Background
The World Health Organization estimates that more than 1.9 billion people are overweight, with 600 million people classified as obese [
1]. Worldwide, the proportion of individuals with a body mass index (BMI) ≥25 kg/m
2, which includes overweight and obese individuals, has risen to encompass 38% of women and 22.6% of girls [
2]. Obesity has been shown to increase the risk of several different types of cancer in women, including postmenopausal breast cancer [
3]. Weight gain following menopause significantly increases breast cancer risk and is correlated with poor prognosis in patients with breast cancer [
4‐
8]. In contrast, obesity in premenopausal women has been shown in multiple studies to mildly reduce the risk of breast cancer [
9‐
11]. The underlying mechanism for this divergence in breast cancer risk before and after menopause is not well-understood.
Mammary epithelium is composed of luminal cells, which express receptors to respond to ovarian hormones, including estrogen receptor alpha (ERα), and basal/myoepithelial cells, which are contractile and contact the basement membrane. Epithelial cells in both the luminal and basal lineages are derived from stem/progenitor cells. These stem/progenitor cells can be enriched in cell fractions that are characterized by their relative expression of cell surface markers, and their activity can be quantified by in vitro assays (for reviews see [
12‐
16]). Current hypotheses suggest that the risk of developing breast cancer during a woman’s lifetime may be determined in part by the number of breast stem/progenitor cells that can serve as targets for transformation [
17]. Recent studies using transgenic mouse models and human tissue xenografts have suggested that the most common types of breast cancer arise from mutations in progenitor cells in the luminal lineage [
18‐
20]. In contrast, cells of the basal/myoepithelial lineage may have a critical tumor suppressive role in preventing the progression of ductal carcinoma in situ (DCIS) to invasive ductal carcinoma (for review, [
21‐
23]). Understanding how obesity alters the complement of stem/progenitor cells within the mammary gland may provide insight into how obesity modulates breast cancer risk.
Data from multiple large epidemiological studies have demonstrated that postmenopausal women have an increased risk of developing ERα
+ luminal breast tumors [
24‐
26]. Postmenopausal women with higher BMI are more likely to develop tumors of the luminal type B molecular subtype, which are characterized by ERα expression, higher proliferative rates and reduced relapse-free survival compared to the luminal A subtype [
27,
28]. However, obese premenopausal and postmenopausal women also have an increased likelihood of being diagnosed with ERα
- tumors compared with lean women [
29,
30]. Here, we investigate changes in the mammary epithelium in both mice and humans to understand how obesity might impact the normal breast epithelium. We show that the timing of obesity and weight loss impact breast epithelial cell populations, which may lead to alterations in lifetime risk of breast cancer.
Methods
Primary human tissue
All human breast tissue procurement for these experiments was in compliance with the laws and institutional guidelines, as approved by the Institutional Review Board (IRB) committee from the University of Wisconsin-Madison. Disease-free, de-identified breast tissues were obtained from patients undergoing elective reduction mammoplasty with informed consent through the Translational Science BioCore BioBank at the Carbone Cancer Center at the University of Wisconsin-Madison. This research study was approved by the IRB as “Not Human Subject Research” with a limited patient dataset including patient age, date of service, and BMI. Tissue samples from patients aged 18–45 years were included in these studies.
Mouse studies
All animal procedures were conducted in accordance with a protocol approved by the University of Wisconsin-Madison Institutional Animal Care and Use Committee. FVB/N (001800) and C57Bl/6 (000664) female mice were purchased from Jackson Laboratories and housed and handled in accordance with the Guide for Care and Use of Laboratory Animals in AAALAC-accredited facilities. All mice were given food and water ad libitum. For obesity studies, 3-week-old or 8-week-old mice were fed either low-fat maintenance chow (5% kcal from fat; Teklad Global #2018) or a high-fat diet (60% kcal from fat; Test Diet #58126) for 16–17 weeks. For weight loss studies, 8-week-old C57Bl/6 female mice were fed the high-fat diet for 15 weeks, then switched to the control diet for 5 weeks. Weight gain was calculated by subtracting the weight on the day of diet initiation from the weight measured each week. One hour prior to euthanasia, mice were injected with 200 mg/kg body weight 5-Bromo-2′-deoxyuridine (BrdU; Fisher Scientific, ICN10017101) diluted in PBS to label cells undergoing DNA synthesis. Thoracic and inguinal mammary glands were collected for analysis. The inguinal glands were utilized for histological assessment and whole mounts, and the remaining glands were collagenase-digested for stem/progenitor assays and flow cytometry.
