful and practical advantage. Another significant advantage of light traps is that they do in fact collect a lot of insects, both in terms of numbers and diversity. Together with the above advantages, this can make light traps the tool of choice for initial general surveys. Perhaps no other single vector surveillance tool gathers a large variety of species so easily. While some biases are known with regard to physiological condition (see below), the light traps may collect at least some insects that may not be receptive to host cues, such as gravid or blood-fed female flies. Simultaneously, since the main targets for vector sampling are nematocerous Diptera, males do not feed on blood and may not respond to host chemical cues like CO2. Light traps may be the only easy way researchers can collect males for studies of that sex or for purposes of identification of vector species where the females are difficult to recognize morphologically. Despite these advantages, certain limitations of light traps merit caution when interpreting collection data. Limitations of Light Trap Efficiency Decreased Attraction With Ambient Light Since the 1930s, the reduced ability of light-baited suction traps to collect hematophagous Diptera during nights on or near the full moon has been documented repeatedly (Bradley and McNeel 1935, Horsfall 1943, Pratt 1948, Provost 1959, Barr et al. 1963, Bidlingmayer 1967, Miller et al. 1970, Stryker and Young 1970, Bishop et al. 2000, Meiswinkel and Elbers 2016). This may be true even though some nocturnal biting flies fly more abundantly under full moon conditions, as was shown in flight interception studies of the bluetongue virus (BTV) vector Culicoides sonorensis Wirth and Jones (Diptera: Ceratopogonidae) (Nelson and Bellamy 1971, Akey and Barnard 1983, Linhares and Anderson 1990), as well as for several mosquito species (Bidlingmayer 1964). Perkin et al. (2014) recently showed that substantial artificial lighting (streetlights) can increase general activity of insects, including aquatic Diptera in the area, relative to darker zones. This increased activity does not appear to necessarily translate into larger trap catches. It is thought that ambient light reduces the contrast around the light trap, and so reduces its attractiveness to insects, or that artificial light sources compete with light traps. Even moderate amounts of ambient light may be enough to reduce collections in specific light traps, as those light traps only efficiently collect insects when ambient light intensity measures 0 lux (Meiswinkel and Elbers 2016). Because smaller catches on brighter nights could be misinterpreted as changes in the overall vector abundance in an area, it is important to note that simultaneously deployed, alternative trapping methods, including animal-baited collections (Pratt 1948), truck-traps (Provost 1959), and un-baited suction traps (Bidlingmayer 1967), do not show the same variation. While the reduced ability of light traps to collect insects a few nights a month near the full moon might be a minor problem, global light pollution exacerbates these concerns. It is estimated that light pollution increases by 6% every year, and extends even into historically rural or isolated areas (Gaston et al. 2012). Increased levels of nightly ambient light decrease nocturnal insect collections. As social progress has brought power and a better standard of living to many developing areas of the world, the amount of ambient light in areas that were previously rural and dark has increased dramatically. The ability to detect vector activity using light-baited suction traps in these areas will likely decrease over time, and research on the effect of artificial lighting on vector surveillance in urban areas is needed. Limited Range of Attraction Perhaps related to their reduced effectiveness with ambient light, light traps may vary drastically in their range of attraction. Studies with Culicoides biting midges differ in their conclusions about the range of light traps. Using CDC light traps in Denmark, Kirkeby et al. (2013) estimated their range of attraction to be ~15 m. Rigot and Gilbert (2012) found the range for Onderstepoort traps, which have a more powerful light and fan than CDC traps, to be as great as 30 m. However, other field studies have demonstrated far smaller ranges for Culicoides, even using the same traps. Venter et al. (2012) for example, demonstrated that the range of attraction of Culicoides to an Onderstepoort light trap in South Africa was only 2–4 m. Studies on Phlebotomine sand flies have also demonstrated that light traps have a short attraction range of 2–6 m (Killick-Kendrick et al. 1985, Valenta et al. 1995, Alexander 2000). Odeyoyinbo (1969) found that the range of attraction of Anopheles spp. in Africa to CDC incandescent light traps was ~5 m, while Costantini et al. (1998) found their range of attraction to be even less. These differences in the observed range of attraction are likely due to a number of potential confounding variables, including the type of trap used (and by extension, type of light used), species, ambient light, and the heterogeneity of the surrounding landscape. Because of this