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Modulation of Energy Expenditure by Estrogens and Exercise in Women

Gavin, Kathleen M.1,2; Kohrt, Wendy M.1,2; Klemm, Dwight J.2,3; Melanson, Edward L.1,2,4

Exercise and Sport Sciences Reviews: October 2018 - Volume 46 - Issue 4 - p 232–239
doi: 10.1249/JES.0000000000000160
Articles
Video Abstract

Reducing estrogen in women results in decreases in energy expenditure, but the mechanism(s) remain largely unknown. We postulate that the loss of estrogens in women is associated with increased accumulation of bone marrow–derived adipocytes in white adipose tissue, decreased activity of brown adipose tissue, and reduced levels of physical activity. Regular exercise may counteract the effects of estrogen deficiency.

In women, loss of estrogens induces changes in several pathways that contribute to reductions in energy expenditure.

1Division of Geriatric Medicine, School of Medicine, University of Colorado Anschutz Medical Campus;

2Eastern Colorado VA Geriatric Research, Education, and Clinical Center, Denver;

3Division of Pulmonary and Critical Care Medicine, and

4Division of Endocrinology, Metabolism, and Diabetes, University of Colorado Anschutz Medical Campus, Aurora, CO

Address for correspondence: Kathleen Gavin, Ph.D., MS B179, 12631 East 17th Ave, Room 8111, University of Colorado Anschutz Medical Campus, Aurora, CO 80045 (E-mail: Kathleen.Gavin@ucdenver.edu).

Accepted for publication: April 24, 2018.

Editor: Nancy I. Williams Sc.D., FACSM.

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Key Points

  • Women transitioning through the menopause experience a decrease in fat-free mass, including bone and skeletal muscle, and an increase in fat mass, which is predominantly in the abdominal region. Studies in animals suggest that this is related to the loss of estrogens.
  • In humans, we have shown that reducing endogenous estradiol results in decreases in energy expenditure and increases in visceral adiposity. Our research is focused on understanding the mechanisms by which this occurs.
  • We have found that estrogen deficiency increases the accumulation of bone marrow–derived adipocytes in white adipose tissue of mice and decreases physical activity in women. Our preliminary data also suggest that estrogen may regulate brown fat activity. We postulate that collectively, these changes contribute to the observed decreased energy expenditure and increased central adiposity.
  • We hypothesize that regular exercise may help to counteract the effects of estrogen deficiency by attenuating the observed detrimental metabolic alterations.
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INTRODUCTION

The menopause transition is a period associated with changes in body weight and composition and fat distribution. Women transitioning through the menopause experience a decrease in fat-free mass (FFM), including bone and skeletal muscle, and an increase in fat mass (FM), which is predominantly in the abdominal region (1). Studies of both animals (2,3) and humans (4,5) suggest that this is related to the loss of ovarian function and, specifically, the loss of estrogens (5). Because the increase in central fat increases risk for several chronic diseases (6), understanding how estrogens contribute to these changes has important public health implications. A focus of our lab over the past decade has been to understand how the loss of estrogens influences body weight regulation and body fat distribution.

Our working model is that the loss of estrogen leads to decreased energy expenditure (EE), thereby disrupting energy balance and contributing to fat gain. This model is supported by studies of animals demonstrating that ovariectomy (OVX) causes excess fat gain and a decrease in total energy expenditure (TEE) that is the result of a dramatic decrease in physical activity (PA) and a suppression of basal metabolic rate (EE during light hours, when PA is low) (7). In addition, recent data demonstrate that the OVX-associated decrease in estradiol (E2) also influences energy homeostasis by decreasing bioenergetic function at the level of the mitochondrial membrane (8). These effects of OVX are prevented or reversed by E2 treatment.

Using a pharmacological approach, we demonstrated that, similar to OVX in animals, suppression of ovarian function with gonadotropin-releasing hormone (GnRH) analog therapy in women causes decreases in resting EE (REE), TEE, and PA and an increase in abdominal adiposity. In our first study of premenopausal women, 6 d of GnRH antagonist treatment to suppress ovarian function resulted in a reduction in REE (1334 ± 36 kcal·d−1, mean ± SE) when compared with measurement during the mid-luteal phase (1405 ± 42 kcal·d−1) and early follicular phase (1376 ± 43 kcal·d−1) (9). However, we could not determine if the decrease in REE was specifically due to a reduction in E2.

