Many environments present some degree of seasonal water limitations; organisms that live in such environments must be adapted to survive periods without permanent water access. Often this involves the ability to tolerate dehydration, which can have adverse physiological effects and is typically considered a physiological stressor. While having many functions, the hormone corticosterone (CORT) is often released in response to stressors, yet increasing plasma CORT while dehydrated could be considered maladaptive, especially for species that experience predictable bouts of dehydration and have related coping mechanisms. Elevating CORT could reduce immunocompetence and have other negative physiological effects. Thus, such species likely have CORT and immune responses adapted to experiencing seasonal droughts. We evaluated how dehydration affects CORT and immune function in eight squamate species that naturally experience varied water limitation.

We tested whether hydric state affected plasma CORT concentrations and aspects of immunocompetence (lysis, agglutination, bacterial killing ability, and white blood cell counts) differently among species based on how seasonally water limited they are and whether this is constrained by phylogeny. The species represented four familial pairs, with one species of each pair inhabiting environments with frequent access to water and one naturally experiencing extended periods (>30 days) with no access to standing water.

The effects of dehydration on CORT and immunity varied among species. Increases in CORT were generally not associated with reduced immunocompetence, indicating CORT and immunity might be decoupled in some species. Interspecies variations in responses to dehydration were more clearly grouped by phylogeny than habitat type.

To test the effects of hydration state on immune function and CORT response, we collected blood samples from serially hydrated, dehydrated, and, in some cases, rehydrated captive individuals of the following squamate species: Gila monster (Heloderma suspectum; n = 8), Mexican beaded lizard (Heloderma horridum; n = 7), Children’s python (Antaresia childreni; n = 8), ball python (Python regius; n = 8), Sonoran gopher snake (Pituophis catenifer affinis; n = 8), diamond-backed water snake (Nerodia rhombifer; n = 8), western diamond-backed rattlesnake (Crotalus atrox; n = 8), and northern cottonmouth (Agkistrodon piscivorus; n = 8). These species represent confamilial pairs (within Helodermatidae, Pythonidae, Colubridae, and Viperidae) that inhabit environments with varied seasonal water limitations (Table 1). While the species within each familial pair have varied degrees of relatedness, each is substantially more related to the confamilial species than to the other species; in fact, the range in divergence between species may provide a broader and more useful view of the effects in question, as we are not limiting our focus to species that diverged evolutionarily for any set duration. To validate differences in their adaptations to water limitation, we also measured the rate of cutaneous evaporative water loss for each species, as cutaneous evaporation can be influenced by native habitat aridity (e.g., in snakes: Lahav and Dmi’el, 1996; Dmi’el, 1998). Snout-to-vent length (SVL) was only measured in animals which were already being anesthetized, during which an accurate reading could be obtained (Gila monsters and beaded lizards), but all animals were of adult size though the beaded lizards, were on the smaller side of adult range (Beck, 2009) and of a size similar to that of the adult Gila monsters. None of the subjects were siblings­. In studies with non-model vertebrates, it can be difficult to access large sample sizes. We found significant effects previously, using 8 subjects (in Gila monsters) and wanted to minimize numbers for animal safety, thus we had a maximum of 8 animals per species. All research was conducted with approval from the ASU Institutional Animal Care and Use Committee (protocol 14-1044R).