variation, comparisons of light trapping data across studies should be made with caution. Wavelength Differences The technological progression from incandescent light bulbs to UV to LEDs in insect trapping was driven by their increasing ability to collect both more species, as well as higher overall numbers of insects (Belton and Pucat 1967, Rowley and Jorgensen 1967, Cohnstaedt et al. 2008, Venter et al. 2009). Burkett et al. (1998) found that several Florida woodland mosquito species showed distinct color preferences, especially towards blue or green, when encountering CDC traps baited with both LEDs and CO2, though in many cases, incandescent light was equally attractive. In South Korea, LED baited traps producing a 365 nm peak were compared to traps using a 4 W UV bulb producing a less intense, but more diffuse mix of short wavelengths (Kim et al. 2017). The LED traps collected twice as many mosquitoes (188,125 vs 92,230) as well as more genera (12 vs 10), and more species (17 vs 14). LEDs have the advantage over traditional black lights of being available in very specific wavelengths, so they can emit not only in the traditional UV range, but in other colors as well. The availability of numerous wavelengths provides researchers with more tools to improve trapping methodology. The use of specific wavelengths could be an important study design consideration as some species are able to differentiate wavelength changes of only 10 nm (Snyder et al. 2016). Green light has been shown to be more attractive to at least some species of Culicoides than blue light is (Bishop et al. 2006, Hope et al. 2015, Gonzalez et al. 2016). In one study the important BTV vector, Culicoides obsoletus (Meigen) (Diptera: Ceratopogonidae) responded equally to most wavelengths, but less so to red light (Hope et al. 2015). In Egypt, phlebotomine sand flies were shown to be more attracted to red light. There, CDC suction traps baited with blue or green LEDs collected only about one-fifth the number of sand flies (Phlebotomus and Sergentomyia spp.) collected by traps baited with red LEDs (Hoel et al. 2007). Triatomine bugs are more attracted to blue lights, rather than green, yellow or red (Pacheco-Tucuch et al. 2012), but actually show a stronger attraction to incandescent light than UV (Minoli and Lazzari 2006). Certain species of insects may respond to different wavelengths of light (Bishop et al. 2006), and even feeding status may influence attraction (Snyder et al. 2016). In general, different wavelengths are often differentially attractive to different vector species. One must test and appreciate this in interpreting field light trap collections epidemiologically, if inter-species comparisons are intended. As data on these differing responses accumulates, LEDs will become increasingly valuable vector surveillance tools. Catch Size Although LED-baited traps sometimes enhance vector catch size over incandescent or UV-light baited traps, light trap collections in general are often smaller than those from traps baited with host cues, or from the host itself (Newhouse et al. 1966, Kline and Mann 1998, Gerry et al. 2009). Even species that can be collected well in light-only traps are often collected in significantly larger numbers in traps baited with semiochemicals, like CO2, lactic acid, or 1-octen-3-ol (Newhouse et al. 1966, Snoddy and Hays 1966, Stryker and Young 1970, Herbert et al. 1972, Alexander 2000, Andrade et al. 2008, Farajollahi et al. 2009, Kasap et al. 2009, Harrup et al. 2012). These chemical baits mimic the natural cues that host-seeking insects use for activation or orientation in the process of seeking a blood meal. As a result, these traps collect larger numbers of active insects of epidemiological interest, such as host-seeking females. Increasing collection sizes is a critical component of trap efficiency. Small collections are difficult, if not impossible, to analyze statistically. In cases where natural pathogen infection rates in vector populations are low (as is often true for arboviruses), small collections are far less likely to include any positive individuals than larger collections. Limitations of Light Trapping for Population Data Sex Biases Because male and female insects may respond differently to light, light trap collections are frequently biased towards one sex. Typically, these biases seem to be towards females (Belton and Pucat 1967, Venter et al. 2009, McDermott et al. 2016). Females of the sand fly, Lutzomyia whitmani (Coutinho and Antunes) (Diptera: Psychodidae) are much more attracted to light than males. In Brazil, suction traps with light were 76% L. whitmani females, whereas females comprised only 22% of catches of that species when the light was missing (Campbell-Lendrum et al. 1999). In some instances, males may make up a large proportion of collections, but even when considering the same species and same attractant, differences in the sex ratio between trapping locations may exist. One study found that C. sonorensis males made up 13.8% of UV trap collections on average at one dairy site in California, but over 45% of UV collections at a second dairy. In comparison, the proportion of males in simultaneously