Ecoimmunological patterns and processes remain understudied in wild primates, in part because of the lack of noninvasive methods to measure immunity. Secretory immunoglobulin A (sIgA) is the most abundant antibody present at mammalian mucosal surfaces and provides an important first line of defense against pathogens. Recent studies show that sIgA can be measured noninvasively in feces and is a good marker of mucosal immunity. Here we validated a commercial ELISA kit to measure fecal IgA in baboons, tested the robustness of its results to variation in collection and storage conditions, and developed a cost‐effective in‐house ELISA for baboon fecal IgA. Using data from the custom ELISA, we assessed the relationship between fecal IgA concentrations and gastrointestinal parasite burden, and tested how sex, age, and reproductive effort predict fecal IgA in wild baboons. We find that IgA concentrations can be measured in baboon feces using an in‐house ELISA and are highly correlated to the values obtained with a commercial kit. Fecal IgA concentrations are stable when extracts are stored for up to 22 months at −20°C. Fecal IgA concentrations were negatively correlated with parasite egg counts (Trichuris trichiura), but not parasite richness. Fecal IgA did not vary between the sexes, but for males, concentrations were higher in adults versus adolescents. Lactating females had significantly lower fecal IgA than pregnant females, but neither pregnant nor lactating female concentrations differed significantly from cycling females. Males who engaged in more mate‐guarding exhibited similar IgA concentrations to those who engaged in little mate‐guarding. These patterns may reflect the low energetic costs of mucosal immunity, or the complex dependence of IgA excretion on individual condition. Adding a noninvasive measure of mucosal immunity will promote a better understanding of how ecology modulates possible tradeoffs between the immune system and other energetically costly processes in the wild.

To validate the commercial kit, develop and validate the in‐house assay, and to measure natural variation in IgA, we collected fecal samples from wild baboons opportunistically, immediately after defecation, from known individuals in Amboseli. Samples were homogenized, and an aliquot of each sample was placed in an empty tube, stored in a cooler with ice packs, brought to our camp site (within 8 hr), and stored in camp at −20°C. Samples were then sent to University of Nairobi where they were freeze dried and stored at −20°C until shipment to the United States. In this study, we analyzed fecal IgA concentrations of 310 fecal samples from 168 baboons collected in May–June 2013, May–Jul 2014, and May–June 2015. The samples collected in years 2013 and 2014 (206 samples) were analyzed using the commercial kit, and all samples from year 2013 to 2015 (310 samples) were analyzed using the in‐house ELISA. For a small subset of these samples (31 samples), we also collected an aliquot in a tube containing 10% of buffered formalin for parasite determination.

To explore whether field storage conditions affected measured IgA concentrations and to test inter‐assay variation of the commercial kit), we collected one fecal sample from each of four captive baboons held at the Division of Laboratory Animal Resources, Duke University Medical Center. Samples were collected immediately following defecation, after which each sample was homogenized and divided into six aliquots: one was collected in 95% ethanol and five were collected in empty tubes; all six aliquots were frozen at −20°C. The aliquots were then shipped on dry ice to Princeton University and stored at −20°C until processed as indicated in Goal 2 below to simulate processing of field samples in Kenya. Processed samples were freeze dried and sifted to a fecal powder (Gesquiere et al., 2008; Khan, Altmann, Isani, & Yu, 2002), before being extracted and assayed.