Positive Regulation of Estrogen Receptor Alpha in Breast Tumorigenesis

Abstract

Estrogen receptor alpha (ERα, NR3A1) contributes through its expression in different tissues to a spectrum of physiological processes, including reproductive system development and physiology, bone mass maintenance, as well as cardiovascular and central nervous system functions. It is also one of the main drivers of tumorigenesis in breast and uterine cancer and can be targeted by several types of hormonal therapies. ERα is expressed in a subset of luminal cells corresponding to less than 10% of normal mammary epithelial cells and in over 70% of breast tumors (ER+ tumors), but the basis for its selective expression in normal or cancer tissues remains incompletely understood. The mapping of alternative promoters and regulatory elements has delineated the complex genomic structure of the ESR1 gene and shed light on the mechanistic basis for the tissue-specific regulation of ESR1 expression. However, much remains to be uncovered to better understand how ESR1 expression is regulated in breast cancer. This review recapitulates the current body of knowledge on the structure of the ESR1 gene and the complex mechanisms controlling its expression in breast tumors. In particular, we discuss the impact of genetic alterations, chromatin modifications, and enhanced expression of other luminal transcription regulators on ESR1 expression in tumor cells.

Full text

cells
Review
Positive Regulation of Estrogen Receptor Alpha in Breast
Tumorigenesis
Lucas Porras, Houssam Ismail and Sylvie Mader *


Citation: Porras, L.; Ismail, H.;
Mader, S. Positive Regulation of
Estrogen Receptor Alpha in Breast
Tumorigenesis. Cells 2021,10, 2966.
https://doi.org/10.3390/
cells10112966
Academic Editors: Stephen Yarwood
and Tuula Kallunki
Received: 16 August 2021
Accepted: 24 October 2021
Published: 31 October 2021
Publisher’s Note: MDPI stays neutral
with regard to jurisdictional claims in
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iations.
Copyright: © 2021 by the authors.
Licensee MDPI, Basel, Switzerland.
This article is an open access article
distributed under the terms and
conditions of the Creative Commons
Attribution (CC BY) license (https://
creativecommons.org/licenses/by/
4.0/).
Institute for Research in Immunology and Cancer, Universitéde Montréal, Montreal, QC H3T 1J4, Canada;
lucas.porras@umontreal.ca (L.P.); houssam.ismail@umontreal.ca (H.I.)
*Correspondence: sylvie.mader@umontreal.ca
Abstract:
Estrogen receptor alpha (ER
α
, NR3A1) contributes through its expression in different
tissues to a spectrum of physiological processes, including reproductive system development and
physiology, bone mass maintenance, as well as cardiovascular and central nervous system functions.
It is also one of the main drivers of tumorigenesis in breast and uterine cancer and can be targeted by
several types of hormonal therapies. ER
α
is expressed in a subset of luminal cells corresponding to
less than 10% of normal mammary epithelial cells and in over 70% of breast tumors (ER+ tumors),
but the basis for its selective expression in normal or cancer tissues remains incompletely understood.
The mapping of alternative promoters and regulatory elements has delineated the complex genomic
structure of the ESR1 gene and shed light on the mechanistic basis for the tissue-specific regulation
of ESR1 expression. However, much remains to be uncovered to better understand how ESR1
expression is regulated in breast cancer. This review recapitulates the current body of knowledge
on the structure of the ESR1 gene and the complex mechanisms controlling its expression in breast
tumors. In particular, we discuss the impact of genetic alterations, chromatin modifications, and
enhanced expression of other luminal transcription regulators on ESR1 expression in tumor cells.