More recently, we studied the longer term effect of ovarian suppression on body composition and EE in premenopausal women (5,10). Because GnRH antagonists are short-acting, we used a GnRH agonist (GnRHAG, leuprolide acetate 3.75 mg, Lupron; TAP Pharmaceutical Products, Inc; Lake Forest, Ill) to suppress ovarian function. A single injection of leuprolide acetate produces an initial stimulation of anterior pituitary and gonadal sex steroids (for several weeks), which eventually leads to down-regulation of the GnRH receptor via a negative feedback loop, producing prolonged suppression of anterior pituitary gonadotropins and gonadal sex steroids (11). Repeated monthly dosing maintains ovarian hormone suppression. We also studied whether the effects of GnRHAG were prevented by E2 treatment (0.075 mg·d−1 of transdermal E2). Seventy premenopausal women were randomized to 5 months of GnRHAG plus E2 (GnRHAG + E2) or placebo (GnRHAG + PL) transdermal therapy. Change in FFM was different between the groups. FFM decreased in the GnRHAG + PL group (mean; 95% confidence interval; −0.6 kg; −1.0, −3.0) and was preserved in GnRHAG + E2 group (+0.3 kg; −0.2, 0.8; Fig. 1A). Although FM did not change in either group, abdominal subcutaneous (SubQ) and visceral fat areas (measured using computed tomography (CT)) increased in response to GnRHAG + PL but not GnRHAG + E2 (Fig. 1B). There was also a decrease in REE (adjusted for changes in FM and FFM) in the GnRHAG + PL group (~ −50 kcal·d−1) that was prevented by E2 add back (Fig. 2). Total daily EE (measured using whole-room indirect calorimetry) was decreased in response to GnRHAG + PL (~ −110 kcal·d−1), but this also decreased in the GnRHAG + E2 (~ −115 kcal·d−1). These results demonstrate that suppression of ovarian hormones in premenopausal women results in the loss of FFM, increased abdominal adiposity, and decreased REE, all changes that are specifically related to the loss of E2.

Figure 1

Figure 1

Figure 2

Figure 2

We are interested in the potential therapeutic benefit of exercise to attenuate the metabolic and bioenergetic consequences of the loss of estrogens. As an exploratory study, a subset of participants in each drug group of the 5-month GnRHAG intervention discussed previously were randomized to undergo progressive resistance exercise training during the drug intervention. Because of the exploratory nature of this study, no inferential statistics were performed. In this proof-of-concept study, resistance exercise seemed to be beneficial in attenuating the decreases in FFM and bone mineral density in response to the suppression of ovarian hormones (5). However, exercise did not seem to attenuate the declines in REE and TEE or increase in abdominal adiposity that occurred with ovarian suppression (10). Although preliminary, these observations suggest that maintaining a regular exercise program during the menopausal transition may be an effective strategy for minimizing some, but not all, of the consequences of the loss of estrogens.

We also have obtained preliminary evidence that E2 contributes to the regulation of PA in women (12). In the same intervention study described previously, PA was assessed by accelerometry for 1 wk before and during each month of the intervention. The GnRHAG + PL and GnRHAG + E2 groups had similar levels of PA at baseline and after 1 month of intervention, but the GnRHAG + E2 group had higher levels of moderate-to-vigorous PA (MVPA) during the final 4 months of intervention (Fig. 3). This divergence coincides temporally with when GnRHAG treatment would be expected to suppress ovarian function. Although the results of this study support the hypothesis that PA level is regulated by estrogens, as has been documented in studies of animals, the study was not powered to detect differences in PA and the findings should be interpreted cautiously. We are currently conducting a study with a larger sample size and more complete and objective assessment of PA and sedentary behavior to better understand the regulation of PA by E2.

Figure 3

Figure 3

In summary, preclinical and clinical studies have provided evidence that the loss of estrogens leads to unfavorable alterations in body composition (increase in central FM, decrease in FFM) and decreases in TEE, REE, and PA. In our studies, E2 therapy during ovarian suppression attenuated the changes in body composition and fat distribution and may preserve PA but seemed to have little or no effect on preserving REE or TEE. Our group is performing several lines of research to further elucidate the impact of E2 on body fat and EE regulation. The following sections discuss two adipose tissue level mechanisms that we believe may be important determinants of EE and body composition in women; production of bone marrow (BM)–derived adipocytes and brown adipose tissue (BAT) activity.

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ESTROGEN AND BM-DERIVED ADIPOCYTE PRODUCTION

Adipocyte Development

White adipocytes, the predominant fat cells in human adipose tissue, turn over at a rate of ~10% per year, as assessed by 14C turnover in genomic DNA (13), indicating that a source of progenitor cells is required for continued development of new adipocytes. Dogma has long held that all adipocytes arise from a common mesodermal precursor. These precursors give rise to adipose tissue-resident progenitor cells of a mesenchymal origin, which then commit to the preadipocyte lineage and subsequently develop into mature adipocytes (14).