Helodermatidae: Gila Monster and Mexican Beaded Lizard

Gila monsters are large lizards that inhabit desert scrub habitats in southwestern North America. In the Sonoran Desert, which encompasses most of the species’ range, Gila monsters experience an extended hot, dry season prior to the monsoon season in late summer. During the dry season, Gila monsters experience a 60 to 80-day drought, during which there typically is no rainfall or standing water (Beck, 2009). Mexican beaded lizards are large-bodied lizards found in Mexico from Sonora to Oaxaca, mainly in tropical deciduous forests. Overall, these areas experience more rainfall and are more humid than those where Gila monsters live. While beaded lizards experience periods with little rain that can last at least four months (Murphy and Lugo, 1986), unlike the habitat of Gila monsters, standing water is usually available during this dry season (Beck, 2009). We used eight adult Gila monsters (SVL 274 – 338 mm; mass 302 – 556 g) received from the Arizona Game and Fish Department (AZGFD; permits #SP577864, SP684760) and seven captive-bred small adult Mexican beaded lizards (SVL 347 – 352 mm; mass: 513 – 719 g) of a slightly larger size compared to the Gila monsters. Both species were held under an AZGFD Holding Permit (#SP598954; #SP666234).

Pythonidae: Children’s Python and Ball Python

Children’s pythons are small-bodied pythons that live in the wet-dry tropical rainforests of northern Australia. While their habitat receives extensive rainfall during the wet season, it also experiences a dry season that lasts for several months between May and October, during which these snakes have elevated plasma osmolality (Brusch and DeNardo, 2017). Ball pythons are heavier bodied, medium-sized pythons that live in tropical grasslands and rainforests of sub-Saharan Africa. These areas have a cool, dry season, when rain is infrequent (from November to May), but there are no extended dry periods, thus ensuring sufficient water availability throughout the year. We used eight captive-bred adult Children’s pythons (mass: 362 – 433 g) and eight captive-born adult ball pythons (mass: 732 – 1058 g) in this experiment.

Colubridae: Sonoran Gopher Snake and Diamond-backed Water Snake

Sonoran gopher snakes are medium-sized snakes that inhabit the southwestern United States and northern states of Mexico, especially in the Sonoran Desert. Most of their range experiences extended hot, dry periods with no rainfall (Stebbins and McGinnis, 2018). Diamond-backed water snakes are small to medium-sized snakes that are restricted to marshes, ponds, or other habitats along bodies of water in the middle southern United States. They primarily eat fish and amphibians, and thus rely heavily on constant access to water (Powell, Conant and Collins, 2016). We used eight wild-caught gopher snakes (mass: 223 – 454 g) from the Sonoran Desert in southern Arizona (AZGFD permit #SP609640) and eight wild-caught diamond-backed water snakes (mass: 213 – 251 g) obtained by Stephen Secor in Itta Bena, MS (Mississippi Department of Wildlife, Fisheries, and Parks collecting permit #1127121).

Viperidae: Western Diamond-backed Rattlesnake and Northern Cottonmouth

Western diamond-backed rattlesnakes are medium to large venomous snakes found in the southwestern United States and central and northern Mexico. They are widely distributed across a variety of habitat types, including the Mojave, Sonoran, and Chihuahuan deserts, where they withstand extended hot, dry seasons with no access to water (Stebbins and McGinnis, 2018). Cottonmouths are medium to large venomous snakes that live in the southeastern United States. They are found in marshes, swamps, and other slow-moving water habitats. Cottonmouths eat fish and amphibians, but they also eat more terrestrial prey, including mammals and birds. These snakes depend on ready access to water year-round (Powell, Conant and Collins, 2016). We used eight wild-caught western diamond-backed rattlesnakes (mass: 292 – 479 g) obtained as nuisance animals near Phoenix, AZ, and eight wild-caught northern cottonmouths (mass: 246 – 308 g) obtained from Steven J. Beaupre in Arkansas. All venomous species were held under AZGFD permits #SP598954 and #SP666234.

