Abstract
Environmental changes result in physiological responses of organisms, which can adversely affect population dynamics and reduce resistance to disease. These changes are expressed in chronic levels of stress. The measurement of glucocorticoid (GC) concentrations in faeces is a non-invasive method for monitoring stress in wildlife. The metabolism and excretion of steroids differ significantly between species and, as a consequence, non-invasive methods must be physiologically validated for each species. Koalas (Phascolarctos cinereus) are declining in numbers through much of their range. The role of chronic stress in koala populations has not been identified. Prior to the assessment of faecal GC concentrations in wild koala populations, the excretion timing and concentrations of GCs need to be determined. In this study, we assessed a method for identifying and measuring the concentrations of GC metabolites in faecal pellets of captive koalas following ACTH treatment. The results show that an elevation of plasma cortisol concentrations, using sustained release of ACTH, results in elevated concentrations of faecal cortisol/cortisol metabolites. Taking into account the excretion time lag, an increase in faecal cortisol metabolite concentrations corresponds to the release of GCs from the adrenal cortex as early as 36 h before faecal pellet collection. The calculations of steroid partitioning of plasma cortisol showed that the ACTH-stimulated values were significantly different from the control values for the concentrations of free, corticosteroid-binding globulin-bound and albumin-bound cortisol. This study validates the use of faecal cortisol analysis to assess the activity of the hypothalamo–pituitary–adrenocortical axis in freshly collected koala faecal pellets and indicates that the method should be suitable to assess the adrenocortical status of koalas in wild populations.
Introduction
Understanding the physiological responses of organisms to changes in their environment is of great importance to conservation biologists as stress can alter behaviour, reduce resistance to disease and adversely affect population performance (including impacting survival and reproduction; Munck et al. 1984, Wingfield & Farner 1993, Millspaugh & Washburn 2004). Vertebrates cope with unfavourable conditions by initiating a stress response that is generally characterised by the release of glucocorticoid (GC) hormones (mainly cortisol or corticosterone in mammals) by the adrenal cortex (Seyle 1935, Axelrod & Reisine 1984, Mostl & Palme 2002, Stewart 2003). Short-term GC secretions, resulting from acute stress (e.g. predator attack), are beneficial, as they assist an animal in coping with a stressor by mobilising energy reserves during the stressful situation (Lee & Cockburn 1985, Wingfield et al. 1998, Sapolsky 2002, Mateo 2006). However, if the stress persists over a long period (i.e. chronic stress), the sustained high levels of GCs can lead to deleterious effects on the physiology of the animal and greater susceptibility to disease and reduced fecundity and survivorship (Munck et al. 1984, Sapolsky et al. 2000, Mostl & Palme 2002, Stewart 2003, Lane 2006). The release of GCs is routinely used as a measure of the stress response in ecological and conservation studies to assess the health of animals (Romero 2004). The measurement of GC concentrations can reveal how stressors affect the survival and reproductive success of wild animals and how natural environmental challenges (e.g. predators, conspecifics and weather), climate change and habitat disturbance have an impact on wild populations (Boonstra & Singleton 1993, Wasser et al. 1997, Creel et al. 2002, Thiel et al. 2008, Sheriff et al. 2009).
In plasma, cortisol exists primarily in three states: unbound, bound to albumin and bound to corticosteroid-binding globulin (CBG; Smith et al. 1980). CBG binds to cortisol with a high affinity and facilitates transport in the blood, while cortisol binds to albumin with a lower binding affinity (Hiramatsu 1983, Chu & Ekins 1988). Under normal conditions, ∼70–85% of the secreted cortisol is bound to CBG (Hiramatsu 1983) and 10–15% to albumin (Chu & Ekins 1988), and the remaining 5–10% of cortisol is unbound and free to diffuse across cell membranes and bind to intracellular GC and mineralocorticoid receptors (Mendel 1989). Steroid hormones bound to carrier proteins (CBG and albumin) are considered to be biologically inactive. They provide a reservoir of inactive circulating hormones and regulate the amount of free hormones available for diffusion into tissues (Mendel 1989). The unbound cortisol reflects the biologically active concentration of the hormone that is capable of interacting with intracellular steroid receptors (Willcox et al. 1985). It is important to investigate the partitioning of steroids because changes in the plasma concentration of biologically active GCs are the foundation of physiological action, and when in excess (e.g. chronic stress), it can cause deleterious effects on the health of an animal.
