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. Author manuscript; available in PMC: 2022 Sep 1.
Published in final edited form as: Stress. 2021 Jan 22;24(5):584–589. doi: 10.1080/10253890.2020.1868431

Infrared thermography is an effective, noninvasive measure of HPA activation

Jenny Q Ouyang 1, Paul Macaballug 1, Hao Chen 1, Kristiana Hodach 1, Shelly Tang 1, Jacob S Francis 1
PMCID: PMC8295405  NIHMSID: NIHMS1676309  PMID: 33480292

Abstract

Infrared thermography (IRT) is increasingly applied as a noninvasive technique for measuring surface body temperature changes related to physiological stress. As a basis for validation of IRT as a tool for diagnostic use, we need to assess its potential to measure hypothalamic-pituitary-adrenal (HPA) axis reactivity. We used experimental manipulations of the HPA axis in house sparrows (Passer domesticus), i.e. adrenal tissue responsiveness to exogenous adrenocorticotropin (ACTH) and the efficacy of negative feedback using the synthetic glucocorticoid dexamethasone (DEX), to test whether IRT is an effective tool for measuring HPA reactivity. Experimental birds showed a pronounced decrease in skin temperatures after ACTH injection and an increase in temperature after DEX injection. However, individual variation in glucocorticoid levels were not related to skin temperatures except after ACTH injection in experimental birds. We show that IRT can be used to measure HPA reactivity but that skin-temperature is not a good index for glucocorticoid secretion at baseline levels. These results suggest that while IRT of skin temperatures is a useful, noninvasive measure of HPA axis reactivity under acute activation, this technique might not be suitable for measuring natural variation of circulating glucocorticoid levels.

Keywords: Glucocorticoids, temperature, IRT, ACTH, DEX, house sparrow

Introduction

The vertebrate stress response is critical to survival yet difficult to quantify (Romero & Wingfield, 2016). It spans multiple levels of organization that ranges from the molecular to the organismal (Feder & Hofmann, 1999). Acute stress triggers two functionally linked physiological systems: the sympathetic-adrenal-medullary (SAM) system and the hypothalamic-pituitary-adrenal (HPA) axis (Charmandari et al., 2005; Johnson et al., 1992). SAM activation causes a rapid drop in skin temperature due to vasoconstriction, and this influx of peripheral blood, along with stress-induced thermogenesis, simultaneously increases core temperature for organs with the greatest metabolic need (Oka et al., 2001). This core temperature increase, termed “stress-induced hyperthermia” (SIH), is proportional to stressor intensity (Groenink et al., 1994) and forms the basis of new pathological screens for biomedical research and animal welfare assessment (Adriaan Bouwknecht et al., 2007; Head et al., 2000; Stewart et al., 2005).

Activation of the hypothalamic-pituitary adrenal (HPA) axis is another critical component of the physiological stress response. Upon perception of stressors, the avian hypothalamus is activated to secrete arginine vasotocin (AVT, a congener of the mammalian arginine vasopressin) and corticotropin-releasing factor (CRF). Both stimulate the pituitary to release adrenocorticotropin (ACTH), in turn, causing the release of glucocorticoids (corticosterone in birds) from the adrenals (Harvey et al., 1984). This response is believed to be aimed at maintaining or restoring homeostasis, thereby helping the animal to survive a stressful episode (McEwen & Wingfield, 2010).

Measuring the stress response via SAM or HPA activity in free-living vertebrates is not simple. Implanting a probe to measure core temperature or taking a blood sample to measure glucocorticoid levels are both invasive, and can change SAM and HPA activity. Capturing and handling illicit a stress response, so simply by using these invasive techniques to measure animals’ physiological state, we change the very response we are interested in quantifying (Angelier et al., 2010). Recently, a novel method has been proposed to non-invasively measure the stress response using skin temperatures (Jerem et al., 2015; Stewart et al., 2005). Studies show that skin temperatures correlate to the intensity and duration of stressors, and are a good proxy for body condition of free-living and captive birds (Herborn et al., 2015; 2018; Jerem et al., 2018). These skin temperature changes could be the direct result of an activation of the HPA axis, which then activates the SAM system (Stewart et al., 2008). Studies both in vivo and in vitro demonstrate this possibility, i.e. glucocorticoids are required for catecholamine-mediated mobilizations of free fatty acids, for shivering responses, and for vasoconstriction (Deavers & Musacchia, 1979). While there is evidence for a link between HPA and SAM activation and skin temperature, these three systems are not always linked mechanistically and can respond independently to different noxious stimuli. Thus, if skin temperature were to be used to measure HPA axis activation, one would need to directly stimulate the HPA axis rather than providing a general external stressor.