Cutaneous Evaporative Water Loss Measurements

We measured cutaneous evaporative water loss (CEWL) rates under standard room conditions (25.8 ± 0.1 °C, 23.3 ± 0.1 % relative humidity) for all subjects. To measure CEWL, we used an AquaFlux AF200 (Biox Systems Ltd., London, England, UK), which derives water vapor flux from humidity gradient measurements using a condenser within a closed chamber. Using AquaFlux version 6.2 software, we calibrated the unit prior to each use and recorded CEWL rate (g·m⁻²·h⁻¹) in real time. To maintain tight contact between the standard measurement cap (orifice diameter of 7 mm) and an animal’s dorsolateral skin at mid-body, we fitted the measurement cap with a donut-shaped piece of closed-cell foam with the hole in the center of the foam being of the same size as, and lined up with, the orifice of the measuring cap. We maintained contact between the unit and the animal’s skin until CEWL rates stabilized (± 0.02 g·m⁻²·h⁻¹ for 10 sec). We ran all measurements in duplicate to verify repeatability.

Hydration States Experiment

We used all individuals to serially examine the effects of hydration state on immune function and stress responses. For Gila monsters, Mexican beaded lizards, and both python species, we collected blood samples from all subjects in a hydrated state (290 – 305 mOsm kg^-1; HYD), with elevated but well-tolerated increased plasma osmolality, or “dehydrated” (315 – 360 mOsm kg^-1; DHY), and when rehydrated (265 – 300 mOsm kg^-1; RHY). In the other species, due to logistical limitations (e.g., difficulty of timed bleeding), we only collected two samples from each snake – one when hydrated (either initial or rehydrated) and the other when dehydrated, but the plasma osmolality range for each hydration state was the same in all species.

To begin the experiment, we fasted animals for two weeks to achieve a post-absorptive state. We then placed them in individual cages (75 X 35 X 13 cm; Freedom Breeder, Turlock, CA, USA) in an environmental chamber at 30 °C. We gave them two weeks to acclimate with water ad libitum. For the hydrated state assessments, we then collected an initial 0.7 ml blood sample from each individual from the caudal vein or by cardiocentesis using a heparinized 1 ml syringe. All initial samples were collected within three minutes of disturbance to avoid elevation of plasma CORT associated with handling (Romero and Reed, 2005). Blood samples were usually taken in the morning, between 0800 and 1100 hours. Following each animal’s initial sample, we administered a standardized 30-minute perturbation that commonly induces a CORT response in reptiles (Schuett et al., 2004; Romero and Reed, 2005; Langkilde and Shine, 2006). We induced stress by either restricting the animal’s movement and prodding it gently with a stick or by agitating the animal in a closed container. We administered the stressors for 10 out of every 30 seconds of the treatment period. Immediately after the perturbation (31 to 34 minutes after handling started), we collected a 0.1 ml blood sample to assess stress reactivity (i.e., the difference in CORT concentration between the pre- and post-stressor samples).

After we collected the hydrated set of samples, we immediately removed water from the animal’s cage and began the dehydration treatment for this experiment. For lizards, we also removed all fluid from the urinary bladder as needed, as some lizards, including Gila monsters, use the urinary bladder as a water reservoir (Davis and DeNardo, 2007). Removing urinary fluid thus increased the rate of dehydration in the lizards. While lizards were anesthetized (using 2% Isoflurane), we removed bladder fluid using transurethral catheterization according to Davis and DeNardo (2007). We then confirmed the lack of bladder fluid using ultrasonography (Concept/MLV, Dynamic Imaging, Livingston, UK). As snakes lack a urinary bladder, we only catheterized lizards.

Once animals were deprived of water, we checked them visually every day and weighed them every 1-7 days to estimate water loss and dehydration. Frequency of weighing depended on the species (e.g., semi-aquatic species dehydrate more quickly) and the time since water was last provided (i.e., frequency increased as animals dehydrated). To monitor plasma osmolality, we typically collected small blood samples (0.1 ml) after two weeks of dehydration and then once a week thereafter. The semi-aquatic species were an exception; we collected their first blood samples four days after water was removed and subsequent samples more frequently. When an animal approached an elevated but well-tolerated plasma osmolality, the animal was not disturbed for four days to reduce the chance of recent stress affecting CORT in initial samples. After four days without disturbance, we collected the dehydrated blood sample set (as above, in the hydrated state; 0.7 ml initial, 0.1 ml post-stressor), then gave the animal water.