In recent years, a number of laboratories have used lineage analysis and fate mapping strategies in mice to demonstrate that specific subsets of adipocytes are produced via distinct developmental pathways both within and beyond the traditional mesenchymal lineage (15–17), which may contribute to the biological diversity of regional adipose tissue depots. Of specific interest is the presence of a unique subpopulation of BM-derived adipocytes in white adipose tissue of mice (18) and humans (19,20) that arise from cells of myeloid rather than mesenchymal lineage (21,22).

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Physiological Relevance of BM-derived Adipocytes

BM-derived adipocytes arise from progenitors that develop in the BM, traffic though the blood as myeloid intermediates, and ultimately reside in the adipose tissue where they trans-differentiate into a mesenchymal-like progenitor and contribute to adipogenesis (Fig. 4) (21). We (19) and Ryden et al. (20) recently confirmed the presence of BM-derived adipocytes in humans. In independent investigations, donor-host chimerism analyses identified the presence of donor DNA in adipocytes isolated from SubQ adipose tissue samples of allogeneic bone marrow transplant (BMT) recipients.

Figure 4

Figure 4

These discoveries have shifted the dogma in adipose tissue biology, demonstrating that not only are some adipocytes produced from non-resident progenitors, but from an unexpected origin — the hematopoietic lineage. Importantly, BM-derived adipocytes can comprise from 5% to 25% of the total adipocyte pool in mice depending on sex (more in females) and depot (more in visceral) (23). Similarly, up to 35% of adipocytes in humans seem to be of BM origin (19,20). Obesity and time since BMT also seem to be important factors in humans, as both were significantly associated with increased presence of BM-derived adipocytes (20). In fact, Ryden et al. (20) found that the BM progenitor contribution to the adipocyte population was 2.5 times greater in severely obese compared with lean individuals. These results suggest that these adipocytes of novel lineage are sufficiently prevalent to influence physiology.

The most common fate mapping method for measurement of BM-derived adipocytes in mice uses lethal irradiation followed by transplantation of BM from donors expressing a fluorescent or luciferase protein (23). However, we have also used the myeloid-specific LysM gene promoter to indelibly label cells arising from the myeloid lineage with LacZ, identifying LacZ+ adipocytes at levels similar to our previous observations in BMT models (21). This non-BMT dependent model confirms that BM-derived adipocyte production is not dependent on irradiation or other insult associated with BM transplantation. Nevertheless, the lack of a unique biomarker for BM-derived compared with conventional mesenchymal lineage adipocytes precludes their measurement humans that have not undergone BMT. This limitation impedes the wider translation of these findings to the general human population. Development of a measurement tool for BM-derived adipocytes in humans is a critical step in carrying this field forward.

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The Biology of BM-derived Adipocytes

Global gene expression and principal component analyses reveal that BM-derived adipocytes are distinct from either conventional white or brown adipocytes and from stromal or circulating myeloid cells (21). Distinctive characteristics of BM-derived adipocytes include decreased expression of the genes for leptin, mitochondrial biogenesis, electron transport and ATP synthesis, and lipid oxidation, and increased expression of the genes for a number of inflammatory cytokines (e.g., Interleukin-6 and CXCL9) (21). If this gene expression profile translates into a metabolic phenotype, it suggests that BM-derived adipocytes are likely to have very low capacity for EE and potentially contribute to the metabolic sequela of obesity via their inflammatory and low leptin profile.

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Sex- and Depot-related Differences in the Production of BM-derived Adipocytes

BM-derived adipocytes are present in all major white fat depots of the mouse (19), but the prevalence is greatest in the gonadal (i.e., visceral) and para/epicardial depots (19,21). This regional specificity has led us to postulate that the accumulation of BM-derived adipocytes is a mechanistic underpinning for the metabolic dysfunction and chronic disease risk associated with abdominal and para/epicardial fat accumulation. Sex differences in the production of BM-derived adipocytes are also apparent. Although both sexes have a similar regional distribution of BM-derived adipocytes, the accumulation of BM-derived adipocytes is greater in female mice than males (Fig. 5) (21,24). This sex difference raises the possibility that sex hormones regulate the development of BM-derived adipocytes.