To validate the use of IRT in relation to HPA axis activation and deactivation, we tested the relationship between skin temperature and glucocorticoid levels after activation of the adrenal tissue via exogenous adrenocorticotrophic hormone and efficacy of negative feedback of the HPA axis using the synthetic glucocorticoid dexamethasone (DEX). Additionally, we assessed the relationship between SAM and HPA activity by exploring inter-individual glucocorticoid-temperature correlations throughout the experiment (with skin temperature used as a proxy for SAM activity). If SAM and HPA activity are connected, we would expect experimental treatment to affect skin temperature levels. If the two systems are disconnected, experiment treatment would not affect skin temperature and there would be no correlation between temperature and glucocorticoid levels.

Methods

All house sparrows were captured as juveniles fall of 2017 and stayed in outdoor aviaries until the start of the experiment (February 2018). Each day, 4 house sparrows (total sample size: 14 males and 11 females) were captured at random and transported to an indoor facility. They were placed in cages (23 cm × 23 cm × 13 cm; more detail below) with ad libitum water and food and allowed to acclimate for 1 hr (see Figure 1 for experimental procedure). We previously tested the acclimation procedure with respect to GC levels at 1, 2, 3, and 24 hr intervals and noted that GC levels remained the same after 1 hr of acclimation inside (Supplementary Figure S1; Rich & Romero, 2005). We took an initial blood sample from the brachial vein after 1 hr (time point 0) and another blood sample after 30 mins (time point 1; to confirm stable corticosterone and temperatures). Following the blood sample at time 1, the bird was randomly assigned to treatment or control groups. Treatment birds received an injection of ACTH (100 IU/kg) and control birds received an injection of saline. 30 mins after the injection of ACTH or saline, we took the third blood sample (time point 2) and gave experimental birds an injection of dexamethasone (7 μg/kg; Sigma Aldrich reconstituted crystalline form) and control birds an injection of saline. After 60mins, we took the final blood sample (time point 3). All weight-dependent doses were standardized for a 28 g bird, and given in 50 μl (Rich & Romero, 2005) and injections were given intramuscularly. Sampling time points were determined for house sparrows for maximal and minimum activation and deactivation of the HPA axis (Rich & Romero, 2005). DEX was chosen as the compound to measure negative feedback, as it has been shown to bind to GR receptors in house sparrows with high affinity (Breuner & Orchinik, 2009). Each blood sample was less than 20 μl for a total of 80 μl maximum of whole blood. Blood samples were immediately placed on ice and centrifuged at 16,000 g for 10mins. Plasma was stored in −80 °C until analyses. Protocols were approved by Institutional Care and Use Committees (IACUC) of the University of Nevada, Reno, Nevada Department of Wildlife, and Federal banding permits.

Figure 1.

Figure 1.

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Experimental procedure. House sparrows were monitored via infrared thermography for skin temperatures and blood samples collected for glucocorticoid levels. Blood samples were taken at four time points: T0 = initial sample after 1 hr in the cage, T1 = second sample after 30 min to test acclimation process, T2 = 30 min after ACTH or saline injection, T3 = 1 hr after DEX or saline injection.