Traditionally, GC concentrations have been measured in blood plasma. However, this procedure is invasive and blood GC levels are ‘point’ samples that are highly variable and are strongly affected by the time of day, food intake and any stressful situation or environmental disturbance that may have occurred shortly before sampling (including capture and handling techniques; Sapolsky 1982, Wingfield et al. 1992, Beerda et al. 1996). Therefore, specific point measurements may be unrepresentative of long-term hormone exposure levels (Harper & Austad 2000, Davenport et al. 2006). Furthermore, blood sampling can be difficult to carry out under field conditions. More recently, non-invasive methods for measuring GC metabolite concentrations in faeces and hair have become widely accepted tools for gaining important information about an animal's endocrine status (Mostl & Palme 2002, Young et al. 2004, Palme et al. 2005, Davenport et al. 2006, Bosson et al. 2009, Sheriff et al. 2009). The employment of non-invasive techniques to assess the physiological status of wild populations can be advantageous for both practical and ethical reasons (Lane 2006).
Faecal GC measures provide an integrated measure of fluctuating blood concentrations during the time the product was being formed and stored (usually a good indicator of GC levels of the previous 24–48 h), thus providing a more representative measure of an individual's average hormone secretion (Goymann et al. 1999, Harper & Austad 2000, 2001). Furthermore, faecal GC measures can be obtained easily without handling or observing the animals (Mostl et al. 1999, Palme et al. 1999, Harper & Austad 2000, Mostl & Palme 2002, Millspaugh & Washburn 2003). The metabolism and excretion of steroids differ significantly between species and sexes (Touma et al. 2003, Palme 2005, Palme et al. 2005, Touma & Palme 2005); therefore, non-invasive methods must be physiologically validated for each species (Rehnus et al. 2009). This validation is typically carried out using an adrenocorticotropic hormone (ACTH) challenge, which involves pharmacologically inducing physiological changes in circulating GC levels to evaluate how these changes are reflected in the concentrations of faecal GC metabolites (Touma & Palme 2005). Although a large proportion of the conjugated GC (cortisol in marsupials) is known to be excreted in urine, in this study, we used faecal and plasma samples to assess the short-term elevation of cortisol levels in the blood plasma and the long-term delayed elevation of cortisol metabolite levels in faeces.
The koala (Phascolarctos cinereus) is a widely distributed, arboreal marsupial folivore, with its range extending across 30 bioregions from tropical Queensland to temperate Victoria and South Australia. Koalas are declining in numbers through much of their range. The drivers of these declines include habitat loss and fragmentation, thermal stress, fodder quality reduction, and water availability and disease. To appropriately quantify the impact of environmental and physiological stressors on koala populations, the validation of non-invasive techniques and assays is a necessary first step.
The aims of this study were to examine the adrenocortical status of wild koalas held captive under controlled conditions and to compare this with the adrenocortical status following a 24-h ACTH challenge. This was achieved by measuring cortisol concentrations in plasma and faeces as well as by employing glucuronidase cleavage of polar bonds to liberate unconjugated cortisol (Mostl & Palme 2002). The establishment of the temporal relationship between ACTH-stimulated GC concentration elevation in plasma and that in faeces would validate the use of fresh faecal pellets from wild koalas to enable an estimate of their adrenocortical status to be made. We did not collect urine sample since this would be impractical for any studies on koalas in the wild.
Materials and methods
Animals
The research was conducted at the Australia Zoo Wildlife Hospital (Beerwah, Queensland). The koalas examined in this study were wild animals being rehabilitated as part of the hospital's standard procedures for koalas prior to release. The koalas were at the later stages of rehabilitation when any injuries had healed and they no longer appeared to be disturbed by the presence of veterinary and nursing staff. The use of animals in an unstressed condition eliminated any influence of past trauma or impacts of treatment on GC concentrations.
The five koalas (four males and one female) used in this study were aged between 2 and 12 years. The daily diet of these koalas was fresh leaves of various Eucalyptus species and was the same for each koala. The study was conducted between March and May 2012. Each koala was held in a large wire mesh cage (∼3×3×3 m) and provided with 2–3 solid tree trunks and connecting horizontal poles. One to two experienced animal handlers lightly restrained each koala, enabling ACTH or saline (control) injections or blood sample collection for this study.
ACTH challenge experiment
Each experiment was carried out over a 7-day period, and each koala was subjected to both a control treatment (injection of saline) and an ACTH challenge treatment (ACTH in saline), with repeat treatments separated by 1 month to allow for the recovery of physiological parameters.
For the control treatment, the koalas were injected intramuscularly with 0.5 ml of saline on the morning of the first day (0 h (T0)). Faeces (one pellet from pellets voided during the collection period) and a blood sample taken by venepuncture of the cephalic vein were collected at 0, 12, 24 and 36 h. Faecal pellets were also collected daily (in the morning) from day 0 to 7 of the experiment. The samples were stored at −20 °C until analysis. The weights and body condition scores of the koalas were also recorded.