Figure 5

Figure 5

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Estrogen Regulation of BM-derived Adipocytes

The importance of the sex of the mice in our previous studies led us to investigate the role of ovarian hormones in regulating the production of these novel, and potentially metabolically harmful, adipocytes. Interestingly, we found that the loss of gonadal hormones in female mice induced by OVX resulted in greater accumulation of BM-derived adipocytes. This was particularly notable in the gonadal depot, where BM-derived adipocytes were increased by 40% to 100% when compared with wild-type (WT) mice (Fig. 6) (24). E2 treatment in OVX mice resulted in full attenuation of the increased accumulation of BM-derived adipocytes. Further, E2 seemed to prevent the production of BM-derived adipocytes through an estrogen receptor alpha (ERα)–mediated mechanism, because female ERα knockout mice also had 36% to 100% more BM-derived adipocytes than WT mice (Fig. 6).

Figure 6

Figure 6

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Exercise and BM-derived Adipocytes

To date, no studies have investigated how exercise may impact the accumulation of BM-derived adipocytes. However, we know that exercise training reduces the obesity associated infiltration of macrophages into adipose tissue and can decrease visceral adiposity and its associated inflammatory profile (25). Interestingly, BM-derived adipocytes appear to arise from the myeloid lineage, specifically via the transdifferentiation of macrophage (22). These adipose tissue resident macrophage arise from myeloid lineage cells of the BM (represented by CD45+/CD11b+ cells in Fig. 4) that traffic to the adipose tissue where they undergo transdifferentiation into adipocytes. Thus, lower macrophage infiltration resultant to exercise training may lead to fewer adipose tissue resident BM progenitors available to contribute to adipogenesis. Determining if exercise can prevent the production of BM-derived adipocytes, particularly in the estrogen deficient state, is a goal of our future investigations.

In summary, BM-derived adipocytes are present in both mice and humans and appear to accumulate preferentially in visceral and para/epicardial fat depots. Their gene expression fingerprint includes a lower contribution to EE, low leptin release, and an elevated inflammatory profile. Importantly, BM-derived adipocyte production seems to be regulated by E2 signaling through ERα. Therefore, we hypothesize that increased production of BM-derived adipocytes in post-menopausal women may be a mechanism underlying the observed increased visceral adiposity and decreased EE in this population. We are currently conducting studies to characterize the in vivo metabolic consequences of increased production of BM-derived adipocytes as well as developing methods to detect these novel adipocytes in non-BM transplanted humans.

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ESTROGEN AND BAT ACTIVITY

Evidence for BAT in Adult Humans

The primary function of BAT is to produce heat. BAT is highly vascularized and richly innervated by the sympathetic nervous system (SNS). When activated, BAT generates heat through mitochondrial uncoupling of oxidation and phosphorylation, mediated by uncoupling protein 1 (UCP1), which is uniquely and abundantly expressed in BAT mitochondria. BAT activity is stimulated by factors that increase SNS activity, including acute cold exposure. When active, BAT takes up both glucose and circulating non-esterified fatty acids (26).

BAT is present in rodents and some mammals throughout the lifecycle (27). Until recently, BAT was thought to be present in significant volume and activity only during infancy in humans. This view began to shift in the early 2000s, when radiological reports documented non–tumor-related 18F fluoro-2-deoxyglucose (18FDG) uptake in tissues with low radiodensity (indicative of an adipose tissue) (28). In these studies, bilateral, symmetrical 18FDG uptake was often observed in the cervical, clavicular, and paraspinal regions. Because these areas were within fat depots (based on CT Hounsfield units) and were more pronounced when patients were not kept warm, some nuclear imaging departments began referring to these as BAT. Subsequent studies established that 18FDG uptake in these areas was stimulated by cold exposure (29), and biopsy studies confirmed these areas as BAT via morphological assessment and identification of UCP1 (30).

The reported prevalence of adults with detectable BAT varies markedly. Retrospective analyses of large cohorts that had undergone clinical positron emission tomography/CT (PET/CT) scanning suggest that 5%–10% of adults have spontaneously detectable BAT (i.e., detected without purposeful cold exposure) and have identified a number of factors associated with BAT activation, including colder outdoor temperature at the time the scans were performed, female sex, younger age, and lower body mass index (BMI) and body fat content (31). Studies of cold-exposed individuals have since shown that the prevalence of adults with metabolically active BAT measured by 18FDG uptake ranges from 30% to 100% (32–34). Cold-induced BAT activity decreases with age (35) but has been observed in humans up to the age of 64 yr (32).

Cold-induced BAT activity is inversely associated with body fatness (29), suggesting that BAT plays a role in body weight regulation, but whether a decline BAT activity contributes to the development of obesity in postmenopausal women is unclear. BAT may play a role in reducing disease risk beyond its potential impact on body weight. Rodents with high levels of BAT activity have lower fasted glucose and triglycerides levels (36), and in humans higher levels of BAT are favorably associated with indices of insulin action and glucose tolerance (37). Furthermore, BAT activation reduces atherosclerosis in rodent models and is associated with reduced arterial inflammation and reduced risk of cardiovascular disease in humans (38).

