Anoplura

To confirm morphologic identifications of ectoparasite species, we subjected a representative subpopulation (≈20%) of the ectoparasites to DNA extraction and amplification of target genes (Appendix Table). For ticks, fleas and lice, we isolated genomic DNA (gDNA) by using the DNeasy Blood and Tissue Kit (QIAGEN, https://www.qiagen.com) according to the manufacturer’s instructions. We isolated gDNA from a small portion of the idiosoma of ticks (23 (link)) and from the anterior dorsal part of the abdomen of fleas (24 (link)). We selected individual lice and mites under an optical microscope and extracted gDNA by using a QIAamp DNA Micro Kit (QIAGEN).

Two similar but independent experiments were conducted at the Sea Lice Research Centre (Experiment 1) and the Institute of Marine Research in Bergen (Experiment 2) to determine the number of chalimus stages in L. salmonis. Experiment 1 used observations of abandoned L. salmonis exuviae in incubators to identify molting events prior to the preadult 1 stage. In experiment 2 morphometrics were used to identify the number of distinct morphological groups of L. salmonis chalimus larvae and to assess molting in incubators.
In both experiments Atlantic salmon (Salmo salar) were stocked in 500L fish tanks (1x1x0.5 m) with flow-through full salinity seawater (34.5‰, 10±0.3°C). The fish were infected with L. salmonis copepodids as described by Hamre et al. [17 (link)] using 150-200 copepodids fish^-1. The lice used belong to the laboratory strains LsGulen and LsOslofjord [17 (link),18 ]. Both experiments were carried out in strict accordance with Norwegian legislation and the experiment was approved by the Norwegian Animal Research Authority (permits nr. 2010/245410 and 2009/186329).

The reference spectra of each species were compared to evaluate the database. A blind test was then performed using new adult specimens from our laboratory-reared colonies and field-caught mosquitoes from Senegal that had corresponding reference spectra in our database (figure 1). For each species, 1 to 4 new specimens were used. Both molecular forms of An. gambiae (M and S) were carefully evaluated to identify a match with the corresponding database. We also tested 2 specimens of different arthropod species that were not included in our database, including ticks (Ixodes ricinus), fleas (Ctenocephalides felis), lice (Pediculus humanus corporis) and bed bugs (Cimex lectularius). The results are presented in the MALDI-Biotyper software as Log Score values that correspond to a matched degree of signal intensities of mass spectra of the query and the reference spectra, and these score values for species identification were obtained for each spectrum of the tested samples [35] (link).

We assessed the efficacy of permethrin by estimating blow fly (Protocalliphora sialia) load in each control and treated nest. Following past work (De Simone et al., 2018 ; Grab et al., 2019 (link)), we collected all nests in sealed bags after fledging (19–21 days post‐hatch). Nests were then dissected in the laboratory, by one observer (SZ) blind to the experimental treatment group, to count blow fly larvae and pupae. Permethrin effectively reduced ectoparasite load (zero‐inflated negative binomial regression: p = .003), where blow flies were found in 0% of permethrin‐treated nests and 43.8% of water‐treated nests (9.5 ± 4.27 per nest across all control nests; 21.71 ± 7.76 per nest across control nests with blow flies; minimum = 1; maximum = 54). Note that feather mites and lice were likely present in small numbers but were not quantified, although we did note several random observations of lice on nestlings. Therefore, any effects of permethrin on morphology and telomere length can only be attributed to blow fly reduction, although it is possible that removal of other ectoparasites also contributed. In addition, our estimates do not distinguish sibling variation in ectoparasite exposure and therefore, only estimates a nest‐average exposure for our nestlings of interest.