For the ACTH challenge, the koalas were treated in the same way, but injected intramuscularly with 25 IU 0.25 ml of depot ACTH (Novartis Pharmaceuticals Australia Pty Ltd.) in 0.5 ml of 0.9% saline at 0 h to increase circulating GC concentrations and again at 12 h. The Synacthen Depot is supplied as a suspension in which the active hormone is adsorbed onto an inorganic zinc complex that enables protracted release. This was used to elevate plasma GC concentrations for a 24–36-h period to allow for the appearance of GC metabolites in the faecal samples. In all the studies on marsupials in which it has been used, the long-acting synthetic β-1–24 amino acid form of ACTH has been shown to be potent in stimulating cortisol secretion from the adrenal cortex.
Blood samples were collected in lithium heparin vials, and plasma was immediately separated by centrifugation for 5 min at 1500 g. Faecal pellets and plasma samples were stored at −20 °C until analysis.
Faeces: extraction and analysis
Faecal pellets from individual koalas were oven-dried at 70 °C for 10 h. The dried samples were then crushed between two 10 mm PVC plates, passed through a fine sieve (1 mm) to remove leaf particles and then collected in 5 ml flat-bottomed polypropylene screw-capped vials. For each sample, 200 mg (±1 mg accuracy) of a dry faecal powder were weighed into 16×100 glass test tubes. Subsequently, 2 ml of borate buffer (pH 6.5; 0.1 M) were added to the dry powder and vortex-mixed, and then 50 μl of β-glucuronidase (β-d-glucuronoside glucuronosohydrolase, EC 3.2.1.31, Sigma Chemical Co.) containing ∼4000 units were added to each test tube. β-Glucuronidase is derived from Helix pomatia and is active in cleaving both glucuronidase and sulphatase. The test tubes were then incubated for 4 h at 37 °C on an orbital mixer. Then, 3 ml of redistilled diethyl ether were added to each test tube and vortex-mixed for 2 min, after which the test tubes were allowed to stand for 2 min. The lower aqueous phase was frozen in liquid nitrogen and the supernatant ether was decanted into 12×75 mm glass test tubes and evaporated to dryness at 40 °C in a hot block evaporator in a fume hood. The residue containing the extracted steroid was re-dissolved in 200 μl of a diluted zero cortisol calibration solution (Demeditec Saliva Free Cortisol Kit; diluted 1:10) and placed on an orbital mixer at 37 °C for 60 min, followed by short, high-speed vortex (20 s). Then, 100 μl of test samples, standards and controls were pipetted into wells of a 96-well Saliva Free Cortisol Kit (Demeditec DES6611, Demeditec Diagnostics GmbH, Kiel, Germany) and run as per kit directions. The efficiency of the extraction process was progressively tracked by addition of 30 000 d.p.m. 3H-cortisol (1,2,6,7 3H-cortisol 160 Ci/mmol, PerkinElmer, Melbourne, Victoria, Australia), and the final assay concentration for cortisol was corrected for this efficiency.
The specificity of the assay was as follows: 100%, cortisol; 0.38%, corticosterone; 1.85%, cortisone; 0.88%, 11-deoxycortisol and 9.89%, prednisolone; with all other steroids of a similar structure having specificity <0.20%. Inter-assay variation was 5.4% coefficient of variation (CV), while intra-assay variation was 5.5% CV. The analytical sensitivity of the assay was 0.014 ng/ml or 4.2 pg per loaded sample.
Serial dilutions of glucuronidase-treated koala faecal extracts run against the Demeditec assay kit calibrator standards gave a satisfactory degree of parallelism for the assay.
Assay data were analysed employing a four-parameter logistic fit using MyAssays Analysis Software Solutions (www.myassays.com).
To assess the effect of enzymatic cleavage of glucuronide and sulphate polar bonds using β-glucuronidase, the pellets collected 36 h after ACTH injection were pooled from four koalas and cortisol was extracted as described above. From the common pool of dry faecal powders, 16 extracts were prepared. Eight samples were extracted without enzymatic treatment and eight after enzymatic treatment.
Plasma: extraction and cortisol RIA
A commercially available cortisol antiserum (product number: C8409-100TST, Sigma Chemical Co.) was diluted with buffer G (0.05 M phosphate buffer containing 0.1% porcine gelatin and 0.1% sodium azide, pH 7.4) to result in ∼50% of the total 3H-cortisol being bound in the absence of competing unlabelled steroid. The optimal titre was determined to be 1/30 000.
