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A review of available methods and description of a new method for eliminating ectoparasites from bird nests

2015; Association of Field Ornithologists; Volume: 86; Issue: 3 Linguagem: Inglês

10.1111/jofo.12113

1557-9263

Amanda K. Hund, Jessica T. Blair, F. Hund,

Avian ecology and behavior

Bird nests offer an ideal situation to manipulate ectoparasites and study how they impact hosts. Several methods are available to eliminate parasites from nests and each has its own suite of advantages and disadvantages. For example, recent toxicity research has revealed that some commonly used insecticides may not be suitable for use in experiments with nestlings. This highlights the need for investigators to control for the effects of methods used to eliminate nest parasites within experimental designs. Methods that can be used across treatment groups are also often needed to study the effects of variation in parasite intensity. To aid investigators in deciding which method(s) to use, we provide a comprehensive review of available methods for eliminating nest ectoparasites and also describe a new heat gun method. We tested the effectiveness of the heat-gun method with nests of Barn Swallows (Hirundo rustica) to which 100 nest mites were added and then quantified the number of surviving mites and other naturally occurring arthropods. We found that fully heated nests had significantly fewer mites and other arthropods than partially heated or control nests. Use of the heat gun had no negative effects on nestling growth or mortality rates. In studies of avian nest ectoparasites, investigators need to consider methods that can be used across treatment groups to ensure that unaccounted for toxicity effects are not influencing results and leading to underestimation of the often subtle effects of ectoparasites on birds. Una revisión de los métodos disponibles y descripción de un Nuevo método para eliminar ectoparásitos de nidos de aves Los nidos de aves ofrecen una situación ideal para manipular ectoparásitos y estudiar cómo impactan en los hospedadores. Varios métodos están disponibles para eliminar ectoparasitos y cada uno tiene sus ventajas y desventajas. Por ejemplo, estudios recientes sobre toxicidad han revelado que algunos insecticidas comúnmente usados pueden no ser adecuados para su uso en experimentos con pichones. Esto destaca la necesidad por parte de los investigadores de controlar el efecto de los métodos usados para eliminar ectoparásitos de los nidos. Los métodos que puedan ser usados a través de grupos de tratamiento son con frecuencia necesarios para permitir el estudio de los efectos de la variación del número de parásitos. Para ayudar a los investigadores a decidir cual método(s) usar, proveemos una revisión de los métodos disponibles para eliminar de ectoparásitos y también describimos un nuevo método de pistola de calor. Hemos probado la efectividad de el método de pistola de calor en los nidos de la Golondrina común (Hirundo rustica) a los cuales se le agregaron 100 ácaros y luego se cuantificaron el número de ácaros y otros artrópodos sobrevivientes. Encontramos que los nidos que habían sido totalmente calentados tenían menos ácaros y otros artrópodos que nidos parcialmente calentados y nidos control. También encontramos que este método de pistola de calor no tiene efectos negativos en las tasas de crecimiento o mortandad de los pichones. En estudios de ectoparásitos de nidos de aves, los investigadores necesitan considerar métodos que puedan ser usados a través de grupos de tratamiento a fin de asegurar que efectos no considerados de toxicidad no estén influenciando los resultados y haciendo que uno subestime el muchas veces sutil efecto de los parásitos en las aves. Parasites are ubiquitous and can impose strong selection pressures on hosts that can have significant ecological and evolutionary consequences (Price et al. 1986, Poulin and Thomas 1999, Karvonen and Seehausen 2012). Parasites have long been known to influence the morphology, behavior, survival, and life history traits of their hosts (Poulin 1995, Windsor 1998). To understand these important and complex effects in wild host-parasite systems, studies that go beyond simply quantifying parasite abundance, prevalence, or diversity are needed (Poulin 2007). Experimental designs that include controlled manipulation of parasites are the most powerful way to determine causal relationships in host-parasite interactions (Clayton and Moore 1997, Owen et al. 2010). Birds have long been used as models to study host-parasite interactions, and many manipulative field techniques have been employed to understand these relationships (e.g., Loye and Zuk 1991, Møller 1991, Owen et al. 2010). For birds with altricial nestlings, nests can harbor ectoparasites and offer a unique situation for studying how parasites impact hosts during a critical developmental and life-history period (Rendell and Verbeek 1996). Because hosts (nestlings) are confined to nests, designing