deployed CO2 baited traps was more similar between farms (4.8 vs 9.7%) (McDermott et al. 2016). It can be difficult to ascertain causes of location-specific differences in relative collections of males versus females. Males of a number of blood-feeding Diptera are more likely to swarm or seek mates close to larval developmental sites, however, and some species will orient to host animals or related chemical cues such as CO2 in order to locate females (Yuval 2006). The consequences of a sex-biased collection vary depending on the goals of the study. In the nematocerous Diptera, males are less epidemiologically important, and so their underrepresentation in light trap collections may not be a major concern. However, broader studies of biology or ecology of a given vector species may suffer from a sole reliance on light traps. In studies where the collection of male individuals is vital, such as for species identification or to understand mating patterns, male-targeted (e.g., seeking out swarming sites) or less biased collection methods (e.g., vehicle-mounted nets, or aerial sweeping) should be employed (Sanders et al. 2012). Parity Biases Differences in the proportions of parous and nulliparous females collected in light traps present more of a problem for epidemiological studies. Many of the pathogens of interest for human and animal health are not frequently or ever transovarially transmitted, and so only parous females (which have previously taken a blood meal from a host) are potentially infected. Because of this, and because of the costs associated with testing insect samples for pathogens, testing only parous females increases both economic efficiency, and the chances of detecting pathogens. Culicoides midges are unique in that they can be sorted visually by parity rather quickly and easily based on parous pigment deposited in the abdominal cuticle (Dyce 1969, Akey and Potter 1979). Other groups of hematophagous insects require more involved, subjective, or laborious techniques such as dissection of the ovaries, or inexact indicators such as wing fray, to determine parity or age (Hayes and Wall 1999, Hugo et al. 2008). Parity sorting thus can quickly become cost- and time- prohibitive, although it can provide invaluable population-level information on vector survival, and thus may be worth the effort. Even for large collections of Culicoides, the time required for visual parity sorting is a concern. Traps that preferentially collect parous females could significantly improve the efficiency of detecting pathogens. Nulliparous female C. sonorensis in the United States and Culicoides brevitarsis Kieffer (Diptera: Ceratopogonidae) in Australia seem to respond relatively poorly to light, and therefore may make up a smaller proportion of catches in UV-baited traps than in CO2-baited traps (Bellis and Reid 1996, McDermott et al. 2016). In a recent study, the proportion of parous females (compared to nullipars and males) collected did not differ between those two attractants (McDermott et al. 2016), so there may not be an advantage to using UV baited traps in terms of pathogen detection. Gravid C. sonorensis do seem to be fairly attracted to light (Anderson and Linhares 1989), so UV traps are useful in their collection. Alternatively, light traps have been criticized for their bias towards unfed/nulliparous female mosquitoes (Reisen and Pfuntner 1987, Githeko et al. 1994, Williams and Gingrich 2007). It is especially important to consider parity rates in collections used to estimate infection rates of mosquito-borne viruses as collections are almost always tested as unsorted pools of parous and nulliparous females. This can negatively affect infection rate estimates. For example, Williams and Gingrich (2007) compared West Nile virus (WNV) infection rates of mosquitoes collected using simultaneously deployed gravid traps and CO2-baited CDC light traps in Delaware. They found that gravid trap collections had estimated infection rates 32 times higher than the light traps, despite light traps collecting 57,000 more insects. Even though their collections are smaller, gravid traps have been recommended as a more efficient means of detecting pathogens in collected mosquitoes (Reisen and Pfuntner 1987, Williams and Gingrich 2007) because barring autogeny, all females have taken at least one blood meal, minimizing the need to sort by parity. Another important consequence of parity error is its effect on survivorship estimates, which are especially relevant for pathogens biologically transmitted by hematophagous Diptera. Probability of daily survival determines how well vectors survive the required extrinsic incubation period, and is highly influential in vectorial capacity calculations (Reisen 2009). In a simple but commonly used form, daily survival is derived by raising the proportion of parous females to the nth power, with n being one divided by the number of days required for a gonotrophic cycle (Davidson 1954). Using one 10-wk long California suction trap study on C. sonorensis (Anderson and Linhares 1989) as an example, a trap with CO2 (no light) had 32% parity, a trap with a UV light plus CO2 had 51% parity, and a trap