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The Contribution of BAT to EE in Humans

Recent studies demonstrated that BAT oxidative metabolism contributes to the increased EE during acute cold exposure in humans. Using 11C acetate infusion during dynamic PET/CT scanning, these studies found that oxidative metabolism is increased in BAT, but not skeletal muscle or SubQ adipose tissue, during cold exposure (32–34). Interestingly, BAT oxidative activity has been observed in participants exposed to room temperature (~22°C) (32,33), suggesting there is a measurable level of BAT activity in humans even without cold exposure. The contribution of BAT activity to REE in humans is not known, although studies using [15O] oxygen PET suggest that BAT contributes to ~1% of whole body oxygen consumption (39).

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Sex Differences in BAT

In rodents, BAT activity is typically higher in females than males (40). In retrospective studies of humans, the prevalence of BAT is greater in women than men, but the sex difference diminishes with age, suggesting a potential role for the mid-life loss of ovarian hormones in women (31). Conversely, studies of acute cold exposure in humans suggest that the volume of metabolically active BAT and BAT glucose uptake is similar in men and women (41). These seemingly incongruent findings may reflect the differences in the cooling protocols used in these studies and limitations with using static PET/CT 18FDG scans, as discussed as follows.

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The Effects of Exercise on BAT

The effects of exercise on BAT are not well understood. Although there is some evidence that exercise training leads to “browning” of white adipose tissue in mice (42), the physiological relevance of these “brite” or “beige” cells in humans is not clear. There is a paucity of studies that have examined the effects of exercise training on BAT activity in humans. Vosselman et al. (43) compared cold-induced BAT activity measured using 18FDG PET/CT in endurance trained and lean, physically inactive men. Cold-induced BAT 18FDG uptake was lower in the endurance athletes. Furthermore, there were no differences in gene expression markers of classical BAT or beige adipocyte markers in SubQ white adipose tissue. Thus, the limited evidence does not suggest that exercise training increases BAT activity in humans. However, as discussed by Stanford and Goodyear (44), this does not rule out other effects of exercise training on BAT, such as altering BAT secretory proteins, which may have downstream effects on EE or metabolism.

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Evidence for Regulation of BAT by Estrogens

Several lines of evidence suggest that sex hormones regulate BAT activity. For example, androgens and estrogens exert opposing effects in brown adipocytes; E2 increases UCP1 expression and thermogenic activity, whereas testosterone has the opposite effects (45). BAT function is compromised when ER signaling is disrupted (7,46). In addition, OVX rats have decreased BAT thermogenic activity and UCP1 expression in brown adipocytes, but these effects are reversed by E2 treatment (47,48).

We recently performed a pilot study in healthy, premenopausal women (n = 5; 31 ± 6 yr; BMI, 24.3 ± 4.5 kg·m−2) to determine the effects of ovarian suppression on REE, cold-induced thermogenesis (CIT, the increase in EE induced by cold exposure), and BAT activity (49). Participants underwent a 2-hr cooling protocol using a cooling vest filled with water (~16–18°C) before and after 3 months of GnRHAG. After GnRHAG, there were no changes in body composition. However, as in our previous studies, unadjusted REE decreased (~40 kcal·d−1). CIT was also attenuated (350 ± 228 vs 302 ± 139 kcal·d−1). We also performed 18FDG PET/CT measurements to demonstrate the feasibility of measuring BAT activity with this protocol. After GnRHAG, there were no significant changes in mean (3.9 vs 4.0) or maximal (12.3 vs 12.2) standard uptake values (SUV) for 18FDG, but mean and maximal SUV decreased in three of the five women.

There were several limitations in this pilot study. First, the 3-month period of hormone suppression may have been too short to invoke substantial changes in BAT activity, because a single injection of GnRHAG produces an initial sex hormone flare (for several weeks) followed by continuous suppression with additional doses of GnRHAG. Second, there are limitations to using static 18FDG PET/CT scans to study BAT metabolism. As discussed by Blondin et al. (26), this approach yields information about the distribution of glucose during cold exposure but is not a quantifiable index of BAT oxidative metabolism. Third, we used a simple cooling protocol involving a water-filled vest that did not have precise temperature control. Our inability to deliver a consistent and repeatable cooling stimulus may have contributed to variability in the PET/CT measures. Finally, we did not quantify the thermoregulatory response to the cooling protocol (skin temperature, core temperature, and shivering responses using electromyography). As reviewed by Blondin et al. (26), recording and reporting these thermoregulatory responses are vital to critically evaluate the reliability of the cooling method and subsequent BAT metabolic responses.