Hormone assays

To measure plasma corticosterone, we used enzyme-linked immunosorbent assay kits (Enzo Life Sciences; Farmingdale, NY, USA) following the manufacturer’s instructions, with a standard curve on each plate. To validate this assay for use with house sparrow plasma, we first removed endogenous hormones from the plasma by incubating it for 20 minutes in a solution of 1% charcoal and 0.1% dextran. We then added sufficient corticosterone standard from the assay kit so that the concentration of corticosterone in each stripped plasma sample was equal to 500 pg/mL. We assayed each stripped and spiked sample at three dilutions (1:20, 1:30, and 1:40) and each dilution with two concentrations of steroid displacement reagent (SDR; 0.5% and 1% of plasma volume). Based on this optimization, we determined that for subsequent assays, house sparrow plasma should be diluted 1:40 with 0.5% SDR. We randomly assigned samples across four plates, with the exception that all samples from the same individual were on the same plate. We included a standard curve on each plate, which ranged from 32 pg/mL to 20,000 pg/mL. The assay sensitivity was 2.1 pg/ml. To calculate intra- and inter-plate coefficient of variation (CV), we also included three pooled house sparrow samples on each plate, and each pool was assayed in triplicate. The intra-plate CV was 10.3% and inter-plate CV was 3.3%.

Temperature measurements

We used a Flir T420 to measure external thermal temperature (FLIR systems Inc, Wilsonville Oregon) and a Mastech MS8269 digital multimeter with a type K thermocouple to measure the room temperature continuously during the experiment. The cage was insulated with black foam to reduce infrared reflection and had one side made of wire mesh, encouraging the bird to position itself in front of the camera, and allowing for measurement of infrared emissions. A camera was placed 43 cm away from the cage, such that the entire cage fit within the camera’s view. The observers were not visible to the birds during the course of the experiment. We analyzed the recordings using Flir Tools + Program. Briefly, we picked 4 points localized at the skin around the bird’s eye and averaged these points to obtain the skin temperature (Jerem et al., 2015). We used an constant emissivity of 0.97 for estimating birds’ skin temperature, standard values for animal skin range from 0.95 to 0.98 (Best & Fowler, 1981). We collected skin temperatures continuously throughout the experiment, via fully radiometric video, but focus our analyses on 4 time points that correspond with the experimental treatment and blood sampling (2 mins before blood sampling).

Statistical analysis

We conducted statistical analysis using R v.3.6 (R core team). We used linear mixed models (LMMs) to test whether skin temperatures and corticosterone levels were different between control and experimental groups with the interaction of treatment (experiment or control) and time point. We ran Tukey post-hoc multiple comparison tests for the interaction to test whether each treatment group was different from others. We included sex as a fixed effect and individual as a random effect to account for repeated measures. Room temperature was also included as a covariate, but was removed as it was insignificant in both models. To see if individual corticosterone levels were correlated to skin temperatures throughout the experiment, we ran spearman’s correlation for the different time points. All final models met assumptions of normality (corticosterone was log transformed) and homoscedasticity of residual errors, and significance was taken at α = 0.05.

Results

Skin temperatures around the eye significantly differed by time point and treatment (Table 1; Figure 2(A)). Specifically, temperatures decreased after ACTH injection in experimental birds (experimental group time 1 to 2: β = −1.0, SE = 0.3, t = −3.8, p = .006). Temperatures increased after DEX injection in experimental birds (experimental group time 2 to 3: β = 0.9, SE = 0.3, t = 3.4, p = .02). Control birds did not differ throughout the experiment (p’s > .2). Control birds had higher temperatures than experimental birds at time 2, 30 minutes after experimental birds received an ACTH injection (β = 1.2, SE = 0.3, t = 4.4, p = .001).

Table 1.

Model estimates for the effect of ACTH and DEX treatment on corticosterone levels and skin temperature in house sparrows.

