Senescence and the stress axis in male ground squirrels

A critical time in the life of a male occurs at reproduction when his behavior, physiology, and resources must be brought to bear for the central purpose of his life. We ask whether reproduction results in dysfunction of the stress axis, is linked to life history, and causes senescence. We assessed if deterioration in the axis underlies variation in reproductive lifespan in males of 5 species of North American ground squirrels whose life history varies from near semelparity to iteroparity. The most stressful and energy-demanding time occurs in spring during the intense 2–3 week breeding competition just after arousal from hibernation. We compared their stress axis functioning before and after the mating period using a hormonal challenge protocol. We found no evidence of stress axis dysfunction nor was there a relationship between reproductive lifespan and stress axis functional deterioration. Moreover, there was no consistent relationship between free cortisol levels and downstream measures. Thus, stress axis function was not traded off to promote reproduction and conclude that it is a pre-requisite for life. Hence, it functions as a constraint and does not undergo senescence.Statistical analysis All data were analyzed using SAS 9.2 (SAS Institute Inc., Cary, NC, USA). For simple comparisons among species, we used one-way ANOVAs and the conservative Tukey-Kramer multiple comparison post-hoc test. For our regressions of postbreeding GC levels against mean reproductive lifespan and probability of surviving more than one breeding season (PROC GLM), we did not test for or control for phylogenetic non-independence. In part, our sample size precluded using phylogenetic contrasts: tests for phylogenetic independence perform poorly on small datasets and if our species were not independent, our sample size would be reduced by one in order to do the contrast. Our decision is further justified by the fact that we selected very closely related species, and because previous comparative studies of GC levels have found that much more diverse species groupings are nonetheless phylogenetically independent. For the prebreeding and postbreeding comparisons, data were first examined for normality. Where significantly non-normal data could not be normalized with a transformation, we used the Wilcoxon-Mann-Whitney non-parametric two-sample test using the EXACT option to generate a Monte Carlo-based exact P-value. Because we released prebreeding Franklin's ground squirrels and some of our postbreeding samples came from recaptured individuals, we analyzed differences between pre- and postbreeding periods using a mixed model in SAS PROC MIXED, with subjects as a repeated measure. All other species comparisons used PROC TTEST; where pre- and postbreeding data had non-homogeneous variances based on the folded-F test, we use Satterthwaite-adjusted degrees of freedom for the t-tests (which can result in degrees of freedom that are not whole numbers). Unless otherwise specified, we present means with 95% confidence intervals.Funding provided by: Natural Sciences and Engineering Research Council of CanadaCrossref Funder Registry ID: http://dx.doi.org/10.13039/501100000038Award Number: RGPIN 6169Study areas and field procedures We trapped prebreeding (within a few days of first emerging from hibernation) and postbreeding males (2–3 weeks later, same site). For the 3 western species (AGS, CGS, and RGS) logistics (i.e. ~2500 km between the AGS and RGS sites), asynchronous timing of emergence and reproduction, and a narrow window of reproduction within a species permitted us to trap only them in 2007 and the 2 eastern species (FGS, TLGS) in 2008. In 2007, we trapped RGS at Kinsella, Alberta (53°N 111°33'W) on March 25–27 and April 15–16, AGS at the Pelly River Ranch, Yukon (62°50'N 137°18'W) on April 9–12 and April 27–30, and CGS at Kananaskis, Alberta (51°2'N 115°2'W) on April 22–24 and May 7–9, respectively. In 2008, we trapped TLGS on April 15–19 and May 5–11 and FGS on May 6–11 and May 28–30 near Portage La Prairie, Manitoba (TLGS at 50°7'N 98°23'W, FGS at 50°12'N 98°14'W). Animals were trapped by placing burrow traps or cage traps (Tomahawk #102, Tomahawk Live Trap Company, Tomahawk, WI, USA) in or next to burrows that animals were seen to have entered. Traps were monitored at least every 30 minutes and captured males were then kept in cage traps covered with a pillowcase in a shaded, quiet location until trapping was completed. We started trapping soon after animals emerged (07h00–09h00) and continued trapping for at most 4 hours or until we had as many animals as we could process at one time (6–9 animals). Field Hormonal and Sampling Procedures Trapped animals were brought to a field laboratory, placed in a cool, quiet location, covered with pillowcases, and left for at least 1 hour to habituate before initiating the hormonal challenge procedure. One animal at a time was removed from its trap, weighed, and anesthetized with isoflurane (IsoFlo, Abbott Laboratories, Saint Laurent, QC, Canada). We took an initial 0.6 mL blood sample (BASE) and injected 0.1 or 0.4 mg/kg dexamethasone (DEX) (Sabex, Montreal, Canada), then returned the animal to the covered trap and processed the next animal. From the BASE sample, we obtained 4 subsamples: 0.3 ml for a complete blood count (Vita-tech Veterinary Laboratory Services, Markham, ON, Canada); 75 µl microhematocrit tubes for hematocrit (packed red blood cell volume, %); a drop for glucose levels (mg/dL, FreeStyle glucose meter, Abbott Laboratories, Alameda, CA, USA); and the rest for hormone and free fatty acid measurements. The latter sample was centrifuged at 8800g for 5 min, and the plasma from the hematocrit tubes was pooled with it and frozen at -20°C. The DEX tests the negative feedback to the pituitary and inhibits ACTH release. As we found no difference in DEX response to the different doses in 2007 (unpublished data), we administered 0.1 mg/kg to all animals in 2008. Two hours after the DEX injection, we anesthetized the animal again and took a 0.2-0.3 mL blood sample ("DEX" bleed) and then injected 4 IU/kg of adrenocorticotropic hormone (ACTH; Synacthen Depot, Novartis Pharmaceuticals Canada Inc, Dorval, Quebec) into the thigh muscle to directly stimulate adrenal GC production. Post-ACTH blood samples of 0.2 mL each were taken at 30, 60 and 120 minutes ("P30", "P60" and "P120" bleeds). Glucose was measured at each sampling point. Blood samples were centrifuged and plasma collected and frozen at -20°C and, on return to the university, stored at -80°C. After the P120 bleed, the animal was euthanized by anesthetic overdose and decapitated. On necropsy, we dissected out and weighed both adrenals (nearest mg), abdominal fat (fat that was easily removed from the abdominal cavity and did not include fat in the mesentery: nearest 0.1g), both testes (nearest 0.01g), and eye lens. Adrenal mass was a rough indicator of adrenal capacity based on the expectation that adrenal hypertrophy would occur from the prebreeding to the postbreeding period reflecting greater capacity to produce GCs. Abdominal fat was a measure of a key energy reserve and expected to decline markedly. Testes mass was a measure of the capacity to produce sperm and testosterone and expected to decline markedly from the prebreeding to the postbreeding period. Age may be a key determinant in how individuals respond to stress and we attempted to age our squirrels by measuring their eye lens weights and by sectioning their femurs and jaws. To quantify the intensity of inter-male aggression during the breeding season, we examined the skin externally and internally for evidence of wounding. The degree of wounding was scored according to the number of punctures (bite marks that penetrated the skin, usually only visible from the inside of the hide) and externally visible wounds (tears in the skin that exposed the muscle below or had scabbed over). Our scoring categories were: "None" for no wounds, "Minimal" for 2 cm2 but