with UV only had 98% parity (mostly gravid flies). The authors did not attempt to calculate survival, recognizing the inherent biases of the trapping, but, extrapolating from their data to make the point (assuming a 4 d gonotrophic cycle length), this would yield rough estimates of daily survival of 75%, 85%, and over 99%, with consequent impacts on calculated vectorial capacity. Certainly the last estimate would be completely unrealistic if used in vectorial capacity, and it would be based on biased parity information that did not accurately reflect either the vector population as a whole or the biting portion of that population. All of these estimates may also differ from a survival estimate calculated from direct animal aspiration collections, an arguably more realistic way to generate a population snapshot. Limitations of Light Trapping for Pathogen Detection Species Composition Perhaps the most critical overall limitation of light traps is their potential to inaccurately reflect the species composition of host-biting insect communities in an area. Many vector species respond poorly or unevenly to light, resulting in species biases in light trap collections. Studies that rely solely on light traps for collections run the risk of missing important species, and drawing false conclusions about transmission risk and putative vectors. The risk of misinterpreting light trap data is most severe in areas where many potential vector species exist, and are of unknown significance for transmission. This occurs regularly in neglected areas, or with emerging pathogens. This problem can even occur in such a well-studied region as Western Europe. Researchers and regulatory agencies there were unprepared for the large, persistent BTV outbreaks during 1999–2009 (Purse et al. 2015) primarily because many countries, without a compelling economic reason, just had not yet studied the main vector genus (Culicoides). Consequently, many locations were at the point of initial exploratory Culicoides community surveys as the outbreaks began, relative to their far better knowledge of mosquitoes or ticks. Studies on both mosquitoes and biting midges have addressed the problem of species diversity in light traps by comparing light-baited trap collections to those from host cue-baited traps, or to concurrent, direct collections from hosts. In the United Kingdom, while UV light-baited suction traps were efficient in collecting large numbers of C. obsoletus and Culicoides scoticus Downes and Kettle (Diptera: Ceratopogonidae), they only collected a small number of another potential BTV vector, Culicoides chiopterus Meigen (Diptera: Ceratopogonidae). Based on light trap data alone, it might have been concluded that C. chiopterus was not an abundant, and therefore important, vector in the United Kingdom. However, concurrent drop-trap collections from live sheep contained large numbers of C. chiopterus (Carpenter et al. 2008). Similarly, in Spain, direct aspiration collections from live sheep resulted in large numbers of the potential BTV vectors, C. obsoletus and Culicoides parroti Kieffer (Diptera: Ceratopogonidae), which were not well represented in collections from light traps deployed at the same time and place as the animal collections were made (Gerry et al. 2009). In France, light traps significantly underestimated the abundance of Culicoides brunnicans Edwards (Diptera: Ceratopogonidae) on sheep (Viennet et al. 2011). In Vietnam, a comparison of CDC traps baited with either CO2, light, or CO2 and light found that of all 23 mosquito species collected, the vast majority of individuals of a given species were collected by CO2/light combination (6.9–100%) or CO2-only (42.2–100%) traps. By comparison, light-only traps collected only 0–13.8% of the individuals of a given species (Herbert et al. 1972). In Florida, CDC traps baited with both CO2 and light collected a total of 17 mosquito species (Aedes, Anopheles, Culex, Culiseta, Psorophora, and Uranotaenia spp.), compared to only seven species in light-only traps. No Psorophora or Uranotaenia spp. were collected in traps without CO2 (Kline and Mann 1998). In Tanzania, CDC light traps were found to underestimate human biting rates of both Anopheles and Culex spp. (Govella et al. 2011), although other studies in Africa have shown light trap collections to be an acceptable measure of attraction to human hosts (Costantini et al. 1998), especially when deployed inside homes. In Korea, a comparison of UV and LED light baited traps for collecting Japanese encephalitis virus (JEV) vectors found that neither was particularly good at collecting Culex tritaeniorhynchus Giles (Diptera: Culicidae), the primary JEV vector in the area. Cx. tritaeniorhynchus females made up only 0.48 and 0.58% of UV and LED trap collections, respectively (Kim et al. 2017). For situations where the vector species are few and well known, such as Anopheles vectors of human Plasmodium in parts of Africa, relating light trap catches to biting and transmission risk becomes somewhat more tractable. Briet et al. (2015) conducted a meta-analysis of 13 studies that used the human landing catch (HLC) together with CDC light trap sampling. Those