Tree swallows are obligate secondary cavity nesters that host a wide variety of ectoparasites, including mites, lice, fleas, and flies (Figure 1). These ectoparasites feed on blood, skin, and feathers (Janovy et al., 1997 ; Rendell & Verbeek, 1996 ) and can negatively impact offspring physiology, immune function, and survival (López‐Arrabé et al., 2015 (link); Martínez‐de La Puente et al., 2011 (link); Merino & Potti, 1995 ; Saino et al., 1998 ). Blow flies (Protocalliphora) have been found to infest 65.9% of tree swallow nests in northeastern US (Roby et al., 1992 ), with loads ranging from 4 to 54 parasites per nest (Grab et al., 2019 (link)). Previous work shows that blow flies feed on nestling blood and can cause anemia, hyperglycemia, and increased metabolic rates in avian hosts (De Simone et al., 2018 ; Grab et al., 2019 (link); Pryor & Casto, 2015 (link); Sun et al., 2020 (link)). One broad‐spectrum insecticide commonly used to remove ectoparasites is permethrin, which attacks the nervous system of larval and adult insects (Edwards, 2006 ). Permethrin treatment is effective against blow flies in nests of tree swallows (De Simone et al., 2018 ; Grab et al., 2019 (link)) and other bird species (Bulgarella et al., 2020 ) and also decreases the abundance of other ectoparasites like fleas and mites (Harriman et al., 2014 ; Pap et al., 2005 ; Pryor & Casto, 2017 (link)).
Any nest with a known hatching day was selected for our experiment, and then randomly assigned to one of two treatment groups, one in which ectoparasites were eliminated via the application of the insecticide permethrin (Permectrin II ©, diluted to 1% with distilled water), the other in which nests were treated with water as a control. In both treatment groups, nests were sprayed on day 0 and again on day 4. To do this, nestlings were temporarily removed from the nest, the bottom and sides of the nest were sprayed thoroughly (to minimize direct contact with nestlings), and the nestlings were returned once the nest had completely dried approximately 5 min later. To the extent possible, permethrin‐ and water‐treated nests were paired by hatch date to avoid the confounding effects that date has on many aspects of tree swallow reproduction (Winkler et al., 2020 ). The final number of nests in our study (n = 16 control, n = 16 insecticide) was less than the initial number sprayed due to brood loss.

Six CD1GM1 and eight CD1GM4 (MU Mutant Mouse Resource and Research Center (MMRRC), Columbia, MO, USA) mice were obtained at 9 weeks of age and set up in same GM profile breeding pairs. These mice were produced by existing colonies of mice generated several years ago and maintained as genetically similar outbred colonies of mice, differing in that each colony harbors a distinct supplier-origin microbiome, with GM1 originating from C57BL/6J mice (Jackson Laboratory, Bar Harbor, ME, USA) and GM4 originating from C57BL/6NHsd mice (Envigo, Indianapolis, IN, USA). In summary, these colonies were generated via the embryo transfer of CD-1 germplasm into pseudopregnant C57BL/6J and C57BL/6NHsd surrogate dams. Pups born to those dams acquired their respective supplier-origin microbiomes and served as founders for outbred colonies that have been maintained at the MU MMRRC since their initial description [22 (link),24 (link)]. Outbred status was maintained via careful rotational breeding and the annual introduction of new genetic stock from the CD-1 vendor via the embryo transfer of a newly acquired CD-1 germplasm into surrogate dams from the existing colonies. Additionally, the microbiome of these colonies was monitored on a quarterly basis via random sampling of 10 cages of adult breeding trios per colony and 16S rRNA amplicon sequencing of fecal DNA. Three litters per microbiome were culled down to eight mice at birth (with a goal of four male and four female pups) to reduce the possible effects of litter size and differential maternal care on preweaning growth. Due to difficulties in accurately sexing neonatal pups, a total of 20 GM1 offspring (10 cages, 5 male, and 5 female) and 22 GM4 offspring (11 cages, 5 male, and 6 female) were available for weaning into same-sex pairs at three weeks of age. Coprophagy leads to a shared cage-level microbiome and the single housing of mice is nonstandard husbandry that is considered stressful to mice. For these reasons, all adult experimental outcomes were based on the cage (i.e., mouse pair) as the experimental unit in an effort to eliminate cage effects and allow for normal activity and feeding behaviors. All mice were housed under barrier conditions in microisolator cages on ventilated racks with pelleted paper bedding and nestlets as enrichment. All mice had ad libitum access to an irradiated diet (LabDiet 5058 for breeder animals and LabDiet 5053 for feed study animals, LabDiet, St. Louis, MO, USA) and autoclaved tap water under a 12:12 light/dark cycle. The mice were determined to be free from opportunistic bacterial pathogens including Bordetella bronchiseptica; cilia-associated respiratory (CAR) bacillus; Citrobacter rodentium; Clostridium piliforme; Corynebacterium bovis; Corynebacterium kutscheri; Helicobacter spp.; Mycoplasma spp.; Pasteurella pneumotropica; Pneumocystis carinii; Salmonella spp.; Streptobacillus moniliformis; Streptococcus pneumoniae; adventitious viruses including H1, Hantaan, KRV, LCMV, MAD1, MNV, PVM, RCV/SDAV, REO3, RMV, RPV, RTV, and Sendai viruses; intestinal protozoa including Spironucleus muris, Giardia muris, Entamoeba muris, trichomonads, and other large intestinal flagellates and amoebae; intestinal parasites including pinworms and tapeworms; and external parasites including all species of lice and mites via quarterly sentinel testing performed by IDEXX BioAnalytics (Columbia, MO, USA).