Plasma samples of 10 μl were combined with 100 μl of phosphate buffer (0.05 M, pH 7.4) in 16×100 mm glass test tubes and the steroid was extracted by vortex mixing for 2 min using 2 ml of redistilled analytical reagent-grade AR dichloromethane. The test tubes were left standing for 2 min to allow separation of the aqueous and organic phases. The aqueous phase was then frozen to the walls of the test tubes by dipping the test tubes into liquid nitrogen for 10 s, and the dichloromethane lower phase was decanted into 12×75 mm glass test tubes and dried in a hot block evaporator at 45 °C in a fume hood. The dried residue was re-dissolved in 100 μl of phosphate buffer (0.05 M phosphate buffer containing 0.1% porcine gelatin and 0.1% sodium azide, pH 7.4).
Extraction efficiency was tracked using 30 000 d.p.m. 3H-cortisol added to the initial extraction tubes. Extraction efficiency was determined to be 90%.
From each sample, 25 μl were transferred into 12×75 mm polypropylene test tubes. To all the test tubes, 25 μl of 3H-cortisol (30 000 d.p.m./25 μl of 0.05 M phosphate buffer containing 0.1% porcine gelatin and 0.1% sodium azide, pH 7.4) were added. To the test tubes containing calibration standards, only 25 μl of buffer were added. Then, 50 μl of steroid-specific antibody were added to all the test tubes, which were then vortex-mixed for 2 min. The samples were then incubated at 0 °C using an ice-water slurry for 4 h. Into each test tube, 500 μl of dextran-coated charcoal (500 mg activated charcoal (fines removed) (Sigma), 50 mg Dextran T70 (Pharmacia), 400 ml phosphate buffer (0.05 M, pH 7.4) and 20 mg Thimerosal (Sigma Chemical Co.)) were pipetted. The test tubes were incubated at 4 °C for 15 min and were then centrifuged at 4 °C in a Beckman GPR refrigerated centrifuge for 10 min at 1500 g. Then, 100 μl of the supernatant containing the antibody-bound steroid were added to 1 ml of Ultima Gold scintillation fluid (PerkinElmer) in a scintillation vial, vortex-mixed and counted using a liquid scintillation spectrometer (Beckman LS 6000 TA, Beckman Coulter, Lane Cove, NSW, Australia). The concentrations of unknown samples were determined using the AssayZap Software (version 2.32, AssayZap, Cambridge, UK).
Plasma cortisol partitioning
Plasma steroid partitioning was determined using methods adapted from the works of Tait & Burstein (1964) and Paterson & Hills (1967) for the determination of the CBG-bound, albumin-bound and free compartments within the plasma. Microequilibrium dialysis has been used to describe the high-affinity and low-affinity binding of cortisol to plasma proteins (Bradley et al. 1980, Bradley 1990). The high-affinity binding constant employed (Ka) was 3.5×107 M−1 and the low-affinity binding constant used was 0.275 for a 1% (w/v) koala albumin solution. These constants were approximately in the middle of the range of published values for marsupials (Bradley et al. 1976, Sernia et al. 1979, Bradley 1982).
Statistical analysis
Baseline concentrations for the control and ACTH treatments were defined as the concentrations at 0 h (T0) of the first morning before injection of saline or depot ACTH. Where necessary, data were logarithmically transformed to satisfy the assumption of normal distribution and homogeneity of variances for the parametric tests. A two-way repeated-measures ANOVA was used to examine the effects of collection time and treatments (ACTH and control) for both plasma (including the partitioning of cortisol into plasma compartments (free, CBG bound and albumin bound)) and faecal samples (Lèche et al. 2011, Legagneux et al. 2011, Fanson et al. 2012). Treatment (ACTH and control) was the main factor and time was the repeated measure. Dunnett's multiple comparison tests were performed for post hoc analysis to test for differences between baseline (T0) and all other times (Fanson et al. 2012). The concentrations of saline-injected controls provided a relative baseline for comparisons with the ACTH treatment. t-Tests were used to examine whether the concentrations for ACTH treatment were higher than those for the control treatment for both plasma and faecal samples after injection (i.e. all times after T0).
Results
Plasma GC levels
There was a significant relationship between treatment (ACTH or control) and time of sample collection for plasma cortisol levels. The effect of the treatment varied with collection times (two-way repeated-measures ANOVA: P≤0.0001, F3, 21=12.93). Post hoc tests indicated that for the ACTH treatment, plasma cortisol levels measured after the challenge (12, 24 and 36 h) were significantly higher compared with the pre-injection levels (T0), while for the control treatment, there was no significant difference between the pre-injection and post-injection levels (Fig. 1).