To assess changes in the estrus cycle in response to obesity, vaginal cytology was performed daily for 14 days prior to tissue collection. Cells from the vaginal epithelium were flushed with 100 μl of PBS onto microscope slides (Fisher Scientific, 22-034979) and stained with Wright’s Stain (RICCA, 9350-16). The stage of the estrus cycle was determined using standard criteria with light microscopy [
31]. Transitional smears were categorized based on the predominant feature on the slide [
31]. Once the vaginal smears were collected, the slides were blinded to the treatment group and evaluated by a veterinary pathologist.
Whole-mount preparation
The inguinal mammary glands were dissected from obese and lean mice and stretched onto microscope slides. The slides were immersed in 10% formalin overnight, defatted in xylenes for 24–48 h, and stained with carmine alum. The whole mounts were dehydrated in graded ethanol and stored in glycerol before analysis. Mammary branching was quantified by identifying branch points from the primary ducts originating from the nipple.
Mammary epithelial cell isolation
Mouse mammary glands and human breast tissue were digested in organoid medium composed of DMEM/Ham’s F-12 (Corning, 10-090-CV) supplemented with 5 ng/ml human epidermal growth factor (EGF; Sigma, E9644), 10 μg/ml insulin (Sigma, I0516), 0.5 μg/ml hydrocortisone (Sigma, H0888), 5% calf serum, and 1% antibiotic/antimycotic solution (Corning, 30-004-CI) with 3 mg/ml collagenase A (Roche, 11088793001) and 100 U/ml hyaluronidase (Sigma, H3506) at 37 °C for either 1 hour (mouse) or overnight (human). Clusters of mammary epithelial cells, called organoids were allowed to settle at room temperature for 20 min, followed by centrifugation for 5 min at 8 × g relative centrifugal force. The supernatant was removed, the contaminating red blood cells lysed (ACK Lysing Buffer, Lonza, 10-548E), and the mammary organoids were washed with 5% calf serum in PBS. Human breast organoids were aliquoted and cryopreserved for future analysis. Mouse mammary organoids were dissociated to single cells following digestion for analysis in progenitor assays.
To isolate epithelial cells, mammary organoids were trypsinized by pipetting in 0.25% Trysin/EDTA (Corning, 25-053-CI). Cells were resuspended in organoid medium with 0.1 mg/ml DNAse (Roche 10104159001) and 5 mg/ml Dispase II (Roche, 04942078001) for 5 min. Single cells were isolated by filtration through a 40-μm cell strainer. Viable cells were counted using Trypan Blue (Corning, 25-900-CI) exclusion prior to further analysis.
Flow cytometry
Flow cytometry of both human and mouse epithelial cells was performed and analyzed according to published guidelines [
32]. Briefly, primary human breast tissue was dissociated to single cells, and endothelial, lymphocytic, monocytic, and fibroblastic lineages were depleted using antibodies to CD31 (MS-353), CD34 (180227), and CD45 (180367) from Thermo Scientific, fibroblast-specific protein IB10 (Sigma, F4771), and pan-mouse IgG Dynabeads (Dynal; Invitrogen, 115.31) according to the manufacturers’ instructions and as described previously [
33]. Lineage-depleted single-cell suspensions or cells from immunomagnetic bead-sorted populations were resuspended at 1 × 10
6 cells/ml in PBS containing 1% calf serum and bound with fluorescently conjugated antibodies to human EpCAM (12 ng/μl; APC; BD Biosciences, 347200) and CD49f (0.5 μg/μl; fluorescein isothiocyanate (FITC); BD Biosciences 555735). Primary mouse mammary epithelial cells were incubated with antibodies to detect CD49f (0.2 μg/μl; PerCP cy5.5; Biolegend, 313618) and EpCAM (0.2 μg/μl; APC; Biolegend, 118213). Endothelial, lymphocytic and monocytic lineages were depleted using antibodies specific to mouse CD31 (0.4 μg/μl; phycoerythrin (PE), 12-0311-83), CD45 (0.5 μg/μl; PE, 12-0451-81) and Ter119 (0.4 μg/μl; PE, 12-5921) from eBiosciences. All cells were incubated with primary antibodies for 20 min at 4 °C. To assess cell viability, all cells were stained with fixable viability dye eFluor 780 (eBiosciences, 65-0865) per manufacturers’ instructions. Antibody-bound cells were washed and resuspended at 1 × 10
6 cells/ml and run on a BD LSR Fortessa (BD Biosciences). Gates were set on fluorescence minus one controls. Flow cytometry data were analyzed with the FlowJo software package (TreeStar).