experiments with different parasite-exposure treatments is possible and such studies have been important in improving our understanding of the short- and long-term costs of parasites on their avian hosts (Moss and Camin 1970, Charmantier et al. 2004, Owen et al. 2010). Many methods have been developed to eliminate parasites from nests to create non-parasitized treatments for comparison with parasitized ones (Table 1). Eliminating parasites is also important for standardizing the addition of parasites to nests. By first eliminating parasites from nests, investigators can control the degree of infestation by adding certain numbers of parasites without having to account for pre-existing infections, and can ensure that nests are not infected by non-focal species of parasites. Several methods for eliminating parasites from nests (or at least reducing their numbers) have been developed, and each has potential advantages and disadvantages. In designing experiments, investigators must consider which method will best serve their needs and, to aid those making such decisions, we present a comprehensive review of available methods and also describe a new method. For each method, we provide a brief description of the technique and how it has been used. We evaluate each method based on: (1) effectiveness in removing/killing nest parasites, (2) persistence, or how long before parasites can re-infest nests, (3) possible use across treatments, (4) known effects on nestlings, (5) ease of use in the field, and (6) risk of use to investigator's safety. A list of studies where each method has been used, along with how it was applied and its effectiveness can be found in Supplemental Table S1. For the sake of brevity, we will refer to "eliminating" nest parasites from nests, but acknowledge that methods we describe may not always be 100% effective at eliminating all parasites from nests. Several insecticides have been applied to nests to kill or deter parasites. For experimental purposes, chemical insecticides persist in the nest for long periods of time (Ebeling 1963). This can be an advantage if experimental designs require nests to stay parasite-free for longer periods, but can be a disadvantage if investigators want to use these methods for parasite addition treatments. The long-lasting action of chemical insecticides also make it nearly impossible to control for the effects that they may have on nestlings because they cannot be used equally across treatments, e.g., nests with parasites and treated nests without parasites (López-Arrabé and Cantarero 2014). Toxicity studies required prior to insecticides being approved for use (overseen by government agencies such as the Environmental Protection Agency), typically involve use of adult birds with domesticated genetic lineages. These birds may have much higher percentages of body fat and may also be less sensitive to chemicals than birds in the wild (Ballantyne and Marrs 2013). In addition, different species of birds vary in their sensitivity to chemicals (Hill 1971, 1982, Tucker and Haegele 1971), and nestlings are often more sensitive than adults (Grue and Shipley 1984, Fry 1995, Wolfe and Kendall 1998, Parker and Goldstein 2000), with altricial nestlings being more sensitive than precocial nestlings (Hoffman et al. 2002). As a result, using insecticides advertised as being non-toxic to birds does not guarantee that they will have no effects on nestlings. Pyrethroids are the most frequently used insecticides for experiments involving nest ectoparasites and are the active ingredient in many flea powders and household and garden insecticides. Pyrethroids are compounds derived from flowers of the chrysanthemum genus (Chrysanthemum cinerariaefolium and C. coccineum) and are potent insect repellents and neurotoxins (Meurer-Grimes 1996). Most commercial pyrethroid insecticides contain permethrin or pyrethrin, and are often mixed with low levels of synergistic piperonyl butoxide (Clayton and Walther 1997). Although potent when applied, the effectiveness of pyrethroids declines as they biodegrade. However, they retain some effectiveness for days to weeks depending on the initial concentration used and exposure to air, water, and light; the soil half-life for pyrethroids is 12 days (Melendez et al. 2006). In many studies, investigators repeat pyrethroid treatments every few days during experiments to increase effectiveness over time (Merino and Potti 1998, Merino et al. 2001, Carleton 2008, Martínez-de la Puente et al. 2011, Harriman et al. 2014, López-Arrabé and Cantarero 2014). Pyrethroids have been found to effective at eliminating a wide array of nest ectoparasites, including mites, lice, fleas, blow flies, ticks, and martin bugs (Table S1). Based on data provided in studies where investigators used pyrethroids (Table S1), treatment with pyrethroids eliminated an average of 76% of nest parasites and repeated application eliminated an average of 96% of nest parasites. Pyrethroids are known to have low toxicity for adult birds and