authors concluded that collections in CDC traps were at least approximately proportional to HLC, although studies were tremendously variable, and reliable generalizations about the relationship were impossible to derive. They argued that such data were still useful despite the imprecision, given the ease of use and safety of light traps, and that it was the very large changes in risk that were most epidemiologically relevant. Another important consideration for the use of light traps to collect mosquito vectors is the growing importance of day biting, highly anthropophilic Aedes spp., especially Ae. albopictus (Skuse; Diptera: Culicidae) and Ae. aegypti (L.; Diptera: Culicidae). In the last 30 yr, several Aedes-transmitted viruses, including dengue virus, chikungunya virus, and Zika virus, have emerged across the globe, causing massive public health problems (Roth et al. 2014). Aedes spp. competent for these viruses are widely established, including in temperate zones (Kraemer et al. 2015). Aedes spp., and Ae. albopictus and Ae. aegypti in particular, are not well collected in light traps (Morris and Defoliart 1969, Krockel et al. 2006, Farajollahi et al. 2009), and even CO2 baited traps are not particularly effective (Krockel et al. 2006). By comparison, human scent-baited suction traps without light, like the BG-Sentinel trap (Biogents AG) have proved to be much more effective collectors of Aedes vector spp. (Krockel et al. 2006, Maciel-de-Freitas et al. 2006, Farajollahi et al. 2009). The BG trap in particular was also shown to collect fewer nullipars than pars or blood-fed females (Ball and Ritchie 2010), making it a potentially more efficient means of sampling day-biting Aedes populations for viruses. As with the highly anthropophilic mosquito species, UV-light baited suction traps may be of little use for collecting anthro- or zoophilic phlebotomine sand flies (Rutledge et al. 1975, Andrade et al. 2008). Host odor cues, including CO2 (Kasap et al. 2009), 1-octen-3-ol, and the BG-lure (Andrade et al. 2008) can increase sand fly collections. For example, Muller et al. (2015) did have success catching large numbers of Phlebotomus papatasi (Scopoli) (Diptera: Psychodidae) using UV or incandescent light with CDC traps, especially if they also were baited with CO2 and other semiochemicals. Similarly, the widespread sole reliance on light traps to collect Culicoides midges has reinforced the common assumption that biting midges are entirely crepuscular and nocturnal, and not active during winter months. While not an unfounded generalization for the genus, this is not strictly true, as there are Culicoides species that are mainly diurnal, such as the suspected Australian BTV vector Culicoides actoni Smith (Diptera: Ceratopogonidae) (Bellis et al. 2004). Light traps on their own could quite possibly miss such species entirely. More importantly, during fall and winter, Culicoides often display a shift in their activity patterns, and a normally crepuscular or nocturnal species in summer can occasionally (or perhaps only) be collected several hours before sunset when temperatures are favorable for flight during cooler months (Lillie et al. 1987, Sanders et al. 2012). A clear demonstration of this occurred in year-round flight interception studies of C. sonorensis activity in Colorado (Barnard and Jones 1980). There, essentially all the adult activity in spring (April) and fall (November) was before sunset, when light traps were expected to be ineffective. In the summer in temperate climates, the BTV vectors C. obsoletus complex and C. chiopterus can be collected using sweep nets throughout the 24-h period, but light traps only collect insects after the sun has completely set and light intensity measures 0 lux (Meiswinkel and Elbers 2016). In fact, Meiswinkel and Elbers (2016) found that an Onderstepoort light trap operating from 2100 to 2200 h (dusk) collected only one C. obsoletus complex individual, while simultaneous aspiration directly off of sentinel cattle during just that 1-h period resulted in collections of 796 C. obsoletus complex and 709 C. chiopterus. C. chiopterus peak activity was found to actually occur more than 2 h before sunset. In Spain, Gerry et al. (2009) caught many female Culicoides before sunset, and numbers of mammal-feeding insects (C. obsoletus and C. parotti) aspirated from sentinel sheep over short (5 min) periods far exceeded the cumulative number taken in UV-baited CDC light traps running continuously nearby. The Gerry et al. (2009) study emphasizes again that one must know something about host feeding patterns when interpreting light trap catches. The most common Culicoides species in the light traps, by far, was Culicoides circumscriptus Kieffer (Diptera: Ceratopogonidae). Without prior knowledge of the species, particularly in the beginning stages of vector investigations, one might assume that it was a major potential vector of a virus like bluetongue. However, this species was entirely lacking from the sheep collections because C. circumscriptus is a bird feeder (Braverman and Linley 1994). With groups like the sand flies, where species tend to be restricted to certain heights within a vertical