Mean (±s.e.m.) plasma cortisol concentrations for the control and ACTH treatments at each collection time. Post hoc tests indicate a significant difference (**** signifies P<0.001) between 0 h and all other collection times for the ACTH treatment and no difference between 0 h and all other collection times for the control treatment.
Citation: Journal of Endocrinology 219, 1; 10.1530/JOE-13-0123
Faecal GC metabolite levels
There was a significant relationship between treatment (ACTH or control) and time of sample collection for faecal GC metabolite levels (two-way repeated-measures ANOVA: P=0.0161, F8, 56=2.630). Post hoc tests indicated that for the ACTH treatment, faecal cortisol levels that reached a peak 36 h after the challenge were significantly higher compared with the pre-injection levels (T0), while for the control treatment, there was no significant difference between the pre-injection and post-injection levels (Fig. 2). Furthermore, there was a significant difference between the ACTH and control treatments for all faecal cortisol concentrations measured after injection (i.e. after T0; t-test: P=0.0091, t=2.684, df=70; Fig. 2).
Mean (±s.e.m.) faecal cortisol concentrations for the control and ACTH treatments at each collection time. Post hoc tests indicate a significant difference between samples collected at 36 h and those collected at 0 h (***signifies P<0.01) for the ACTH treatment and no difference between 0 h and all other collection times for the control treatment.
Citation: Journal of Endocrinology 219, 1; 10.1530/JOE-13-0123
After the assessment of the effect of enzymatic cleavage of glucuronide and sulphate polar bonds using β-glucuronidase, it was found that faecal samples extracted without and with enzymatic pre-treatment had faecal cortisol concentrations of 17.50±5.13 and 223.50±18.20 ng/g (mean±s.d.; t-test: P<0.00, t=30.82, df=14).
Steroid partitioning in plasma
The mean values for free cortisol, CBG-bound and albumin-bound concentrations between the treatments highlight a significant difference in the ACTH treatment compared with the control treatment (two-way repeated-measures ANOVA: free: P=0.0011, F3, 21=7.865; CBG bound: P=0.0001, F3, 21=11.41 and albumin bound: P=0.0002, F3, 21=10.68; Figs 3, 4 and 5 respectively). In all the tests, the mean values in the post hoc tests indicated a significant increase from the pre-injection (T0) and post-injection collection times for the ACTH treatment compared with control treatment, for which the values did not increase after injection (Figs 3, 4 and 5).
The difference in the mean (±s.e.m.) concentration of free cortisol between the control and ACTH treatments over time. It highlights the increase in free cortisol concentrations after injection (T0) for the ACTH treatment compared with no difference in free cortisol concentrations after injection for the control treatment. ****signifies P<0.001.
Citation: Journal of Endocrinology 219, 1; 10.1530/JOE-13-0123
The difference in the mean (±s.e.m.) concentration of CBG-bound cortisol between the control and ACTH treatments over time. It highlights the increase in CBG-bound cortisol concentrations after injection (T0) for the ACTH treatment compared with no difference in CBG-bound cortisol concentrations after injection for the control treatment. ****signifies P<0.001.
Citation: Journal of Endocrinology 219, 1; 10.1530/JOE-13-0123
The difference in the mean (±s.e.m.) concentration of albumin-bound cortisol between the control and ACTH treatments over time. It highlights the increase in albumin-bound cortisol concentrations after injection (T0) for the ACTH treatment compared with no difference in albumin-bound cortisol concentrations after injection for the control treatment. ****signifies P<0.001.
Citation: Journal of Endocrinology 219, 1; 10.1530/JOE-13-0123
The mean values for cortisol partitioning (free, CBG bound and albumin bound) were converted to percentages of the total excreted cortisol for each treatment (control and ACTH) (Fig. 6). There was a significant interaction between treatments (ACTH and control) and the partitioning of cortisol (free, CBG-bound and albumin-bound cortisol) (two-way repeated-measures ANOVA: P≤0.0001, F2, 102=17.54). Post hoc tests indicated that when compared with those during the control treatment, the percentage of albumin-bound cortisol increased and that of CBG-bound cortisol decreased during the ACTH treatment, while that of overall free cortisol increased marginally but not significantly between the treatments (Fig. 6).
The mean (±s.e.m.) percentage of free and bound (CBG and albumin) cortisol for each treatment (ACTH and control).