Progenitor assays
To assess mammosphere forming ability, 40,000 human mammary epithelial cells were plated in triplicate on 6-well ultra-low-attachment plates (Corning, CLS3473) in MEGM media (Lonza, cc-4136). For mice, 4000 mammary epithelial cells were plated in triplicate on 24-well ultra-low-attachment plates (Corning, CLS3473-24EA) in 500 μl EpiCult-B complete media (STEMCELL Technologies, 05610). Mammospheres formed for 7 days at 37 °C with 5% CO2 and were counted and imaged on an inverted microscope. To assess second-generation mammosphere formation, primary mammospheres were collected, trypsinized, and re-plated onto ultra-low-attachment plates for 7 days. Primary and secondary mammospheres were counted from 5 − 8 mice in each treatment group in triplicate.
To assess 3D colony formation, 40 μl of 1 mg/ml Type I rat tail collagen (Corning, 354236) was polymerized in 8-well chamber slides (Falcon, 354108); 1000 mouse mammary epithelial cells were resuspended in 500 μl organoid media containing 2% Matrigel (Corning, 354234) and layered over the collagen in duplicate. Colonies were formed for 7 days at 37 °C with 5% CO2 and were counted and imaged on an inverted microscope. Round and branching colonies were identified from five mice in each treatment group and quantified based on morphology.
To assess colony forming ability on tissue culture plates, viable cells from single-cell suspensions of lineage-depleted human mammary epithelial cells were plated at 10,000 cells/mL in 6-well plates in triplicate for adherent growth in MEGM (Lonza, cc-4136). Colonies were allowed to form for 7 days, after which non-adherent spheres (floating colonies) from adherent cultures were collected for analysis. Floating colonies were pelleted at 150 × g for 5 min and resuspended in 500 μl of growth medium and counted with an inverted light microscope.
To assess mouse mammary epithelial cell colony forming ability, 5000 epithelial cells were plated on 35-mm tissue culture plates (Cellstar, 627160) containing a feeder layer of 100,000 NIH 3 T3 cells (generously provided by Dr. Linda Schuler) treated with 20 mg/ml bleomycin sulfate (Millipore Calbiochem, 20-340) for 30 min. Epithelial cells were cultured in 2.5 ml of EpiCult-B medium supplemented with proliferation supplements (STEMCELL Technologies, 05610), 4 μg/ml heparin (STEMCELL Technologies, 07980), 0.02 μg/ml recombinant human basic fibroblast growth factor (STEMCELL Technologies, 02634), 0.01 μg/ml hEGF (Sigma, E9644) and 5% fetal bovine serum. After 24 h, the medium was changed to serum-free stem cell medium. Colonies were formed in duplicate for 7 days at 37 °C with 5% CO2.
Following colony formation, the plates were fixed with ice-cold methanol for 10 min and stored at -20 °C prior to analysis. To assess luminal and basal lineage markers, plates were immunohistochemically co-labeled for cytokeratin 8 (CK8) and CK14. Briefly, cells were permeabilized with 0.1% Triton X for 10 min and blocked for non-specific antibody binding using 1% BSA in PBS for 1 h. Cells were incubated with CK8 (Thermo Scientific, RM-2107-S0) at a 1:150 dilution in 1% BSA in PBS, followed by 30-min incubation with biotinylated goat anti-rabbit IgG (Vector, BA-1000) at 1:200 dilution in 1% BSA in PBS. Staining was visualized using VECTASTAIN ABC horseradish peroxidase (HRP) Kit (Vector, PK-4000) and VECTOR VIP Peroxidase (Vector, SK-4600). Cells were blocked with the avidin/biotin blocking kit (Vector, SP-2001). Cells were stained for CK14 (Thermo Scientific, MA5-11599) at a 1:150 dilution utilizing the MOM Kit (Vector BMK-2202) according to package instructions. CK14 staining was developed with 3,3-diaminobenzidine (DAB) Peroxidase (HRP) Substrate Kit (Vector, SK-4100). Colonies were counted from five to eight mice in each treatment group in duplicate using a light microscope.