Keywords:
breast cancer; estrogen receptor alpha; luminal breast cancer; mammary gland; ESR1;
FOXA1;GATA3
1. Introduction
Breast cancer is the most frequent malignancy in women worldwide, with outcomes
strongly affected by the stage of the disease [
1
]. It is characterized by the unregulated
proliferation of epithelial cells normally located in the mammary ducts or alveoli. However,
immunohistochemical (IHC) analysis reveals that breast cancer is a heterogeneous disease
as evidenced by the differential detection of two main targetable tumorigenic drivers:
estrogen receptor alpha (ER
α
), a nuclear receptor, and ERBB2/HER2, a membrane receptor
belonging to the ERBB1-4 growth factor receptor family whose gene is frequently amplified
in breast cancer. Use of positivity thresholds for detection of each marker by IHC (and for
detection of ERBB2/HER2 amplification by fluorescence in situ hybridization) identifies
four tumor subtypes, ER+HER2
−
, ER+HER2+, ER
−
HER2+ and ER
−
HER2
−
[
2
]. The
expression of a third marker, the progesterone receptor (PR, NR3C3), an ER
α
target gene,
reflects active estrogenic signaling and is a marker of improved prognosis compared to
PR-negative tumors [
3
]. ER+ and/or PR+ tumors represent more than two-thirds of breast
cancer cases and are currently treated with hormonal therapies that most commonly include
aromatase inhibitors, which suppress endogenous estrogen production, or antiestrogens,
which compete with estrogens for binding to estrogen receptors and inactivate them by
inducing alternative conformations of their ligand-binding domains [
4
,
5
]. On the other
hand, primary tumors with undetectable ER expression levels and activity are intrinsically
resistant to hormonal therapies. These include ER-HER2+ tumors, and also tumors negative
for ER, PR and HER2, called Triple Negative Breast Cancer (TNBC). It is estimated that
approximately 30–50% of patients relapse after one or several lines of endocrine treatment.
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Cells 2021,10, 2966 2 of 25
Expression of ER
α
is maintained in most relapsed tumors, which can remain sensitive
to different hormonal therapy agents. However, about 15–40% of relapsed tumors lose
expression of ER
α
and are therefore insensitive to additional hormonal treatment [
6
–
8
]. This
review will discuss specifically the mechanism controlling ER
α
expression levels in normal
and breast cancer cells. Other mechanisms of resistance to hormonal therapies, which
include activation of other proliferative pathways and modulation of ER
α
or coregulator
activity via genetic or epigenetic alterations, have been reviewed elsewhere (e.g., [
7
,
9
–
11
]).
ER
α
and its paralog estrogen receptor beta (ER
β
), encoded by the estrogen
receptor 1
(ESR1) and the estrogen receptor 2 (ESR2) genes, respectively [
12
] (Figure 1a), mediate the
systemic physiological effects of circulating estrogens [
13
]. ER
α
can be found in three major
protein isoforms differing in molecular weight, ER
α
-66, ER
α
-46 and ER
α
-36
(Figure 1b)
,
while ER
β
has five isoforms, ER
β
1-5. These receptors are part of the broader nuclear
receptor family, composed of transcription factors whose activity can be regulated by small
ligands and/or post-translational modifications [
14
]. Human ERs have only 44% overall
identity in their primary structures but contain a well conserved DNA binding domain
(96% identity between human ER
α
/
β
, Figure 1a) [
15
] composed of two zinc fingers folding
into a single structural entity that directs binding of ER dimers to palindromic estrogen
response elements (EREs) [
16
–
18
]. ER binding sites can be found at promoters of estrogen
target genes but also at enhancers situated in intergenic or intronic sequences and having
long-range impacts on gene transcription [
19
–
23
]. However, only a minority of predicted
binding sites are used under any given experimental conditions [
24
], occupancy being
controlled by chromatin accessibility [
25
] and/or cooperativity with other transcription
factors, some of them acting as pioneer factors to enable access of ERs to cognate binding
sites [
26
]. In addition to direct binding to EREs, ERs can also be recruited onto DNA
by tethering via interactions with other transcription factors. Ligand addition leads to
increased association of ER
α
with DNA and assembly of clusters of ER
α
-bound enhancers,
or super-enhancers, organized around “mother enhancers” that are enriched in strong EREs
and are detected in estrogen-deprived cells. By contrast, “daughter enhancers” appearing
near mother enhancers upon ligand stimulation are enriched in sites for cooperating factors,
suggestive of protein–protein interactions driven by the increased local concentrations of
ERs on DNA [
27
]. Some enhancers are “hotspots” of colocalization of many transcription
factors, which may be assembled in a “MegaTrans” complex marking the most active
enhancers [
28
–
30
]. In particular, interaction with CTCF and components of the cohesin
complex, which play roles in chromatin loop formation [
31
], has been described as a
hall-mark of “hub” enhancers driving local chromatin conformation within topologically
associated domains (TADs) [32].