mammals (Casida 1973, Jackson 1985, Melendez et al. 2006) and studies have revealed no negative effects on adult birds with topical application (Tomás et al. 2008). However, pyrethroids are highly toxic to fish and amphibians (NPIC 1998, Todd et al. 2003). Some skin and respiratory irritation may result from exposure in humans, but these chemicals are generally thought to be safe for people (Melendez et al. 2006). Pyrethroids come in both liquid and powder forms, so nests can be dusted or sprayed. Pyrethroids biodegrade over time so have minimal lasting effects on nests across seasons or years (Meurer-Grimes 1996). Although apparently non-toxic for adult birds, pyrethroids may have negative effects on nestlings. López-Arrabé and Cantarero (2014) compared nestling Pied Flycatchers (Ficedula hypoleuca) in nests heat-treated (microwave), pyrethroid-treated, and control nests with parasites. Compared to nestlings in heat-treated nests, nestlings in pyrethroid-treated nests weighed significantly less, had shorter tarsi and wings, and lower levels of total glutathione, an important biomarker for redox state and cellular detoxification capacity. In addition, Garg et al. (2004) found that long-term exposure to synthetic pyrethroids (Fenvalerate) by young broiler chicks (Gallus gallus) did not affect body mass, but lowered total leucocyte count, reduced concentrations of active macrophages in the blood, raised serum adenosine triphosphate, and decreased the mass of the thalamus and spleen (Garg et al. 2004). Such results suggest that use of pyrethroids in studies of nest parasites may influence the results because possible side effects on nestlings that are not accounted for could introduce undesired systematic variability across treatment groups. This could lead to the effects of ectoparasites being underestimated compared to a pyrethroid-treatment group. Tetrachlorvinphos (sold as Rabon) is a commercially available organophosphate livestock insecticide that has also been used in avian research (Table S1). It comes in several forms, but, in experiments with nest ectoparasites, is typically diluted from a 50% solution to a 0.5% or 1% solution and then sprayed on nests using a hand-spray applicator (Norcross and Bolen 2002, Eggert and Jodice 2008, Eggert et al. 2010). Organophosphate insecticides inactivate acetylcholinesterase, an enzyme essential for nerve function, and are toxic to most insects (Walker et al. 1972). Investigators typically apply this chemical once during breeding and have reported an average of 85% reduction in parasites (soft ticks; Table S1). Tetrachlorovinphos biodegrades due to hydrolysis in natural conditions, with a soil half-life of 6–8 d (Hanley et al. 2006). Tetrachlorovinphos is highly toxic to fish and amphibians (Wolf 2008), was found to be carcinogenic in long-term studies of rodents, and is a cholinesterase inhibitor and endocrine disruptor (Walker et al. 1972, Wolf 2008). Protective clothing is recommended for spray application (Hanley et al. 2006). Tetrachlorovinphos has been used to reduce numbers of soft ticks (Ornithodoros capensis) in nests of Brown Pelicans (Pelecanus occidentalis) (Norcross and Bolen 2002, Eggert and Jodice 2008, Eggert et al. 2010). Based on dietary exposure studies (2000 mg/kg) with adult Ring-necked Pheasants (Phasianus colchicus), Mallards (Anas platyrhynchos), and Chukars (Alectoris chukar), tetrachlorovinphos has been approved as non-lethal for birds (Hanley et al. 2006, Wolf 2008). However, convulsions and tremors were noted in the days following treatment (Hanley et al. 2006, Wolf 2008). Young Domestic Chickens (Gallus gallus) exposed to similar organophosphate insecticides had significantly lower levels of serum acetylcholinesterase, higher levels of serum adenosine triphosphatase, lower total leucocyte counts, and reduced mass of immune organs compared to control chickens (Garg et al. 2004). Additional study is needed to fully understand how this chemical may affect developing nestlings, particularly in wild, non-model species. Chloroform (trichloromethane) is frequently used to remove parasites from the plumage of adult birds (Clayton and Walther 1997, Bush 2009, Hamstra and Badyaev 2009, Koop and Clayton 2013), but less often to remove nest ectoparasites (Saino et al. 1998). Chloroform is an organic compound that comes as a highly volatile liquid and is known to depress the nervous system (Jarboe et al. 2001). Saino et al. (1998) placed material from Barn Swallow (Hirundo rustica) nests in a plastic bag and added chloroform to anesthesize and count, but not kill, louse flies (Ornithomyia biloba) in nest material. Anesthesia from this treatment for louse flies lasted ∼30 min, after which most flies fully recovered (Saino et al. 1998). Chloroform can be useful if the goal is to anesthesize and collect live parasites, particularly insects like flies that may be mobile and difficult to catch and quantify. However, chloroform would not be suitable for reliably eliminating parasites from nests because it is not