Immunohistochemical assessment
Mammary glands and breast tissue were fixed in 10% neutral buffered formalin for 18–24 h, embedded in paraffin and sectioned at 5 μm. Paraffin-embedded sections were rehydrated in graded alcohol and incubated with 0.5% H2O2 for 20 min. Sections underwent antigen retrieval in 0.1 M tris buffer (pH = 9.0) at 95 °C for 20 min. The tissue sections were blocked in 1% gelatin in TBST for 1 h and incubated with anti-mouse ERα antibodies (clone MC-20; Santa Cruz Biotechnology, sc-542) at a 1:500 dilution in 1% gelatin in TBST overnight at 4 °C. The sections were incubated with biotinylated goat anti-rabbit IgG secondary antibodies (Vector, BA-1000) at 1:500 dilution in 1% gelatin for 30 min, followed by VECTASTAIN ABC HRP Kit (Vector, PK-4000) for 30 min. The sections were developed with DAB Peroxidase (HRP) Substrate Kit (Vector, SK-4100), counterstained in hematoxylin, dehydrated in graded alcohol and mounted with Permount. A Nikon Eclipse 80 t microscope and SPOT camera were used for imaging the stained sections. The percentage of ERα+ cells were quantified by dividing the total number of ERα+ cells by the total number of epithelial cells within the ducts and multiplying by 100. Five images of each gland from five mice in each treatment group were analyzed.
Immunofluorescence
Paraffin-embedded sections were rehydrated in graded alcohol and underwent antigen retrieval in 0.1 M tris buffer (pH = 9.0) or 0.01 M citrate buffer (pH = 6.0) at 95 °C for 20 min. Sections were blocked for 1 h and incubated with primary antibodies at 4 °C overnight. Primary antibodies included anti-mouse ERα (1:50, clone MC-20; Santa Cruz Biotechnology, sc-8002), anti-human ERα (1:200, clone F-10; Santa Cruz Biotechnology, sc-8002), anti-human CK14 (1:500; ThermoScientific, RB-9020), anti-human p63 that recognizes the C-terminal region of the protein (1:250, Santa Cruz Biotechnology, sc-6243), alpha smooth muscle actin (SMA, 1:5,000; Sigma, A5228), Ki67 (1:250; Abcam, 15580), or BrdU (clone BU1/75, 1:40; Accurate Chemical and Scientific Corporation, OBT0030S). Tissue sections were then incubated with secondary antibodies diluted 1:250 in 5% BSA in PBS for 30 min. Secondary antibodies were obtained from Life Technologies and included Alexa Fluor 546 goat anti-rabbit IgG (A11010), Alexa Fluor 488 goat anti-mouse IgG (A11001) and Alexa Fluor 488 goat anti-rat IgG (A11006). Sections were counterstained with 250 ng/ml 4′,6-diamidine-2′-phenylindole dihydrochloride (DAPI) for 5 min and mounted with the Slow-Fade mounting kit (Life Technologies, S2828).
The sections were imaged using a fluorescent microscope and the program NIS Elements BR 3.2. For basal/myoepithelial cell stains, including αSMA, p63, and CK14, sections were scored based on continuity of stain surrounding the luminal epithelial cells. The following score was used: 4 = 100%, 3 = 99–75%, 2 = 74–50%, and 1 = <50%. Five images were scored per tissue section. For mouse ERα and BrdU labeled samples, five images were quantified from seven mice per group. For human ERα and Ki67 labeling, ERα
+, Ki67
+, and co-labeled cells were quantified from three images per tissue sample. All cell counting and image alignment was performed using Image J software. To examine ERα expression in breast tissue from postmenopausal women, we utilized an available dataset of previously published data [
34] from normal postmenopausal breast tissue from the Susan G. Komen Tissue Bank. We analyzed the correlation between ERα expression that was quantified as described [
34] and the patients’ recorded BMI values.
Statistical analyses
Results were expressed as the mean ± standard deviation (s.d.), unless stated otherwise. Statistical differences were determined using the Mann-Whitney test, unless stated otherwise. Correlation with BMI in humans was analyzed using two-tailed Pearson correlation. Investigators were blinded to the BMI data until analyses were complete. P values of 0.05 or less were considered to denote significance. Statistical analyses were conducted using GraphPad Prism 6 (GraphPad Software).