The ligand binding domains of ERs are also conserved, although to a lower degree
(59% identity between human ER
α
/
β
) [
15
], which has made it possible to design receptor-
specific synthetic ligands [
33
]. Both receptors are nevertheless activated by circulating
estrogens, which trigger conformational changes in the ligand binding domain inducing
receptor binding to DNA and recruitment of a variety of transcriptional coactivators or
co-repressors [12,15,34–38]. The two receptors diverge more in other domains (Figure 1a),
and as a result differ in their functional properties [
39
] and in their regulation by post-
translational modifications (please refer to Phosphosite.org for a summary of ER
α
/
β
post-
translational modifications and to [
11
,
40
,
41
] for reviews). Modifications can affect diverse
ER functional properties, including subcellular localization, DNA binding, interaction
with other transcriptional regulators or chromatin components, and protein stability. Post-
translational modifications include phosphorylation by diverse kinases at multiple sites in
a manner regulated by ligand binding. In addition, ERs can be acetylated, methylated and
SUMOylated in a ligand-modulated manner. In addition, ubiquitination can be induced
by agonist binding but also by some antiestrogens, such as fulvestrant, the prototype of
Selective Estrogen Receptor Downregulators (SERDs), resulting in enhanced degradation
of ER
α
by the proteasome machinery [
42
]. Finally, palmitoylation regulates localization of
a pool of ER
α
at the plasma membrane, where it initiates non-genomic estrogenic signaling

Cells 2021,10, 2966 3 of 25
that can cross-talk with the nuclear pool of ER
α
[
43
]; mouse models with a mutation
abolishing palmitoylation support a tissue-specific contribution of membrane signaling in
estrogen physiology, while lack of nuclear ERαabrogates most responses [34,39].
Figure 1.
(
a
). ER
α
and ER
β
have a conserved gene and protein organization. Amino acid identity between human ER
α
and ER
β
is indicated for each domain [
12
]. The most frequent ER
α
mutations found in relapsed ER-positive tumors are
indicated. (b). The main ERαprotein isoforms and their corresponding mRNA structures are shown.

Cells 2021,10, 2966 4 of 25
ER
α
is the main tumorigenesis driver in breast cancer, the roles of the ER
β
isoforms
being less clear, although it was shown that they can repress the activity of ER
α
[
44
].
The human ESR1 gene, located at the q25.1-q25.2 locus of chromosome 6, has a complex
structure. A cDNA was first cloned from MCF-7 cells in 1985 and was found to result from
the transcription of eight exons, which together with introns span 296 kb [
45
–
48
]. However,
characterization of transcripts in different estrogen-responsive tissues has revealed that
the overall gene unit spreads over ~447 kb, comprising several alternative promoters and
non-coding exons. Various transcriptional regulators bound to its distinct promoters or
to enhancers acting at a large distance via the formation of chromatin loops control ESR1
gene expression during the development, normal physiology and tumorigenesis of the
mammary tissue and in other estrogen-responsive tissues [
49
]. In this review, we will focus
on the mechanisms known to control ESR1 expression, with a focus on positive regulation
by transcription factors expressed in ER+ breast cancer, also called luminal breast cancer.
2. Expression of ESR1 Is Transcriptionally Regulated in Normal Mammary Tissue and
in Breast Tumorigenesis
2.1. Expression in Normal Tissue
The mammary gland, an organ uniquely found in mammals, is composed of an
epithelial and a stromal compartment [
50
]. The first stage of mammary tissue development
occurs during mammalian embryogenesis and leads to the formation of a rudimentary
ductal tree connected to the nipple [
50
,
51
]. During puberty, the mammary tissue will
further develop to form an expanded tree of epithelial ducts ending in terminal end
buds, embedded in the mammary fat pad. Throughout pregnancy, the ductal tree further
expands and lactogenic differentiation of luminal cells in alveoli enables milk production
for lactation [51].
In normal human breast tissue, ESR1 expression is restricted to the epithelial com-
partment, where ER
α
is detected in only about 7–10% of cells [
52
,
53
]. ER
α
is absent in
differentiated myoepithelial cells and present only in a fraction of luminal cells in the hu-
man mammary gland where its expression levels are variable throughout life, particularly
following puberty, during menstrual cycles and throughout pregnancy and lactation [
54
,
55
].