After seven days of free water access, we collected the rehydrated blood sample set (as above; 0.7 ml initial and 0.1 ml post-perturbation) and removed the animal from the experiment. We immediately refrigerated samples until we could process, aliquot, and freeze them (within six hours of sampling) for later assays.

If the effort to collect a blood sample failed, we thoroughly evaluated that animal and then left it for four days of no disturbance before another attempt at sampling. When repeating the sample collection effort for the dehydrated sample set, we gave animals controlled amounts of water (roughly equivalent to 50% of the mass lost in the previous four days) to slightly rehydrate them, so they would remain safely dehydrated four days later. If three sampling attempts were unsuccessful, we skipped the sample and removed the animal from the Hydration States Experiment.

Dehydration Tolerance

Dehydration tolerance trials were only conducted in animals that had already completed the Hydration States Experiment (above). The Dehydration Tolerance Experiment was conducted last, as it explored critical levels of dehydration, and thus it was uncertain as to the recovery period and possible impact on other trials. Due to logistical limitations, we determined the extent of dehydration (i.e., plasma osmolality) that could be tolerated in only four of the eight species – gopher snakes (n = 5), diamond-backed water snakes (n = 7), western diamond-backed rattlesnakes (n = 6), and cottonmouths (n = 5). After completing the Hydration State Experiment, subjects were fed and then given at least 4 weeks to recover and reach a post-absorptive state before starting tolerance trials. Animals that had been difficult to bleed during the Hydration State Experiment were excluded from this part of the experiment, as obtaining blood samples at greater levels of dehydration would be even more difficult; with a limited number of animals, this reduced tolerance sample sizes. We measured the number of days until the snake showed signs of clinical dehydration, as well as change in plasma osmolality and body mass lost over that time. Animals in a post-absorptive state were held in individual cages in an environmental chamber at 30 °C with no food or water and were monitored daily for health and weekly (at most) for plasma osmolality (same as above, in Hydration States Experiment). When plasma osmolality reached 315 – 360 mOsm kg^-1; DHY), we increased health monitoring to two to four times daily until the animals showed the first symptoms of clinical dehydration (e.g., lethargy, loss of righting response, reduced responsiveness to stimulation (Divers, 1999)). We limited blood samples to 0.1 ml and collected samples every 4 to 14 days (as the trials went on), for a maximum of 10 times over the course of the trial. Blood samples were usually taken in the morning, between 0800 and 1100 hours. As soon as clinical signs appeared, we collected a final blood sample to test plasma osmolality, and we then provided the animal with access to water ad libitum. Two days after each animal was rehydrated, we removed the animal from testing altogether.

Sample Preparation and Assays

We assayed plasma from each initial sample (0.7 ml) to determine hydration state (i.e., plasma osmolality, mOsm kg^-1), initial plasma CORT concentration, and immune function (agglutination, lysis, bacterial killing ability (BKA), and white blood cell differential). We used post-perturbation plasma samples to determine reactive CORT concentrations.

After collecting the initial sample, we used two drops of whole blood to create blood smears for analysis of white blood cell counts (described below). We centrifuged the remaining blood from the initial samples as well as the blood from the post-perturbation sample and divided the resulting plasma into 50 ul aliquots. We stored separated blood cells and plasma at -80 °C until later analysis.

Sample Processing: Hydration State

We analyzed each initial sample for plasma osmolality using vapor pressure osmometry (± 6 mOsm kg^-1; model 5100C; Wescor, Inc., Logan, Utah, USA) as described in Davis and DeNardo (2007).