Discussion
Obesity has divergent effects on breast cancer risk, depending on whether weight gain occurs early in life or following menopause. To understand how obesity alters normal breast tissue, potentially leading to increased risk of breast cancer, we examined the consequences of obesity in a well-characterized HFD mouse model and in human breast tissue samples from reduction mammoplasty surgery. Using these tissues, we identified global changes in both human and mouse epithelial cell populations and in mammary gland architecture that might lead to the observed changes in breast cancer risk over time.
Breast cancer can be divided into distinct subtypes based on gene expression profiling [
62‐
64]. These divergent subtypes have been hypothesized to arise due to differences in mutations and distinct cells of origin within the breast (for review see [
16,
65,
66]). Studies using targeted expression of oncogenes in the mammary epithelium have demonstrated that luminal lineage cells generate tumors that are more aggressive and heterogeneous than epithelial cells from the basal lineage [
20,
67,
68], leading to the hypothesis that luminal progenitor cells are the cells of origin for the most common types of breast cancer [
18,
19]. If breast cancers originate in distinct stem/progenitor cell populations, it also suggests that the risk of cancer development may be related to the size of the progenitor cell pool and its mitotic activity [
17]. Our studies show that obesity significantly enhances luminal cells in mice and mature luminal and luminal progenitor cells in women. While postmenopausal women have an increased risk of developing ERα
+ luminal breast cancers [
24‐
26], both premenopausal and postmenopausal obese women also have an increased likelihood of being diagnosed with ERα
- tumors compared with lean women [
29,
30]. These results suggest that obesity may enhance the risk of development of different subtypes of breast cancer through the expansion of luminal progenitor cells that may give rise to the most common types of breast cancer. Studies utilizing lineage tracing in the context of obesity will be necessary to more directly assess how changes in epithelial cell populations contribute to the formation of different tumor histological types.
Epidemiologic studies have suggested that increased ERα expression in breast epithelial cells increases breast cancer risk, particularly in postmenopausal women [
69‐
71]. Our studies suggest that obesity enhances ERα
+ cells in obese mice and in the breast tissue of premenopausal and postmenopausal women. Interestingly, while weight gain in premenopausal women mildly decreases breast cancer risk before menopause [
9,
10], premenopausal weight gain enhances the risk of development of ERα
+ breast tumors after menopause [
72]. The cells of origin for ERα
+ breast tumors are currently unclear, but tumors may originate in ERα
+ progenitor cells. In both mice and humans, rare ERα
+ cells have been identified that co-label with markers of proliferation [
58‐
60,
73]. These ERα
+ proliferating cells are rare in breast tissue from young women, but are enhanced with age [
74,
75]. Our studies suggest that obesity also enhances ERα
+ cells that co-label with markers of proliferation in both mice and humans. ERα
+ epithelial cell populations have been shown to have limited progenitor activity within the mouse mammary gland [
38,
76,
77]. Recent lineage tracing studies have demonstrated that ERα
+ progenitor cells maintain the ERα
+ luminal cell lineage [
76,
78]. Examination of ERα
+ progenitor cells in the context of obesity using these lineage tracing models could provide insight into how obesity alters ERα
+ progenitor cells and their contribution to ERα
+ breast tumors.
Progression from ductal carcinoma in situ (DCIS) to invasive ductal carcinoma is characterized by invasion of breast cancer cells into the surrounding tissue due to the physical breach of the basal/myoepithelial cell layer and underlying basement membrane [
79]. Our studies have shown that within mammary glands from obese mice, basal/myoepithelial epithelial cell populations are reduced, and SMA
+ basal/myoepithelial cells discontinuously surround luminal cells, compared with control mice. Given their central role in suppression of tumor progression and invasion (for review see [
21‐
23]), discontinuity of the basal/myoepithelial layer under conditions of obesity may weaken local defenses, which could facilitate tumor invasion into the surrounding tissue once cancer arises. Obesity increases the risk of the development of invasive breast cancer in postmenopausal women [
80], and loss of the protective basal/myoepithelial layer may contribute to this risk by allowing more rapid progression of DCIS to invasive breast tumors. Our results suggest that disease models to examine the impact of obesity-induced basal/myoepithelial cell loss on the progression of DCIS to invasive ductal carcinoma may be best addressed using transgenic mouse models of tumor progression or intraductal transplantation in obese mice to model human DCIS [
81,
82].