Citation: Journal of Endocrinology 219, 1; 10.1530/JOE-13-0123
Discussion
This study shows that the elevation of plasma cortisol concentrations in koalas is reflected in the cortisol/cortisol metabolites of faecal pellets. The intestinal passage causes a time delay between the circulation of steroids in plasma and their appearance in faeces, and this delay in faecal GC excretion is correlated with gut passage time (confluence of the bile fluid to the rectum; Schwarzenberger et al. 1997, Mostl et al. 2005, Palme et al. 2005). This time delay can range from 30 min to more than 1 day depending on the species and activity of the animal, which can affect the rate of gut passage (Young et al. 2004, Palme et al. 2005, Thiel et al. 2005, Touma & Palme 2005, Wasser & Hunt 2005). The 36-h delay between the elevation of plasma cortisol concentrations and the appearance of cortisol and its metabolites in the faecal pellets is similar to the findings reported for some vertebrates, but longer than that reported by other studies (Table 1). This is probably a result of the slow gut passage time of koalas. The mean retention time for particle markers for the koala has been recorded between 39 h (Krockenberger & Hume 2007) and 100 h (Cork & Warner 1983), while the mean retention time for solute markers has been recorded between 110 h (Krockenberger & Hume 2007) and 213 h (Cork & Warner 1983). The time delay of 36 h in the appearance of cortisol metabolites in faecal pellets is close to the average mean retention time of particle markers of 39 h reported by Krockenberger & Hume (2007). While the results of this study show a pattern of steroid excretion that is similar to that reported for several eutherian species, a study by Ellis (1997) using injected 14C-cortisol in koalas showed that while gut passage time is slow in koalas, some individuals did show an elevation of faecal cortisol levels as early as 20 h after injection. This timing is similar to the appearance of 14C-cortisol in urine (Ellis 1997), and it may be related to an obligate release of some urine into the cloaca during defaecation, resulting in the contamination of pellets with a small quantity of urine. In females, the urethral opening is deep within the cloaca, which would allow the faecal pellets more contact with any residual or freshly voided urine than is the case with males. Although only one female was included in the study, it is interesting to note that the faecal cortisol concentration detected in pellets from the female was higher than that detected for males at all collection times after 24 h. Consistently, Nayaran et al. (2013) found sex-related differences in faecal cortisol metabolite (FCM) levels in captive koalas. The percentage rise in FCM levels of the males and females after the ACTH challenge was 25.53 and 71.79% respectively. These observations suggest that future studies of faecal steroid metabolite excretion in the koala, and indeed in all marsupials, should consider this possible sex difference. However, the faecal pellet collectors in our study ensured that only freshly deposited pellets that were not near any area containing voided urine were collected for later analysis. Urine contamination is still feasible (the pellets may always have a trace of urine on them from the cloaca), but all possible measures were taken to reduce its impact and this contamination is unlikely to have confounded our results.
Comparison of faecal cortisol concentrations and different delay times of cortisol metabolite appearance in faeces for different species reported in the literature
| Species | Faecal cortisol (basal/control/pre-treatment) | Faecal cortisol (ACTH/peak/post-treatment) | Delay time (h) | References |
|---|---|---|---|---|
| Columbian ground squirrels (Spermophilus columbianus) | 263.03±61.10 ng/g | 416.87±57.78 ng/g | 7±0.53 | Bosson et al. (2009) |
| Snowshoe hares (Lepus americanus) | 1054±151 ng/g | 11 130±3317 ng/g | 10 | Sheriff et al. (2009) |
| African elephants (Loxodonta africana) | 21–168 nmol/kg (median: 48) | – | 20–25.5 | Stead et al. (2000) |
| Spotted hyenas (Crocuta crocuta) | – | – | 26±5 | Goymann et al. (1999) |
| Male: 50.2±20.4 ng/g | Male: 576.0 ng/g | Male: 16 | Benhaiem et al. (2012) | |
| Female: 37.3±13.7 ng/g | Female: 913.9 ng/g | Female: 32 | ||
| African wild dogs (Lycaon pictus) | – | 1200–8000 ng/g | 24–30 | Monfort et al. (1998) |
| Himalayan black bear (Ursus thibetanus) | 107.9±8.0 ng/g | 432.6 ng/g | 24 | Young et al. (2004) |
| Koala (Phascolarctos cinereus) | 29.74±6.99 ng/g | 261±97.93 ng/g | 36 | This study |
| Cheetah (Acinonyx jubatus) | 628.4±95.3 ng/g | 7193.2 ng/g | 24–48 | Young et al. (2004) |
| Black-footed ferret (Mustela nigripes) | 220.2±19.3 ng/g | 548.6 ng/g | 24–48 | Young et al. (2004) |
In this study, following i.m. injection of ACTH, plasma cortisol concentrations were elevated at about 36 h, a delay expected with the administration of ACTH via this route. A recent study by Narayan et al. (2013) investigating non-invasive stress assessment methods in koalas has reported lower ACTH-stimulated plasma cortisol concentrations than those found in our study. This difference may be due to the use of the non-depot form of Synacthen by Narayan et al. (2013) rather than the long-acting Synacthen Depot that we employed. It is also interesting to note that FCM concentrations reported by Narayan et al. (2013) following ACTH stimulation were considerably lower than those that we found. While Narayan et al. (2013) used an assay for FCMs that employed a polyclonal antibody, they did not use enzymatic cleavage using glucuronidase/sulphatase prior to the extraction. Our study employed enzymatic cleavage prior to steroid extraction, which should have liberated free (unconjugated) cortisol, making this available for solvent extraction followed by ELISA known to specifically target cortisol.