space, species biases in light trap collections can also occur. Due to the nature of the attractant, these traps have a relatively limited collection range in vertical space. Phlebotomine sand fly species that are active either lower to the ground or higher up are unlikely to be sampled consistently using light-baited suction traps (Alten et al. 2015). The use of host-cues, or hosts, in place of or in combination with light can increase sand fly numbers in collections. Campbell-Lendrum et al. (1999) placed CDC suction traps for Lutzomyia spp. directly above potential bait animals (humans, chickens, dogs), and found that traps with light collected more L. whitmani and Lutzomyia intermedia (Lutz and Neiva) (Diptera: Psychodidae) than did similarly placed traps without light. Compared to L. whitmani, L. intermedia collections were relatively more enhanced by the addition of light, suggesting that these species differ in their attraction to light. Light traps collected larger total numbers of the potential Peruvian Leishmania vector, Lutzomyia peruensis Velez (Diptera: Psychodidae) than HLC collections, but HLC collections contained a higher proportion of this species (45/955 total sand flies in light traps vs 80/3029 in HLC) (Davies et al. 1995). Once again, prior knowledge of the biology of a particular species is necessary for study design in this case. Infection Prevalence Rarely, cases might arise where the use of light traps has a direct negative impact on pathogen detection or transmission due to vector behavior. One compelling field example is the evidence that BTV infection apparently causes C. sonorensis midges to become averse to UV light. Work by Mayo et al. (2012) and McDermott et al. (2015) in California showed that BTV infection rates in pools of parous C. sonorensis collected using UV-light (315–400 nm, peak at 350 nm) baited CDC suction traps, or traps using both CO2 and UV-light, were significantly lower than in pools collected using CO2-only baited traps. UV trap collections underestimated the infection rates seen in CO2 trap collections by as much as 8.5 times (McDermott et al. 2015). UV wavelengths above 320 nm have essentially no microbiocidal properties and are well above the far higher energy, shorter wavelengths (250–260 nm) used for disinfection (Wolfe 1990, Bintsis et al. 2000), making it highly unlikely that the observed lower infection rates were due to UV degradation of BTV. This is suggestive then of behavioral manipulation of C. sonorensis by BTV, causing infected midges to avoid, or possibly be repelled by, UV light. UV traps were particularly poor at collecting infected midges when the overall number of midges in an area was low. Importantly then, there is the risk of missing active, BTV-infected vectors when vector abundance is low, such as periods of perceived overwintering or following the introduction of infected vectors into virus-free areas, inhibiting the ability to enact early eradication measures. Finally, a second rare, but possibly important, consideration when using light traps is the risk of drawing infected vectors into close proximity with hosts. Although the range of attraction to light for many species is limited (as discussed previously), some vector groups are highly phototactic, and artificial lights can influence natural dispersal behaviors, with unintended consequences for pathogen transmission. In historically rural, Chagas disease-endemic regions, urbanization, improved home construction, and aggressive vector control programs have helped to eliminate the traditional domestic Triatomine kissing bug vector species, but the increase in outdoor electric lighting was accompanied by a subsequent increase in orally transmitted Chagas and shift to sylvatic and peridomestic vector species (Barghini and de Medeiros 2010). These typically outdoor associated species are drawn to houses by bright outdoor lights (Pacheco-Tucuch et al. 2012), where they defecate the infectious metacyclic trypomastigote stage of the parasite onto fruits or other food grown or held nearby. The contaminated food is then consumed by people or animals, resulting in disease (Walter et al. 2005, Barghini and de Medeiros 2010). Although specifically discussed here in terms of highly phototactic Triatomine vectors, all attractive traps could potentially pose this risk, and placement of traps near hosts deserves careful consideration by researchers. Conclusions Light traps are one of, if not the most, widely used vector surveillance methods available. They are both practical and portable, and lend themselves to long-term surveillance studies. In some cases, they may be the only readily available option. However, light traps have potentially serious limitations, especially if used as the sole sampling tool. Researchers must understand the pros and cons associated with light traps when making decisions about their study designs. Ease of use should not outweigh the risks of not identifying vector species or misunderstanding pathogen activity in an area. This holds especially true when light trap data are being used to make important public or livestock health decisions, like when to make pesticide applications or