Variable Estimate SE t p-value
LMM for skin temperatures
 (Intercept) 36.0 0.32 111.80 <.001
 Treatment 0.24 0.36 0.67 .50
 Time point 1 (reference is 0) −0.03 0.28 −0.11 .91
 Time point 2 (reference is 0) −0.28 0.28 −1.00 .32
 Time point 3 (reference is 0) −0.42 0.27 −1.55 .12
 Sex −0.08 0.29 −0.27 .78
 Treatment × Time point 1 −0.12 0.38 −0.31 .75
 Treatment × Time point 2 −0.89 0.39 −2.28 .02
 Treatment × Time point 3 0.17 0.38 0.44 .66
LMM for corticosterone
 (Intercept) 40.71 7.77 5.24 <.001
 Treatment 0.24 8.50 0.03 .98
 Time point 1 (reference is 0) −7.35 5.34 −1.38 .17
 Time point 2 (reference is 0) −14.99 5.35 −2.80 .005
 Time point 3 (reference is 0) −6.96 5.35 −1.30 .19
 Sex −4.40 7.40 −0.59 .55
 Treatment × Time point 1 −8.54 7.56 −1.13 .26
 Treatment × Time point 2 17.56 7.56 2.32 .02
 Treatment × Time point 3 −15.06 7.56 −1.99 .04
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Individual estimates are given from summary statistics of the LMM. Random effects include individual ID. Time point is 0 (initial), 1 (after 30 mins), 2 (after ACTH or saline injection, 3 (after DEX or saline injection).

Figure 2.

Figure 2.

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Temperature (A) and corticosterone concentrations (B) as a function of experimental time points. Time point 0 is initial sample, point 1 is after 30 minutes, point 2 is 30 minutes following an ACTH or saline injection, and point 3 is 60 minutes following a DEX or saline injection. Plotted are 95% confidence intervals and means for control (solid black line) and experimental (dotted red line) birds. Experimental and control points are jittered for clarity.

Corticosterone levels significantly differed by time point and treatment (Table 1; Figure 2(B)). Specifically, corticosterone levels increased after ACTH injection in experimental birds (experimental group time 1 to 2: β = 18.5, SE = 5.4, t = 3.4, p = .02). Corticosterone levels also decreased after DEX injection in experimental birds (experimental group time 2 to 3: β = −24.6, SE = 5.4, t = −4.6, p = .0004). Although corticosterone levels decreased from time point 0 to 1, it was not significantly lower in post-hoc analyses (β = −15.9, SE = 5.4, t = −2.8, p = .07). Control birds’ corticosterone levels did not change throughout the experiment (p’s > .1). Corticosterone levels were higher in experimental birds than control birds at time 2 (β = −17.8, SE = 8.5, t = −3.0, p = .04) and lower in experimental birds than control birds at time 3 (β = 22.0, SE = 5.3, t = 4.1, p = .002).

Skin temperatures and corticosterone levels were not correlated at any time point (e.g. after DEX injection, t = 0.4, R2 = 0.02, p = .7) except after ACTH injection (time point 2) only for the experimental birds (t = –2.19, p = .03). Experimental birds with higher corticosterone levels had lower skin temperatures (Figure 3).

Figure 3.

Figure 3.

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Skin temperature negatively correlated with corticosterone levels only after ACTH injection for experimental birds (solid black line and filled circles; R2 = 0.17). Control birds are shown in open circles.

Discussion

We validated the use of IRT in assessing HPA activity in an avian system. We found that skin temperatures decreased with HPA activation and increased with negative feedback. However, individual variation in skin temperatures and glucocorticoid levels were not correlated except during ACTH challenge, suggesting that a noninvasive IRT method may be most useful as a coarse measure HPA activity during levels of high HPA activation.