Quite clearly, as mentioned by Touma & Palme (2005) and acknowledged by Narayan et al. (2013), antibodies used in some assays might not be ideal because they are produced primarily to measure concentrations of the respective unmetabolised steroid in plasma. To fully answer this question, an exhaustive HPLC study would be required to identify the primary FCMs in koala faeces and to ensure that appropriate antibodies are used to target the FCMs present at the highest concentration. We note that many studies of faecal GC excretion have not included the enzymatic cleavage procedural step to account for the fate of a significant proportion of the free cortisol conjugated by hepatocytes and entering the small intestine in bile secretions. Prior to the extraction of cortisol from faecal pellets, we suggest that it is important to treat samples with glucuronidase, which cleaves polar bonds and liberates extractable unconjugated cortisol. This is important because paired assays using a common pool of faecal extracts with or without prior enzymatic cleavage reveal considerably higher cortisol concentrations in the former compared with the latter. It is likely that FCMs of different molecular structures may not interact equally with the antibody provided with ELISA kits or RIAs designed to sensitively measure cortisol concentrations. In his study, Ellis (1997) employed solid-phase trapping (Sep Pak C18 solid-phase cartridges) in conjunction with glucuronidase cleavage of polar cortisol metabolites to extract cortisol. Faecal 14C-cortisol/cortisol metabolites increased 24 h after injection, and he suggested that the analysis of faecal cortisol/cortisol metabolites would be a very useful method to be employed for the assessment of stress in koalas.
We suggest that it is prudent to use enzymatic cleavage prior to solvent partitioning or solid-phase extraction of the target deconjugated steroid. For these reasons, we are confident that our assay system measured the concentrations of extracted cortisol.
The plasma cortisol concentrations found in this study confirm the finding of generally low plasma concentrations of this steroid in other studies of koalas (Weiss & Richards 1970, Scoggins 1978, McDonald et al. 1990, Ellis 1997) and relatively low concentrations compared with other mammals (e.g. koala: basal 1.101±0.53 ng/ml, ACTH 9.133±0.61 ng/ml (this study); koala: 2.86–7.17 ng/ml (range) (Hajduk et al. 1992); koala: 0.0002–0.18 ng/ml (range) (Ellis 1997); fruit bats: Pteropus hypomelanus 1269±207 ng/ml (basal) and P. vampyrus 596±67 ng/ml (basal) (Windmaier & Kunz 1993); Steller sea lion (Eumetopias jubatus): Russia 151.5±3.2 ng/ml, Southeast Alaska 152.7±2.7 ng/ml and Southwest Alaska 134.9±2 ng/ml (Myers et al. 2010); and Tasmanian devil (Sarcophilus harrisii): male 49±9.19 ng/ml (capture) and female 74.0±3.24 ng/ml (capture) (Jones et al. 2005)).
Most studies have not extended the treatment of GC excretion by calculating the partitioning of the principal adrenocortical hormone into the three plasma compartments: free, CBG bound and albumin bound (Bradley et al. 1980, Bradley 2003, Breuner et al. 2013). This study has shown that plasma free cortisol concentrations are increased significantly in response to ACTH administration and that the extent of this increase is not adequately described by just measuring plasma total cortisol concentrations (Breuner et al. 2013). Sheriff et al. (2010), working with snowshoe hares, reported that plasma free cortisol levels mirrored bile and FCM levels, but plasma total cortisol levels did not. Since it is the free hormone fraction that is removed by the hepatocytes during the passage of blood through the hepatic microcirculation, the free cortisol is clearly most relevant to the appearance of conjugated steroids in both urine and faeces.