During mouse mammary gland development, ductal growth is initiated at terminal end
buds (TEBs); ER
α
is not expressed in cap cells, located around the leading tip of the TEB,
but is detected in a fraction of the underlying body cells, which also express luminal
markers keratin 8/18 [
51
,
56
]. In mice, expression of the Esr1 gene is necessary for the devel-
opment and differentiation of the mammary gland, which is under control by estrogens and
progesterone [
57
–
59
]. Indeed, suppression of Esr1 expression disrupts the morphogenesis
of the mammary gland and the function of the entire reproductive system [59,60].
The synthesis of sex hormones in the ovaries is regulated throughout the hormonal
cycle after puberty and before menopause. While estrogens are produced during the entire
hormonal cycle with a peak between the follicular phase and ovulation, progesterone is
produced mostly during the second half (luteal phase). Increased expression of the ESR1
gene in epithelial cells during the follicular phase correlates with subsequent increased
mitotic activity of epithelial cells in breast lobules [
61
–
63
]. However, ER
α
expression in
individual normal epithelial cells does not correlate with proliferation (KI67 positivity, triti-
ated thymidine incorporation) [
64
], and it has been shown that paracrine signaling via ER
α
stimulates mammary epithelial cell proliferation during mammary morphogenesis [
65
].
However, studies in rodents and rhesus monkey have indicated that ER
α
protein expres-
sion is down-regulated by estrogens, which may explain the apparent lack of association
of ERα-expressing cells with proliferative markers [66,67]. In breast cancer cell lines, ERα
drives cell proliferation by activating expression of proliferative genes, including CCND1,
MYC,E2Fs and FOXM1 [68–71]. In ER+ MCF-7 breast cancer cells, ERαprotein levels are
highest in the S and G2/M phases of the cell cycle [
72
]. The proliferative action of estrogens
in tumors as well as cell lines is confirmed by the observation that hormonal therapies
suppress proliferation of breast tumor cells [73].

Cells 2021,10, 2966 5 of 25
2.2. Expression in Tumoral Tissue
In the clinic, breast tumors have been deemed ER
α
-positive (ER+ tumors) if more than
1% of epithelial cells demonstrate nuclear staining in immunohistochemistry with anti-
hER
α
antibodies, qualifying patients for hormonal therapies, although rates of response
are much lower for tumors with low expression of ER
α
[
74
]. While there is a range of
ER
α
expression patterns between tumors, expression levels of the ER
α
protein are often
higher and more homogeneous than in normal tissue, especially in the LumA subgroup.
Nevertheless, some tumors exhibit heterogeneous expression, with variable proportions of
cells expressing ERαwhile staining is faint or absent in others.
Elevated ESR1 expression has been observed in atypical ductal hyperplasia and
in situ carcinoma, suggesting that deregulation of the ESR1 expression level may be
implicated in early pathogenic changes during breast tumorigenesis [
62
,
75
]. Whether
ESR1 overexpression is sufficient to drive tumorigenesis in the human mammary gland
is unclear. Interestingly, transgenic expression of wild type (wt) murine ER
α
in mice
using an
MMTV-tTA
controlled expression system led to ductal hyperplasia and ductal
carcinoma in situ by four months of age and to a low incidence of invasive carcinoma by
12 months [
76
,
77
]. However, it remains unclear whether deregulated ESR1 expression is
sufficient to drive tumorigenesis in the human mammary gland.
The ESR1 gene is affected by copy number variations, with amplification events
initially reported at a high frequency (20%), correlating with higher ER
α
levels [
78
]. On
the other hand, Brown et al. found amplification in only 1% of tumors [
79
]. Amplification
is predicted by GISTIC in less than 4% of tumors in the TCGA Firehose Legacy dataset
(Figure 2) and at similar low frequencies in other large breast tumor collections such as
METABRIC. However, amplification does not correlate strongly with increased mRNA
(Figure 3a) or protein expression. ER
α
is not frequently mutated in primary cancer, while
recurrent mutations occur in a fraction of tumors progressing to resistance after hormonal
treatment [
80
]. Several of these mutations (e.g., hot spot mutations Y537S/N/C and D538G,
Figure 1a), lead to constitutive ER
α
activity, which generates resistance to aromatase
inhibitors. ER
α
mutations can also differentially impact the potency and/or efficacy of
clinically-relevant antiestrogens [80–84].