Control birds maintained the same temperature and glucocorticoid levels throughout the experiment. The increase in glucocorticoid levels after ACTH treatment showed appreciable corticosterone responses to the dosage used. This indicates that the adrenal tissue has reserve capacity to secrete corticosterone and is not maximally stimulated by the endogenous ACTH release. ACTH treatment significantly lowered skin temperatures for all individuals, consistent with previous research (McCafferty, 2013). However, a study on calves showed that eye temperature did not change after administration of ACTH, and it could be related to the amount of ACTH administered, timing of measurements or differences in endocrine gland sensitivity between an avian or mammalian model (Stewart et al., 2008). This effect in house sparrows happened within 5 minutes of ACTH treatment suggesting that HPA activity causes a rapid cooling of skin temperatures (Herborn et al., 2015). Steroid hormones may take up to 3 minutes to circulate (Baugh et al., 2013) and it seems that within 5 minutes, the effect is already seen in the SAM and stayed cool until 30 minutes later. Free-living blue tits (Cyanistes caeruleus) responded to capture within 10 seconds with a drop of 2 °C (Jerem et al., 2015). ACTH causes an increase in circulating glucocorticoids for at least 30 minutes (Rich & Romero, 2005), and skin temperatures remained cool during the course of HPA stimulation (this study; note we measured skin temperatures 28 and 58 mins after ACTH and DEX injections, respectively). Activation of the HPA axis is known to increase shivering responses and vasoconstriction, cooling extremities (Deavers & Musacchia, 1979). Therefore, stress-induced hyperthermia (SIH) does happen after adrenal activation, most likely due to the influx of circulating glucocorticoids.

The efficacy of negative feedback using DEX was clearly indicated by the drop in corticosterone levels after 60mins of DEX injection, in parallel with a significant increase in skin temperatures. DEX binds glucocorticoid receptors with high affinity in house sparrows, allowing for this synthetic glucocorticoid to be used effectively to test negative feedback of the HPA axis (Breuner & Orchinik, 2009). HPA negative feedback could also affect SAM by decreasing circulating GCs and raising skin temperatures, indicated by skin temperatures increasing after DEX injection. Additionally, ACTH levels would be declining by the time we took temperature measurements after DEX injection and may implicate ACTH itself as inducing the temperature change via a non-steroid mechanism (Lipton et al., 1981).

However, individual variation in glucocorticoid levels were not related to skin temperatures in control individuals and experimental individuals at any time point except after ACTH injection. Core temperatures are linearly related to corticosterone levels in mice (Veening et al., 2004). However, the relationship between skin temperature and glucocorticoid secretion may not be linear for a few reasons. Although skin temperature can be measured noninvasively, it is inherently more variable and dynamic than core temperature (Giloh et al., 2012). Herborn et al. showed that the change in skin temperature due to different stressor intensities is not a linear relationship (Herborn et al., 2015). Moreover, skin temperatures vary with body condition and ambient temperature, further complicating the relationship between corticosterone secretion and magnitude in SIH (Jerem et al., 2018). For example, eye surface temperature response and amplitude was unrelated to baseline corticosterone in one study (Jerem et al., 2018) but negatively correlated to free corticosterone (~96–99% of total glucocorticoids are bound by globulins) in another study (Jerem et al., 2019). We did not measure free corticosterone and perhaps the interaction between HPA and SAM affects binding globulin activity. Glucocorticoids do influence the sympathetic nervous system, but the intensity of the interaction of the HPA and SAM systems likely depend on the stressor intensity (Wadsworth et al., 2019). It is worthwhile to note that there is a negative inter-individual relationship between skin temperature and corticosterone levels after ACTH treatment, suggesting that stimulation of the adrenals can lead to stronger among-individual correlation of SAM and HPA activity, and that IRT may be an effective method for measuring HPA activity during acute stress.

We tested the relationship between SAM and HPA activity with adrenal activation and negative feedback and found that IRT can detect differences in HPA reactivity. However, individual variation in glucocorticoid levels only correlated to skin temperatures during ACTH-stimulated glucocorticoid release, suggesting that the interactions of the two systems at baseline levels are likely more relaxed than when subjected to maximal adrenal activation. We propose that IRT may offer researchers a “coarse ruler” to assess HPA activity with a tradeoff between invasiveness and precision. We caution that strength, duration, and perception of a particular stressor affect HPA and SAM differently and should be taken into consideration before using IRT to quantify physiological stress.

Supplementary Material

supp materials

Acknowledgments

Funding

We thank two anonymous reviewers for their helpful comments. JQO is funded by NIH [P20 GM103650 and R15 ES030548] and PM received the Nevada Undergraduate Research Award and Tri Beta Beta research award for this work.

Footnotes

Supplemental data for this article can be accessed here.

Disclosure statement

No potential conflict of interest was reported by the author(s).

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