Although postmenopausal obesity is a significant risk factor for breast cancer, epidemiological studies suggest that obesity during puberty decreases life-long breast cancer risk [
10,
72,
83], and decreases the risk of developing benign breast disease [
84]. Puberty is an important stage of growth in the normal mammary gland, with the elongation of ducts and secondary branches forming from terminal end buds to fill the mammary fat pad. When 3-week-old mice were fed a HFD, we observed significant reductions in ductal branching. Our results are consistent with a previous study demonstrating significant reductions in terminal end buds and ductal elongation in BALB/C and C57Bl/6 mice fed a HFD for 4 weeks [
85]. Together, these studies suggest that obesity during puberty alters the architecture of the mammary gland, leading to reduced ductal development. Reduced mammary epithelial development may decrease breast cancer risk over a woman’s lifetime by limiting the total number of epithelial stem/progenitor cells that could accrue mutations and form malignant growths. However, the effects of obesity on breast development in the context of obesity during puberty have not been examined in humans, and how these changes might contribute to the decreased breast cancer risk in premenopausal women observed epidemiologically are unknown. Obesity has also been associated with alterations in the menstrual cycle and decreased ovulation in young women (for review see [
86]). We did not observe significant differences in the length of the estrus cycle in lean and obese mice or significant changes in uterine weight, which might suggest imbalances in ovarian hormones. However, more subtle differences in the estrus cycle may be present, which we were unable to detect.
Although we have demonstrated that obesity leads to alterations in mammary epithelial cell populations and enhanced ERα expression, the mechanism underlying these observations may be complex. Leptin, which is secreted by adipocytes, is increased under conditions of obesity and has been shown to enhance ERα expression in both normal epithelial cells and breast cancer cell lines [
87,
88]. Leptin has also been shown to stimulate breast stem cell self-renewal in mammosphere assays utilizing primary human breast epithelial cells [
89]. In addition to enhanced leptin secretion, obesity increases the recruitment of inflammatory macrophages into mammary adipose tissue [
35,
37,
90]. Chronic inflammation has been shown to result in increased aromatase expression within mammary tissue [
90], and increased expression of cytokines, such as interleukin (IL)-6 and IL-8, which have been shown to regulate breast cancer stem cells [
91]. Obesity may also alter transforming growth factor beta (TGFβ) release and bioavailability to mammary epithelial cells [
92,
93]. TGFβ signaling inhibits the proliferation of ERα
+ epithelial cells in mouse mammary glands [
94,
95] and cultured human breast epithelial cells [
96], and loss of TGFβ signaling may play a role in the observed increases in ERα
+ proliferating cells in mammary tissue in obesity. These signaling pathways may act in concert or have divergent effects on separate epithelial cell populations within the luminal and basal/myoepithelial mammary epithelium.
Weight loss is an attractive intervention for reduction of breast cancer risk. Data from the Prospective Nurses’ Health Study suggests that weight loss after menopause is associated with lower incidence of breast cancer [
97]. Additionally, a group of women who lost weight with bariatric surgery were shown to have reduced risk of breast cancer [
98]. Our results suggest that in mice that lost weight, the percentage of ERα
+ cells in normal mammary epithelium was similar to that in control mice. Additionally, there were no significant differences observed in any in vitro progenitor assays. These results suggest that weight loss may reverse the enrichment of progenitor cells induced by obesity, potentially decreasing the cells of origin for breast cancer to similar levels found in lean mice. We also observed increased continuity of the SMA
+ basal/myoepithelial layer, which also suggests that the protective layer surrounding the luminal cells is regained with weight loss. Recent studies in postmenopausal obese women and in obese and overweight breast cancer survivors following treatment suggest that weight loss decreases serum levels of leptin, insulin, and estrogens [
99,
100], and Ki67 expression in normal breast epithelial cells [
100]. These results suggest that weight loss may reduce serum levels of growth factors that may influence progenitor activity within the breast epithelium. Although we observed reductions in progenitor activity with weight loss in mice, another study examining epigenetic changes in the mammary gland in obesity demonstrated that weight normalization did not reverse the effects of obesity on epigenetic reprogramming and inflammatory signals in the mammary gland microenvironment [
101]. Further studies are necessary to understand how the timing and use of weight loss strategies may have maximal impact for the reduction of breast cancer risk in obese women.