Cells 2021, 10, 2966 5 of 26
ing CCND1, MYC, E2Fs and FOXM1 [68–71]. In ER+ MCF-7 breast cancer cells, ERα pro-
tein levels are highest in the S and G2/M phases of the cell cycle [72]. The proliferative
action of estrogens in tumors as well as cell lines is confirmed by the observation that
hormonal therapies suppress proliferation of breast tumor cells [73].
2.2. Expression in Tumoral Tissue
In the clinic, breast tumors have been deemed ERα-positive (ER+ tumors) if more
than 1% of epithelial cells demonstrate nuclear staining in immunohistochemistry with
anti-hERα antibodies, qualifying patients for hormonal therapies, although rates of re-
sponse are much lower for tumors with low expression of ERα [74]. While there is a range
of ERα expression patterns between tumors, expression levels of the ERα protein are often
higher and more homogeneous than in normal tissue, especially in the LumA subgroup.
Nevertheless, some tumors exhibit heterogeneous expression, with variable proportions
of cells expressing ERα while staining is faint or absent in others.
Elevated ESR1 expression has been observed in atypical ductal hyperplasia and in
situ carcinoma, suggesting that deregulation of the ESR1 expression level may be impli-
cated in early pathogenic changes during breast tumorigenesis [62,75]. Whether ESR1
overexpression is sufficient to drive tumorigenesis in the human mammary gland is un-
clear. Interestingly, transgenic expression of wild type (wt) murine ERα in mice using an
MMTV-tTA controlled expression system led to ductal hyperplasia and ductal carcinoma
in situ by four months of age and to a low incidence of invasive carcinoma by 12 months
[76,77]. However, it remains unclear whether deregulated ESR1 expression is sufficient to
drive tumorigenesis in the human mammary gland.
The ESR1 gene is affected by copy number variations, with amplification events ini-
tially reported at a high frequency (20%), correlating with higher ERα levels [78]. On the
other hand, Brown et al. found amplification in only 1% of tumors [79]. Amplification is
predicted by GISTIC in less than 4% of tumors in the TCGA Firehose Legacy dataset (Fig-
ure 2) and at similar low frequencies in other large breast tumor collections such as META-
BRIC. However, amplification does not correlate strongly with increased mRNA (Figure
3a) or protein expression. ERα is not frequently mutated in primary cancer, while recur-
rent mutations occur in a fraction of tumors progressing to resistance after hormonal treat-
ment [80]. Several of these mutations (e.g., hot spot mutations Y537S/N/C and D538G,
Figure 1a), lead to constitutive ERα activity, which generates resistance to aromatase in-
hibitors. ERα mutations can also differentially impact the potency and/or efficacy of clin-
ically-relevant antiestrogens [80–84].
Figure 2. Genetic alterations in ESR1, GATA3 and FOXA1 in the TCGA Firehose Legacy breast invasive carcinoma dataset.
Amplification, deep deletion or mutations in genes for luminal transcription factors ERα, FOXA1 and GATA3 are shown.
The figure was downloaded from cbioportal.org by selecting the TCGA Firehose Legacy dataset, querying the ESR1,
FOXA1 and GATA3 genes and using the OncoPrint tool.
In primary tumors, ESR1 mRNA expression correlates very well with protein expres-
sion, suggesting that regulation of ESR1 expression takes place mainly at the RNA level
(transcription and/or mRNA stability) or that a balance between protein and RNA levels
Pag e 1 of 1
https://w w
w .cb iop o rtal.org/results/oncop rint
4%
4%
4%
14%
Genetic Alteration
Missense Mutation (putative driver) Missense Mutation (unknown significance)
Splice Mutation (putative driver)
Truncating Mutation (putative driver) Truncating Mutation (unknown significance) Amplification
Deep Deletion No alterations
ESR1
FOXA1
GATA3
Figure 2.
Genetic alterations in ESR1,GATA3 and FOXA1 in the TCGA Firehose Legacy breast invasive carcinoma dataset.
Amplification, deep deletion or mutations in genes for luminal transcription factors ER
α
, FOXA1 and GATA3 are shown.
The figure was downloaded from cbioportal.org by selecting the TCGA Firehose Legacy dataset, querying the ESR1,FOXA1
and GATA3 genes and using the OncoPrint tool.