lethal. There is also a risk to researchers because chloroform is a known carcinogen and has immediate, potentially dangerous effects on the central nervous system (Du et al. 2001). Risk to nestlings is low because exposure to the chemical would be limited if chloroform is allowed to fully vaporize before nest material is returned to nests. Citronella oil has been used in nests of Great Tits (Parus major) and Blue Tits (Cyanistes caeruleus) to reduce numbers of flying blood-sucking insects, such as midges (Diptera ceratopogonidae) and black flies (Diptera simuliidae; Martínez-de la Puente et al. 2009, 2013). Citronella is an oil derived from lemon grass (Cymbopogon spp.) and contains several compounds that contribute to its insecticide properties, including aldehydes, terpines, and alcohols (Novak and Gerberg 2005, Kim et al. 2005). Citronella is thought to interrupt olfactory search mechanisms that biting insects use to locate hosts (Peterson and Coats 2001). Investigators that who sprayed citronella on nests reported reductions in the numbers of biting midges (87.5% reduction) and black flies (76% reduction), but it had no effect on the number of blow fly pupae (Martínez-de la Puente et al. 2009). These results suggest that citronella may not be suitable for eliminating many common ectoparasites that live exclusively in nests, such as mites, lice, fleas, and blow flies, but may be useful if researchers want to specifically repel flying insects. Citronella is a fully active repellent for 4–8 h, but may retain some repellent effect for several days after application (Brown and Hebert 1997, Martínez-de la Puente et al. 2009). For consistent high repellent effects in nests for periods longer than 4–8 h, reapplication would likely be necessary. Although citronella has been approved for use with several mammal species, there have been no toxicity tests with birds (Jarboe et al. 2001). Citronella poses no risk for researchers. Yarrow (Achillea millefolium) is a plant insecticide that has been used to repel ectoparasites in nests (Table S1). Yarrow is a perennial herb and is common across many parts of Europe and North and South America (Nemeth and Bernath 2008), and is often one of the plants selected by birds (e.g., European Starlings, Sturnus vulgaris) that incorporate fresh green vegetation into nest substrates (Wimberger 1984, Clark and Mason 1985). One hypothesis to explain this behavior is that these plants have insecticide properties that reduce ectoparasite burdens in nests (Wimberger 1984, Clark and Mason 1988). Aromatic hydrocarbons, mainly monoterpenes and isoprene, in the volatile oil that are part of the plant's defense against herbivory are apparently being used by birds to deter ectoparasites (Wimberger 1984, Clark and Mason 1985). Manipulative experiments with species that do not naturally incorporate greenery into their nests (e.g., Tree Swallows, Tachycineta bicolor) have provided mixed results concerning the effectiveness of yarrow at reducing numbers of ectoparasites (Gwinner and Berger 2006). Although some investigators found that addition of fresh yarrow into nests reduced flea numbers by as much as 50% (Shutler and Petersen 2003, Shutler and Campbell 2007), Dawson (2004b) found that fleas were actually more abundant (32% increase) in nests where yarrow had been added. In studies to date, investigators have reported an average decrease in numbers of fleas in nests of 23% with yarrow compared to control nests, but yarrow had no effect on numbers of blow flies (Shutler and Peterson 2003, Dawson 2004b, Shutler and Campbell 2007). Thus, yarrow may only be effective with certain ectoparasites. Shutler et al. (2003) placed new yarrow clippings in nests every 2–8 d, so frequent retreatment may be necessary to maintain effectiveness. Yarrow poses no apparent risk for birds or humans and, in fact, is a common component of many herbal supplements (Ernst 1998). Use of yarrow by investigators may be limited by the availability of fresh yarrow near nest sites. Diatomaceous earth (DE) is an inert dust consisting of fossilized diatoms and is composed of amorphous silicon dioxide. It is stable, non-toxic, and has been used in water purification systems and for treatment of internal parasites in livestock (Bingham et al. 1991). DE is most effective at killing crawling insects by abrading exoskeletons and absorbing waxy fats and lipids leading to water loss and, ultimately, death (Ebeling 1971). DE has been used primarily to reduce numbers of fleas (Ceratophyllus idius) and blow flies (Protocalliphora sp.) in nest boxes used by Tree Swallows (Dawson 2004a, Bennett et al. 2011). DE remains effective as long as the powder remains in a nest, making it undesirable for use across treatment groups. Some investigators have used a combination of DE mixed with pyretherin, commercially available as a powder mitocide (Shutler and Petersen 2003, Dawson 2004a), to reduce numbers of fleas and blow flies. Investigators using mitocide powder with a DE base have reported declines