Sample Processing: Corticosterone Assays

We assayed plasma samples for CORT in duplicate using enzyme-linked immunoassay kits following the manufacturer’s instructions (ADI-900-097; Enzo Life Sciences, Farmingdale, NY, USA). To validate the assay for these eight squamate species, we conducted a pilot assay to determine that the curves generated by serially diluted (5X to 80X) pooled samples for each species were parallel with the standard curve (Supplementary Figure 1). Based on the pilot, we diluted the plasma samples from each species 40X with assay buffer containing the prepared steroid displacement reagent at a volume equal to 1% of plasma volume before assaying.

We used ten assay plates total, with a standard curve on each plate, and ran them over three days. We assayed all the samples from a species on the same day and randomly assigned samples to a plate on a given day. Each plasma sample was 6.25μl. The average inter- and intra-assay coefficients of variation were 7.1 % and 13.2 %, respectively. The average assay sensitivity was 6.34 pg ml^-1.

Sample Processing: Agglutination and Lysis Assays

We measured agglutination and hemolytic ability of all initial samples following Moeller et al.’s (2013) modified protocol, originally from Matson et al. (2005). We serially diluted 20 µl of each plasma sample from 1:2 to 1:2048 in a 96-well plate and added 20 µl of diluted 50% heparinized sheep blood (SBH050, HemoStat Laboratories, Dixon, CA, USA) to each well. After incubation at 29 °C (the active season mean diurnal body temperature of Gila monsters (Davis and DeNardo, 2010) and an active temperature for many of the species tested), we assayed for agglutination (marked by changes in the formation of a dense RBC pellet), incubated the plates at 29 °C again, then assayed for hemolysis activity (marked by a lack of red pigment pellet presence).

Sample Processing: Bacterial Killing Ability

Following French and Neuman-Lee (2012), we used initial plasma samples to conduct bacterial killing assays. We pipetted 2 µl of thawed plasma in duplicate onto 96-well round-bottom microplates. Negative-control wells held 6 µl phosphate-buffered saline and 18 µl CO₂-independent Medium plus 4 mM L-glutamine and no bacteria. Positive-control wells had 6 µl working bacteria solution, which held 10⁴ colony-forming units of Escherichia coli (ATCC NO. 8739), along with 18 µl medium. We also added 6 µl working bacteria solution and 16 µl of medium to samples, so all wells had a volume of 24 µl. We thoroughly mixed all wells, gently vortexed the microplates for 1 min, then incubated them at 37 °C (the optimal temperature for exponential E. coli growth) for 30 min.

After incubation, we vortexed the plates for another minute, then added 125 µl sterile tryptic soy broth (Sigma-Aldrich NO. T8907;15 g broth per 500 ml Nanopure water) to each well. After an additional minute of gentle vortexing, we read the absorbance of the plates at 300 nm (BioRad xMarkTM Microplate Absorbance Spectrophotometer). We incubated the plates at 37 °C for 12 hr, then vortexed for 1 min and read again. We compared sample absorbance before and after the 12 hr of bacterial growth to the positive controls (0 % bacterial killing). We calculated percent bacteria killed as one minus the mean absorbance for each sample, which we ran in duplicate, divided by the mean absorbance for the positive control (triplicate wells containing only media and bacteria), multiplied by 100.

Sample Processing: Blood Cell Profiles

We created and analyzed duplicate blood smears from each initial sample. After inverting each blood sample several times to remix any settled cells, we smeared one drop of blood on a slide and stored them in a desiccator until they were preserved with methanol and re-stored for up to 6 mo. We then stained with Giemsa-Wright stain for 60 min at the optimal dilution for each species to maximize visualization (1:15 dilution for helodermatids; 1:18 dilution for other species), then rinsed slides in two nanopure water baths for 5 min each. We counted lymphocytes, heterophils, monocytes, and basophils under a light microscope (BX60, Olympus Optical Co., Tokyo, Japan) at 400X magnification, counting the number of each blood cell type (according to Cooper-Bailey et al., 2011) along a mapped grid until the first 100 white blood cells were counted on a slide. Eosinophils and azurophils were not found in most samples so were not counted.