We are currently following up this pilot study to better characterize the effect of estrogen status on BAT activity. BAT volume and activity are quantified using dynamic PET/CT scanning with 18F-fluorodeoxyglucose and 11C-acetate. The use of 11C-acetate combined with dynamic PET acquisitions provides a quantifiable index of BAT activity based on the mono-exponential decay of the 11C radiotracer (32,33,50,51). We are also delivering the cold exposure using a liquid-conditioned suit to provide a consistent and reproducible cooling stimulus (32–34,50). Importantly, we are performing these studies under both thermoneutral and cold-stimulated conditions. By studying women under thermoneutral conditions, we will be able to determine if BAT activity contributes to differences in REE between pre- and post-menopausal women. Determining the specific role of E2 will be the focus of our future work.

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CONCLUSIONS AND FUTURE DIRECTIONS

Evidence from both animal and human studies indicates that estrogens play an important role in regulating bioenergetics, body composition, and body fat distribution, through postulated mechanisms depicted in our working model (Fig. 7). Our model portends that a decrease in circulating E2 causes reductions in BAT activity and PA. These changes contribute directly to reductions in REE and TEE. A reduction in E2 also leads to increased accumulation of BM-derived adipocytes, predominantly in visceral adipose depots. The adverse metabolic profile of these adipocytes is postulated to contribute to an increase in inflammation and decreased mitochondrial content and activity. Furthermore, if BM-derived adipocytes are characterized by low leptin production, this could exacerbate the decreases in REE and TEE. An important avenue of future research will be to understand whether interventions, such as exercise and menopausal hormone therapy, attenuate the adverse consequences of the loss of E2, by reducing the accumulation of visceral fat and reducing the risk of chronic metabolic diseases.

Figure 7

Figure 7

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Acknowledgments

Supported by NIH grants P50 HD073063, R01 AG018198, R01 DK112260, P30 DK048520, UL1 TR001082, and K01 DK109053. Drs. Kohrt, Klemm, and Melanson are supported also by the Geriatric Research, Education, and Clinical Center at the Denver VA Medical Center, and Drs. Kohrt and Gavin are supported by the University of Colorado Center for Women’s Health Research.