In primary tumors, ESR1 mRNA expression correlates very well with protein expres-
sion, suggesting that regulation of ESR1 expression takes place mainly at the RNA level
(transcription and/or mRNA stability) or that a balance between protein and RNA levels
is maintained via tight feedback regulation (Figure 3b). The ESR1 mRNA is differentially
expressed in breast cancer subtypes (Figure 3c). In the transcriptome-based PAM50 classifi-
cation, ESR1 expression is highest in the luminal A and luminal B tumors, the latter being
associated with higher grade and worse prognosis [
85
]. On the other hand, the HER2+
tumors express the ESR1 mRNA at variable and overall weaker levels while the basal-like

Cells 2021,10, 2966 6 of 25
subtype has very weak to undetectable levels of ESR1 mRNA and of the ER
α
protein
(Figure 3c). The existence of discrete “intrinsic” subtypes with differential expression of the
ESR1 gene likely reflects the phenotype of the cell of origin giving rise to these tumors (stem
cell, bi-valent or univalent progenitor) and the inhibitory impact of genetic/epigenetic
aberrations in breast cancer cells on normal epithelial cell differentiation.
ESR1 expression is rapidly lost in primary cultures of normal epithelial cells, either
through epigenetic silencing or lack of proliferation/death of ER-positive cells. In addi-
tion, only a handful of ER
α
-expressing cell lines have been derived from ER+ tumors.
ER+ tumors
are also more difficult to engraft in Pdx models. Thus, our knowledge of ER
signaling in tumors obtained from existing models may be biased toward more aggressive
tumors. Nevertheless, ER+ cell lines such as MCF-7, T-47D and ZR-75-1 cells have proven
very useful to dissect pathways regulating ER
α
expression as well as its role in the control
of BC cell proliferation [86].
Methylation studies of the ESR1 gene have demonstrated that a CpG region located
from the acceptor splice site in the first coding exon to the end of this exon (exon 1,
Figure 4
)
is unmethylated in ER+/PR+ tumors and in ER+ cell lines MCF-7, T-47D and ZR-75-1,
while it is partially methylated in ER
−
MDA-MB-468 cells and completely methylated
in mesenchymal MDA-MB-231 cells [
87
]. Surprisingly, ER+/PR- tumors also presented
some methylated CpG sites [
87
]. These results may be explained by the inclusion in
the ER+ group of tumors with low ER
α
levels, resulting in undetectable expression of
PR, whose gene is under estrogenic regulation. In addition, ESR1 expression is strongly
negatively correlated with DNA methylation downstream of the major transcription start
site in the TCGA breast tumor dataset (Figure 3d). Treatment of triple negative cell
lines with DNA methyltransferase inhibitors such as 5
0
aza-deoxycytidine and/or histone
deacetylase (HDAC) inhibitors was reported to induce expression of the ESR1 gene, leading
to sensitivity to tamoxifen [88–91]. However, other studies have failed to reproduce these
observations [
92
]. On the other hand, treatment of ER
α
-expressing breast and uterine cancer
cells with HDAC inhibitors leads to suppression of ESR1 expression [
93
–
96
]. Recently,
treatment of triple negative breast tumor cells MDA-MB-231 LM2 with EZH2 inhibitors
has been shown to induce GATA3 expression and, more modestly, ESR1 expression, and to
sensitize these cells as well as another TNBC cell line (MDA-MB-468) to treatment with the
antiestrogen fulvestrant [97].
These observations suggest that epigenetic control through DNA methylation and
histone acetylation/methylation plays a role in ESR1 expression in breast cancer cells.
In addition to this, mutations activating or inactivating one of the multiple regulatory
elements in the ESR1 gene [
98
], from changes in expression/activity of upstream regulators
(see below), or a combination of both, may also contribute to the regulation of ESR1
expression. Altogether, the existence of different mechanisms of ESR1 gene expression
activation likely contributes to the heterogeneity of ESR1 expression levels and patterns
between and within tumors.

Cells 2021,10, 2966 7 of 25
Figure 3.
Expression of the ESR1 gene is epigenetically controlled (
a
). Impact of CNVs on ESR1 expression in the TCGA
Firehose Legacy breast invasive carcinoma dataset. (
b
). Correlation between ESR1 mRNA and protein expression in the
same dataset. (
c
). Differential expression of ESR1 within PAM50 breast tumor types (
d
). Correlation between ESR1 gene
expression and DNA methylation in the first intron in the TCGA Firehose Legacy breast invasive carcinoma dataset. Panels
(
a
,
b
,
d
) were downloaded from cbioportal.org by querying the ESR1 gene and using the Plots tool. Panel c was produced
using the PAM50 classifier on the TCGA breast invasive carcinoma dataset in [99].