in numbers of mites (80%), blowflies (66%), and fleas (95%) in nests (Shutler and Petersen 2003, Dawson 2004a, Carleton 2008, Wiebe 2009). Negative impacts of DE on nestlings are likely minimal or non-existent. However, Hill (2000) suggested that using DE in a closed space, such as a nest box, may cause problems because nestlings that flap their wings as they approach fledging age can aerosolize DE, which may cause negative effects if inhaled. For large ectoparasites, such as blow fly larvae that live in nesting material, a physical barrier between nest and nestlings can prevent parasites from reaching and feeding on their hosts. For example, a small section of nylon fitted around a circular wire loop was placed on the floor of Darwin's Medium Ground Finch (Geospiza fortis) nests to prevent botfly larvae (Philornis downsi) that live in the bottom of nests from reaching nestlings (Koop et al. 2011). Koop et al. (2011) found that larvae did occasionally get around or through the lining and reach nestlings, so liners had to be examined, cleaned, and sometimes repaired each time nests were checked. Nylon has also been used to reduce contact between blow fly larvae and nestling Blue Tits (Parus caeruleus; Charmantier et al. 2004). Although this technique eliminates the risk of chemical side effects, there is still a risk that some parasites will reach nestlings. In addition, this method may work well for large, less mobile fly larvae, but smaller nest ectoparasites such as mites, lice, ticks, or fleas might be more likely to crawl around or through the liner to reach nestlings. Nest liners are thought to have minimal effects on nestlings, but the lining may interfere with the thermal environment of the nest because the lining is placed between nestlings and nest material such as feathers (Møller 1984, Lombardo et al. 1995). Birds that reuse old nests from previous breeding seasons may have an increased risk of exposure to ectoparasites with life stages that overwinter in nests (Barclay 1988, Møller 1990). As such, investigators that remove old nests from nest boxes between breeding seasons may reduce the risk of ectoparasite exposure for birds that use the nest boxes the next breeding season (Møller 1989). However, several studies have revealed no significant difference in ectoparasite numbers between nest boxes with and without old nests (Johnson 1996, Pacejka and Santana 1996, Allander 1998, Blem et al. 1999, Mazgajski 2013). Mappes et al. (1994) even reported greater numbers of ectoparasites in nest boxes where old nests had been removed. In contrast, however, leaving old nests in nest boxes may result in increased numbers of fleas that overwinter in nests (Rendell and Verbeek 1996, López-Arrabé et al. 2012). Compared to nest boxes where nests were not removed, cleaned boxes were found to have an average of 16% fewer fleas (Table S1), with the greatest reduction in flea numbers being 52% (Rendell and Verbeek 1996). Investigators have reported no effect of old nest removal on numbers of mites, which are also thought to overwinter in nests (Pacejka and Thompson 1996, Blem et al. 1999, López-Arrabé et al. 2012). The presence and numbers of nest ectoparasites can be manipulated by physically removing nests and disinfecting them in the field and then replacing them. Nests can also be exchanged for ones that have been previously collected and disinfected (Rendell and Verbeek 1996, Charmantier et al. 2004, Moreno et al. 2009, Brommer et al. 2011). This method is typically used with species of birds that nest in nest boxes or natural cavities, where nests are easy to remove and replace, and can be very effective at removing parasites because nests can be microwaved (Rendell and Verbeek 1996, Charmantier et al. 2004, Moreno et al. 2009, Brommer et al. 2011). For example, Richner et al. (1993) found that placing nests in a microwave (750-850W) for 1–5 min killed all arthropods. To prevent desiccation when microwaving, nests can be placed in a sealed plastic bag and even sprayed lightly with water (López-Arrabé et al. 2014). Although this method can effectively eliminate a variety of nest ectoparasites (e.g., mites, fleas, and blow flies), nests can be re-infected as soon as they are replaced (unless recently disinfected with insecticide; Rendell and Verbeek 1996, Charmantier et al. 2004, Moreno et al. 2009, Brommer et al. 2011). Advantages of this method are that it can be used across treatment groups and to standardize ectoparasite addition treatments. To maintain ectoparasite-free nest environments, nests can be replaced several times during the breeding period (Charmantier et al. 2004). For nests in nest boxes, cleaning nest boxes before returning treated nests is also important (Rendell and Verbeek 1996). Investigators have done this using wire brushes and by passing the flame of a butane torch over cracks on the inside of nest boxes to remove or kill any remaining ectoparasites (Rendell and Verbeek 1996, López-Arrabé 2014). Although effective, use of replacement nests is