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References

1. Lovejoy JC, Champagne CM, de Jonge L, Xie H, Smith SR. Increased visceral fat and decreased energy expenditure during the menopausal transition. Int. J. Obes. (Lond). 2008; 32(6):949–58.
2. Rogers NH, Perfield JW 2nd, Strissel KJ, Obin MS, Greenberg AS. Reduced energy expenditure and increased inflammation are early events in the development of ovariectomy-induced obesity. Endocrinology. 2009; 150(5):2161–8.
3. Witte MM, Resuehr D, Chandler AR, Mehle AK, Overton JM. Female mice and rats exhibit species-specific metabolic and behavioral responses to ovariectomy. Gen. Comp. Endocrinol. 2010; 166(3):520–8.
4. Revilla R, Revilla M, Villa LF, Cortes J, Arribas I, Rico H. Changes in body composition in women treated with gonadotropin-releasing hormone agonists. Maturitas. 1998; 31(1):63–8.
5. Shea KL, Gavin KM, Melanson EL, et al. Body composition and bone mineral density after ovarian hormone suppression with or without estradiol treatment. Menopause. 2015; 22(10):1045–52.
6. Expert Panel on Detection, Evaluation, and Treatment of High Blood Cholesterol in Adults. Executive Summary of The Third Report of The National Cholesterol Education Program (NCEP) Expert Panel on Detection, Evaluation, And Treatment of High Blood Cholesterol In Adults (Adult Treatment Panel III). JAMA. 2001; 285(19):2486–97.
7. Camporez JP, Jornayvaz FR, Lee HY, et al. Cellular mechanism by which estradiol protects female ovariectomized mice from high-fat diet-induced hepatic and muscle insulin resistance. Endocrinology. 2013; 154(3):1021–8.
8. Torres MJ, Kew KA, Ryan TE, et al. 17beta-estradiol directly lowers mitochondrial membrane microviscosity and improves bioenergetic function in skeletal muscle. Cell Metab. 2018;27(1):167-79.
9. Day DS, Gozansky WS, Van Pelt RE, Schwartz RS, Kohrt WM. Sex hormone suppression reduces resting energy expenditure and {beta}-adrenergic support of resting energy expenditure. J. Clin. Endocrinol. Metab. 2005; 90(6):3312–7.
10. Melanson EL, Gavin KM, Shea KL, et al. Regulation of energy expenditure by estradiol in premenopausal women. J. Appl. Physiol. (1985). 2015; 119(9):975–81.
11. Belchetz PE, Plant TM, Nakai Y, Keogh EJ, Knobil E. Hypophysial responses to continuous and intermittent delivery of hypopthalamic gonadotropin-releasing hormone. Science. 1978; 202(4368):631–3.
12. Melanson EL, Lyden K, Gibbons E, et al. Influence of estradiol status on physical activity in premenopausal women. Med. Sci. Sports Exerc. 2018;50(8):1704-9.
13. Spalding KL, Arner E, Westermark PO, et al. Dynamics of fat cell turnover in humans. Nature. 2008; 453(7196):783–7.
14. Majka SM, Barak Y, Klemm DJ. Concise review: adipocyte origins: weighing the possibilities. Stem Cells. 2011; 29(7):1034–40.
15. Berry R, Rodeheffer MS. Characterization of the adipocyte cellular lineage in vivo. Nat. Cell Biol. 2013; 15(3):302–8.
16. Tran KV, Gealekman O, Frontini A, et al. The vascular endothelium of the adipose tissue gives rise to both white and brown fat cells. Cell Metab. 2012; 15(2):222–9.
17. Sanchez-Gurmaches J, Hung CM, Sparks CA, Tang Y, Li H, Guertin DA. PTEN loss in the Myf5 lineage redistributes body fat and reveals subsets of white adipocytes that arise from Myf5 precursors. Cell Metab. 2012; 16(3):348–62.
18. Crossno JT Jr, Majka SM, Grazia T, Gill RG, Klemm DJ. Rosiglitazone promotes development of a novel adipocyte population from bone marrow-derived circulating progenitor cells. J. Clin. Invest. 2006; 116(12):3220–8.
19. Gavin KM, Gutman JA, Kohrt WM, et al. De novo generation of adipocytes from circulating progenitor cells in mouse and human adipose tissue. FASEB J. 2016; 30(3):1096–108.
20. Ryden M, Uzunel M, Hard JL, et al. Transplanted bone marrow-derived cells contribute to human adipogenesis. Cell Metab. 2015; 22(3):408–17.
21. Majka SM, Fox KE, Psilas JC, et al. De novo generation of white adipocytes from the myeloid lineage via mesenchymal intermediates is age, adipose depot and gender specific. Proc. Natl. Acad. Sci. U. S. A. 2010; 107(33):14781–6.
22. Gavin KM, Majka SM, Kohrt WM, Miller HL, Sullivan TM, Klemm DJ. Hematopoietic-to-mesenchymal transition of adipose tissue macrophages is regulated by integrin beta1 and fabricated fibrin matrices. Adipocyte. 2017; 6(3):234–49.
23. Majka SM, Miller HL, Sullivan T, et al. Adipose lineage specification of bone marrow-derived myeloid cells. Adipocyte. 2012; 1(4):215–29.
24. Gavin KM, Sullivan TM, Kohrt WM, Majka SM, Klemm DJ. Ovarian hormones regulate the production of adipocytes from bone marrow derived cells. Front. Endocrinol. 2018;9:276.
25. Goh J, Goh KP, Abbasi A. Exercise and adipose tissue macrophages: new frontiers in obesity research? Front. Endocrinol. (Lausanne). 2016; 7:65.