Cells 2021,10, 2966 8 of 25
3. ESR1 Gene Organization and Regulatory Sequences
3.1. ESR1 Gene Structure and Alternative Transcripts
The analysis of different cDNA clones for the ESR1 gene has led to the characteri-
zation of seven promoters controlling nine upstream exons (A, B, C, D, T1/T2, F, E2/E1,
Figure 4) [49,100,101]. The splicing of exons E1, T1, T2, D, C, and B to a common acceptor
splice site located at position +163 in the first exon transcribed from promoter A yields 7
ESR1 transcript isoforms that can all be translated into the same 66 kDa protein [
49
,
100
,
101
].
The seven alternative promoters are regulated in a tissue-specific manner and generate
transcripts with unique 50untranslated regions (50UTRs).
Nevertheless, several isoforms of the ER
α
protein can result from alternative splicing.
A truncated form of ER
α
with an alternative, shorter N-terminal end, ER
α
-46 (
Figure 1b
),
has been identified in breast cancer cells. It results from transcription initiated at the
E2 or
F exons
, spliced to exon E1 and then directly to exon 2, deleting exon 1 [
102
,
103
].
Expression of this isoform is regulated by specific transcription factors [
104
,
105
]. The
homeobox transcription factor BARX2 has been shown to bind to the ESR1 gene at the
level of the E1 and F promoters and to upregulate transcription of this isoform [
104
]. The
high mobility group A protein 1a (HMGA1a) is involved in the alternative splicing mecha-
nism of this transcript by the recruitment of the RNA-protein complex U1 snRNP at the
acceptor splice site [
105
]. ER
α
-46 translation is intiated at an ATG codon in a favorable
Kozak consensus sequence in exon 2 and lacks most of the AF1 transcriptional activation
domain, acting in a dominant negative manner to suppress genomic estrogen signaling and
estrogen-dependent proliferation [
106
]. Furthermore, a shorter 36 kDa isoform has been
characterized [
107
]. This protein, encoded by exons 2 to 6 and an alternative exon 9, located
64 kbp downstream of the ESR1 gene, is truncated both in the AF1 (
lack of exon 1
) and
AF2 (lack of
exons 7–8
) transcriptional activation domains but preserves the DNA binding
domain, as well as a truncated ligand binding domain with a different 27 aa C-terminus.
The function of these truncated isoforms is not yet fully elucidated, although different
lines of research suggest both short isoforms are localized preferentially at the plasma
membrane [
108
]. This may result from the absence of exon 1, placing three potential myris-
toylation sites at residues 25–30 (GVWSCE), 76–81 (GMMKGG), and 171–176 (ELLTNL)
in both shorter isoforms, and therefore much closer to the N-terminus of the protein than
in the full length ER [
107
]. Moreover, the expression level of ER
α
-36 has been shown to
correlate with a worse prognosis, the proposed mechanisms being, in spite of the LBD
truncation, estrogen-dependent stimulation of mitogen signaling activity, activation by
antiestrogens such as tamoxifen, and increased stemness and metastasis of breast cancer
through upregulation of the aldehyde dehydrogenase 1A1 (ALDH1A1) gene [108,109].
Rearranged forms of the ESR1 gene have also been described. A duplication of
exons 6 and 7
, encoding part of the E domain, has been reported, leading to an ER
α
isoform of 80 kDa [
110
]. This longer isoform was found in a subclone of the MCF-7
cell line, MCF-7:2A, which contains four to five copies of the ESR1 gene and is estrogen
independent. Moreover, 88 ESR1 gene fusions with a break point in or near intron 6,
resulting in truncation of the LBD, have been characterized in metastatic ER-positive
disease with a frequency estimated at more than 1% [
111
]. Contrary to hER
α
-36, which
lacks exons 7 and 8 but was found to be activated by estrogens [
108
], these fusions are
reportedly devoid of estrogen-dependent activity but have different levels of constitutive
activity dependent on their fusion partners.
3.2. ESR1 Alternative Promoters Have Tissue-Specific Activity
Alternative transcripts of the ESR1 gene have been detected in both normal and can-
cerous tissues. In human cell models of ER+ breast tumors, ESR1 expression is driven pre-
dominantly by the proximal A promoter and the C promoter located 1.9 kb upstream [
112
].