limited to species of birds where nests can be easily replaced. It is not feasible for many species of birds because nest integrity, structure, and stability could be compromised by the exchange. Dry heat is an effective way to kill small arthropods such as mites, fleas, bed bugs, and lice (Nicholson and Rotberg 1996, Mourier and Poulsen 2000), and an effective way to deliver dry heat in the field is with an industrial heat gun. Heat guns can be found at most hardware stores. Several models are available, but one with multiple temperature settings, particularly in the 95°C–450°C range and with a high-powered fan to deliver hot air throughout nests is best. If field sites have electricity, an electric heat gun and extension cord can be used, but butane and battery-powered heat guns are also available for use at sites without access to electricity. We have successfully used both models in our studies (Fig. S1). To eliminate ectoparasites from nests, nests are heated in situ by holding the heat gun about 15 cm away and moving it slowly, but continually, over the entire surface of a nest. With Barn Swallow (H. rustica) nests, we first heated the inside of the nest cup and then the outer shell, bottom of the nest, and area around the nest, and then repeated this pattern to insure thorough heating. Nest temperature is tracked during heating with a battery-powered digital infrared thermometer. Following guidelines from the pest-control literature, we heat all parts of nests to at least 145°C, which usually takes <5 min (Nicholson and Rotberg 1996). All nest material is left in nests during this process. Barn Swallow nests in our studies are made of mud and lined with twigs, horsehair, and feathers. During the heating process with a heat gun setting of 260°C, we have found that the horsehair occasionally deforms when exposed to the heat, but the other nest materials are not affected. Auto-ignition temperatures of most nesting materials used in Barn Swallow nests, such as feathers and small sticks, are above 200°C, which is higher than our treatment temperature range (Shen et al. 2006). However, continual motion of the heat gun is needed to prevent ignition. When nests cool to <29°C, usually within ≤10 min, eggs or nestlings can be safely returned. To insure that nests remain ectoparasite free, repeat treatments may be necessary depending on the risk of ectoparasites becoming re-established in nests after initial treatment. To test the effectiveness of the heat-gun method, we conducted a study to quantify elimination of ectoparasites. We also used this method in a reciprocal cross-fostering experiment and quantified the reduction in numbers of ectoparasites and any negative effects on nestlings in heat-treated nests. Our study was conducted in July 2014 at a Barn Swallow breeding colony in Boulder County, Colorado. We used inactive nests because they had to be removed, but these nests were similar in structure to active nests. All nests in our study were still lined with horsehair, sticks, twine, and feathers. We added 100 live mites (Ornithonyssus sylviarum) to each of 15 nests to insure the presence of ectoparasites. We collected mites from active nests less than 48 h before they were added and then allowed them 24 h to infiltrate nest structures. We did not treat nests prior to adding mites, so nests had 100 mites plus any other ectoparasites and arthropods already present. After 24 h, we returned to the site and randomly assigned five nests to each of three treatments: not heated (control), heated to 60°C (half-heat treatment), and heated to 120°C (full-heat treatment). After treatment, nests were removed, placed in sealed plastic bags, and transported to the lab. Each nest was placed in a berlese funnel and the light and heat source (150 W halogen flood light 25 cm above nests) was turned on for 24 h. All living mites and other arthropods that emerged from the nests fell down the funnel and were collected in alcohol. Alcohol samples were examined under a dissecting scope and all mites and other arthropods were identified and quantified. Differences between treatment groups in the number of mites and other arthropods in nests were analyzed using a one-way ANOVA. For pair-wise comparisons, we used Tukey's HSD test. From June through September 2013, we monitored 752 nests at 27 locations in Boulder, Jefferson, and Weld counties in Colorado. Nests were checked every 3 or 4 d except near the end of the incubation period when they were checked every day to determine hatch date. We banded and measured nestlings at 12 d post-hatching (near fledging). The heat-gun method was used to experimentally manipulate ectoparasite presence in a subset of these nests (N = 94) as part of a reciprocal cross-fostering experiment. Experimental nests were heated to 120°C 2 d after hatching. Nestlings were removed during the heating process and returned when nests cooled to <29°C. Half of these nests then had 100 live mites added to them to create two treatment groups: treated with no mites ad