26. Blondin B, Labbe SM, Turcotte E, Haman F, Richard D, Carpentier AC. A critical appraisal of brown adipose tissue metabolism in humans. Clin. Lipidol. 2015; 10(3):259–80.
27. Cannon B, Nedergaard J. Brown adipose tissue: function and physiological significance. Physiol. Rev. 2004; 84(1):277–359.
28. Nedergaard J, Bengtsson T, Cannon B. Unexpected evidence for active brown adipose tissue in adult humans. Am. J. Physiol. Endocrinol. Metab. 2007; 293(2):E444–52.
29. van Marken Lichtenbelt WD, Vanhommerig JW, Smulders NM, et al. Cold-activated brown adipose tissue in healthy men. N. Engl. J. Med. 2009; 360(15):1500–8.
30. Virtanen KA, Lidell ME, Orava J, et al. Functional brown adipose tissue in healthy adults. N. Engl. J. Med. 2009; 360(15):1518–25.
31. Ouellet V, Routhier-Labadie A, Bellemare W, et al. Outdoor temperature, age, sex, body mass index, and diabetic status determine the prevalence, mass, and glucose-uptake activity of 18F-FDG-detected BAT in humans. J. Clin. Endocrinol. Metab. 2011; 96(1):192–9.
32. Blondin DP, Labbe SM, Noll C, et al. Selective impairment of glucose but not fatty acid or oxidative metabolism in brown adipose tissue of subjects with type 2 diabetes. Diabetes. 2015; 64(7):2388–97.
33. Blondin DP, Labbe SM, Phoenix S, et al. Contributions of white and brown adipose tissues and skeletal muscles to acute cold-induced metabolic responses in healthy men. J. Physiol. 2015; 593(3):701–14.
34. Ouellet V, Labbe SM, Blondin DP, et al. Brown adipose tissue oxidative metabolism contributes to energy expenditure during acute cold exposure in humans. J. Clin. Invest. 2012; 122(2):545–52.
35. Yoneshiro T, Aita S, Matsushita M, et al. Age-related decrease in cold-activated brown adipose tissue and accumulation of body fat in healthy humans. Obesity (Silver Spring). 2011; 19(9):1755–60.
36. Bartelt A, Heeren J. Adipose tissue browning and metabolic health. Nat. Rev. Endocrinol. 2014; 10(1):24–36.
37. Lee P, Smith S, Linderman J, et al. Temperature-acclimated brown adipose tissue modulates insulin sensitivity in humans. Diabetes. 2014; 63(11):3686–98.
38. van den Berg SM, van Dam AD, Rensen PC, de Winther MP, Lutgens E. Immune modulation of brown(ing) adipose tissue in obesity. Endocr. Rev. 2017; 38(1):46–68.
39. U Din M, Raiko J, Saari T, et al. Human brown adipose tissue [(15)O]O2 PET imaging in the presence and absence of cold stimulus. Eur. J. Nucl. Med. Mol. Imaging. 2016; 43(10):1878–86.
40. Rodriguez-Cuenca S, Pujol E, Justo R, et al. Sex-dependent thermogenesis, differences in mitochondrial morphology and function, and adrenergic response in brown adipose tissue. J. Biol. Chem. 2002; 277(45):42958–63.
41. Chen KY, Brychta RJ, Linderman JD, et al. Brown fat activation mediates cold-induced thermogenesis in adult humans in response to a mild decrease in ambient temperature. J. Clin. Endocrinol. Metab. 2013; 98(7):E1218–23.
42. Bostrom P, Wu J, Jedrychowski MP, et al. A PGC1-alpha-dependent myokine that drives brown-fat-like development of white fat and thermogenesis. Nature. 2012; 481(7382):463–8.
43. Vosselman MJ, Hoeks J, Brans B, et al. Low brown adipose tissue activity in endurance-trained compared with lean sedentary men. Int. J. Obes. (Lond). 2015; 39(12):1696–702.
44. Stanford KI, Goodyear LJ. Exercise regulation of adipose tissue. Adipocyte. 2016; 5(2):153–62.
45. Rodriguez-Cuenca S, Monjo M, Gianotti M, Proenza AM, Roca P. Expression of mitochondrial biogenesis-signaling factors in brown adipocytes is influenced specifically by 17beta-estradiol, testosterone and progesterone. Am. J. Physiol. Endocrinol. Metab. 2007; 292(1):E340–6.
46. Xu Y, Nedungadi TP, Zhu L, et al. Distinct hypothalamic neurons mediate estrogenic effects on energy homeostasis and reproduction. Cell Metab. 2011; 14(4):453–65.
47. Nadal-Casellas A, Proenza AM, Llado I, Gianotti M. Effects of ovariectomy and 17-beta estradiol replacement on rat brown adipose tissue mitochondrial function. Steroids. 2011; 76(10–11):1051–6.
48. Pedersen SB, Bruun JM, Kristensen K, Richelsen B. Regulation of UCP1, UCP2, and UCP3 mRNA expression in brown adipose tissue, white adipose tissue, and skeletal muscle in rats by estrogen. Biochem. Biophys. Res. Commun. 2001; 288(1):191–7.
49. Blondin DP, Swibas T, Wayland L, et al, editors. Brown adipose tissue metabolism in pre- and post-menopausal woman. Keystone Symposia on Molecular and Cell Biology: Bioenergetics and Metabolic Disease. 2018; Keystone, CO.
50. Blondin DP, Labbe SM, Tingelstad HC, et al. Increased brown adipose tissue oxidative capacity in cold-acclimated humans. J. Clin. Endocrinol. Metab. 2014; 99(3):E438–46.
51. Klein LJ, Visser FC, Knaapen P, et al. Carbon-11 acetate as a tracer of myocardial oxygen consumption. Eur. J. Nucl. Med. 2001; 28(5):651–68.
Keywords:

humans; estrogens; resistance exercise; gonadotropin-releasing hormone agonist; adipocytes; brown fat

© 2018 American College of Sports Medicine