transport animals. The efficacy of light traps for vector collection is highly dependent on assumptions about vector behavior. It is critical to take into account the published data that challenge these assumptions when choosing traps or attractants for medical and veterinary entomology field studies. Light trap collections can be biased towards certain species, sexes or physiological states. Competent vector species may not be strongly phototactic, and their low numbers in light trap collections could lead to their dismissal as important for pathogen transmission. Likewise, when light traps preferentially collect nulliparous females or males, the ability to detect pathogens in pooled insect samples drops. Light traps do not efficiently collect day active or highly anthro- or zoophilic species, so light trap collections may lead to a misunderstanding of vector activity patterns. In some cases, pathogen infection may cause vectors to alter their phototactic behavior, resulting in errors in transmission risk estimates, as has been suggested for BTV-infected midges. Because these ‘subtle effects’ of pathogen manipulation (those that have no obvious impact on transmission) are largely unstudied, other vector-pathogen interactions in other disease systems may exist that limit the effectiveness of light traps. For highly phototactic species, the use of light traps may put people and animals at risk by drawing large numbers of vectors into areas near hosts, as has been hypothesized with Chagas disease. In rural areas, researchers may want to consider whether to place light traps away from livestock enclosures and habitations. In areas where light traps are the only logistically realistic option for vector surveillance, the use of LEDs may improve efficiency by allowing for the use of targeted wavelengths. LED baited traps have been shown to collect higher numbers of midges, mosquitoes, and sand flies compared to UV or incandescent lights, and may also collect a higher diversity of insects. LEDs in specific wavelengths can be selected for surveillance programs or field studies in order to effectively target certain vector species of interest. That being said, ambient light, which is increasingly a problem in our developing world, significantly decreases the effectiveness of all light traps. Even small amounts of ambient light have been shown to reduce the number of insects collected in a given trap. The effects of light pollution across the globe have been discussed in any number of contexts, but perhaps it is time medical and veterinary entomologists began to consider more seriously the effect it may have on vector surveillance. There is no such thing as a perfect insect sampling technique, especially when the stakes are as high as they can be with vector borne diseases. All trapping methods will have advantages and disadvantages. That does not mean, though, that critical flaws in a method should be ignored because of convenience. Today, it is clear that medical and veterinary entomology could benefit from innovative alternatives or supplements to the light baited suction trap. The best way to improve the interpretation of light trap collections, and indeed all trap collections, is to continue to collect data on how they compare to other trapping methods and other light trap studies for as many species and in as many situations as possible. When conducting surveillance studies, researchers should consider including multiple attractants (e.g., UV vs CO2 or UV vs LED) for comparison to traditional light traps. When running long-term vector surveillance programs using light traps (such as those conducted by vector control districts), periodic collections using host-cue baited traps, or other non-light methods, to check light trap assumptions should be considered. Light traps will continue to have a place in medical and veterinary entomology, and in some cases may truly be the most appropriate choice. However, more broad and critical discussion about the validity of the epidemiological conclusions drawn from these collections is needed in order for entomologists and epidemiologists to avoid the dark side of light traps. Acknowledgments We would like to thank William Reisen and Christopher Barker for their input and discussion on mosquito surveillance trends and helpful comments on this article, as well as Alec Gerry and Lindsey Garver, who also provided generous feedback to improve the article. The material to be published reflects the views of the authors and should not be construed to represent those of the United States Department of the Army or the United States Department of Defense. References Cited Akey, D., and Potter H.. 1979. Pigmentation associated with oogenesis in the biting fly Culicoides variipennis (Diptera: Ceratopogonidae): determination of parity. J. Med. Entomol . 16: 67– 70. Google Scholar CrossRef Search ADS PubMed  Akey, D. H., and Barnard D. R.. 1983. Parity in airborne populations of the biting gnat Culicoides variipennis (Diptera: Ceratopogonidae) in northeastern Colorado. Environ. Entomol . 12: 91– 95. Google Scholar CrossRef Search ADS   Alexander, B. 2000. 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