While the plasma cortisol concentrations that occur in the koala are at the low end of the spectrum reported for other mammals, steroid partitioning within the blood plasma reveals a pattern that is quite similar to that of other mammals with most of the steroid being tightly bound to CBG and about 30% being bound more loosely to albumin, while about 10% appears as free or unbound hormone. However, it is clear that following ACTH stimulation, the relative sizes of the free and albumin compartments increase to a greater extent than does that of the CBG compartment. McDonald et al. (1990) described high-affinity binding between cortisol and CBG in koala plasma; however, they appear to have omitted an exponent from the stated affinity constant. Based upon the discussion in their book chapter in which a binding affinity that is much lower for the koala than for other marsupials is mentioned, we assume that the affinity constant stated should have been 0.90×107/M−1. If this is correct, then our chosen mid-range affinity constant of 3.5×107/M−1 would be too high and as a consequence the partitioning of cortisol within the blood plasma for koalas in our study would have resulted in much higher free and albumin-bound cortisol values than those appearing in our graphs. Clearly, dialysis experiments using koala plasma should be repeated to resolve this uncertainty.
It is apparent that there is a roughly tenfold increase in both plasma total and free cortisol concentrations from 12 to 36 h after ACTH injection. FCM levels increase significantly by 36 h after ACTH injection; however, the increase in FCM levels from time 0 is much smaller than changes in plasma cortisol concentrations, although FCM levels do remain elevated above FCM levels for controls for several days. This difference exists because hepatic clearance and excretion of polar GC metabolites in mammals via the renal route usually account for around 70% of all GC metabolites, excreted as β-glucuronides and sulphates (cortisol metabolites, CMLs), while the remaining 30% is excreted as polar conjugates in bile into the upper gastrointestinal tract (GIT). For this reason, FCMs appearing in the faecal pellets should represent only a small part of the total cortisol metabolites excreted. The FCMs are also, understandably, lower in concentration than the plasma cortisol.
At the commencement of the experiment, the FCM levels for the controls were higher than those for the ACTH-treated group. This observation, together with the gradual decrease in FCM levels for the controls during the experiment, is consistent with some of the control animals being exposed to some form of stressor 1 or 2 days prior to the commencement of the experiment. The FCM concentrations detected in this study following ACTH administration were considerably higher than those reported by Narayan et al. (2013), and this may, in part, be related to our use of the depot form of ACTH, which when administered via the i.m. route in other mammals is known to cause a significant elevation of plasma cortisol concentrations for 24–36 h (Novartis Pharmaceuticals Australia Pty Ltd.) after injection.
Passage time through the GIT of the koala is relatively slow (Krockenberger & Hume 2007), and this may allow time for some bacterial degradation of the CMLs to occur, resulting in the levels of extractable FCMs from faecal pellets being lower than might otherwise be found. When this FCM analysis is being considered as a stress assessment tool, it is important to note that 36 to 144 h after ACTH injection, there is a consistent elevation of FCM levels in faecal pellets.
We conclude that the results of the ACTH challenge in rehabilitating wild koalas validate this non-invasive technique for the monitoring of adrenocortical activity in free-ranging wild koalas. This study also validates the use of faecal cortisol analysis for the assessment of the activity of the hypothalamo–pituitary–adrenocortical axis of koalas using freshly collected koala faecal pellets.
Future studies could implement this non-invasive faecal sampling method to assess the physiological stress of koalas in wild populations that differ in exposure to various environmental and physiological (e.g. disease or injury) stressors. Further assessment should be made by suppressing endogenous GC concentrations using dexamethasone and examining the plasma and CML response to exposure of koalas to a predator such as a dog (Sheriff et al. 2009). This technique could also be extended to evaluate the impact of climate change, which is contributing to population declines, and how environmental stressors may affect the survival and reproductive success of koala populations throughout Australia.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
This work was supported by the Australian Koala Foundation, South West NRM, and the Australian Research Council (ARC Linkage Project 0882090).
Author contribution statement
N D and A B contributed to the design of the study, sample analysis, statistical analysis and interpretation and manuscript drafting. A G preformed the ACTH challenge on the animals. A G, L S, G B, D L and C M participated in the conception and design of the study and helped to draft the manuscript. All authors read and approved the final manuscript.
Acknowledgements
The authors gratefully acknowledge the generous financial support provided by the Australian Koala Foundation, Australian Research Council (ARC Linkage Project 0882090) and South West NRM. They are extremely grateful for the assistance provided by all of the Australia Zoo Wildlife Hospital staff and volunteers while running the experiment. They also thank Jonathan Rhodes for his input. The approved protocol for this project was obtained from the Animal Ethics Committee at The University of Queensland (GPA/603/08/ARC) and the Environmental Protection Agency (now the Department of Environment and Resource Management) (Scientific Purposes Permit WISP10160211).
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