Experiments from Grandien et al. demonstrated that both the A and C mRNA isoforms
were also expressed in endometrial tissue and uterine adenocarcinoma cell lines. Interest-
ingly, the mRNA levels of the A and C transcripts are widely different in breast cancer cells,

Cells 2021,10, 2966 9 of 25
with a ratio of approximately 20:1 [
112
]. The A transcript is more highly expressed in breast
tumors than in the normal mammary gland, but the opposite is observed for the C mRNA
isoform [
112
]. All transcripts except the T-transcript have been cloned from the MCF-7 cell
line and have been shown to be expressed in the mammary gland, in endometrium and/or
in liver [112].
Promoter A:
Despite a degenerate TATA box (TACTTAAAG), the A transcript is
the most abundant, whether in healthy mammary tissue or in luminal tumors [
113
,
114
].
DNA methylation analyses performed on a large panel of breast cancer cell lines revealed
that none of the 22 CpG sites identified in promoter A were methylated [
115
]. However,
data from ENCODE indicates that these sites are highly methylated in the HeLa cell line,
consistent with the lack of expression of ESR1 in these cells.
ENCODE ChIP-Seq data reveals binding of several transcription factors (TFs) within
500 bp of the transcriptional start site (TSS) of promoter A. FOXA1, GATA3 and ER
α
interact with the promoter in the T-47D BC cell line and CTCF, MYC and POLR2A interact
in the MCF-7 cell line. A predicted binding site for FOXA1 coincides with a FOXA1 ChIP
peak 400 bp upstream of the promoter A TSS. The existence of a half ERE in promoter A
(arrow, Figure 4) could possibly contribute to the recruitment of ER
α
[
114
,
116
]. ChIP-Seq
assays targeting the breast cancer type 1 susceptibility protein (BRCA1) have shown that
this protein binds the A promoter indirectly via Oct-1, encoded by the POU2F1 gene [
117
].
Oct-1 binds a region of promoter A between -385 and -169 bp and recruits BRCA1 as
a coactivator, resulting in positive transcriptional regulation of ESR1 [
117
]. In addition,
functional genomic studies have shown the presence of the p53 protein in the region
between
−
128 to
−
40 bp of promoter A, accompanied by other factors such as Sp-1, c-Jun
and the coactivator CBP, suggesting regulation of ESR1 by p53 via this promoter [
118
]. This
is consistent with the higher rate of p53 mutations in ER-negative vs. ER-positive and in
luminal B vs. luminal A tumors, although the extent to which p53 controls ESR1 expression
remains unclear. The recruitment of Sp-1 within the
−
245 to
−
192 bp region was found
to be essential for ESR1 expression and to mediate positive regulation of promoter A by
exogenous Sp-1 expression in Drosophila Schneider SL2 cells [119].
The presence of the repressive histone methylation mark H3K9me3 has been reported
at promoter A in basal breast cancer cells. ChIP-qPCR on FOXC1 wt and knock-out
BT549 basal-like breast cancer cells showed a correlation between trimethylation of H3K9,
FOXC1 expression and loss of RNA PolII recruitment at the ESR1 regulatory sequence.
This negative regulation of ESR1 gene expression reinforces the hypothesis that ESR1
heterogeneity between different breast cancer types is due to the establishment of distinct
epigenetic marks [120].
Promoter B:
As there are only about 360 bp between the B and A transcript start sites,
several of the TF binding sites upstream of promoter A overlap with promoter/exon B. The
B transcript has been detected in breast tissue at very low mRNA levels compared to the A
transcript. The B promoter and exon B include 18 CpG sites that are unmethylated both in
normal MCF-7 cells or in antiestrogen and anti-aromatase inhibitors resistant MCF-7 cells,
which express lower levels of ERα[115].
Promoter C:
Exon C presents two main transcription start sites located at
−
1974 and
−
2007 bp, generating C-transcripts with different 5
0
-ends [
113
]. C transcripts are ten times
less abundant than the A-transcript in MCF-7 cells and absent from ZR-75 cells. Promoter
C is highly methylated in ZR-75-1 cells, correlating with the absence of the C-transcript in
that cell line, while it is unmethylated to moderately methylated in ER-negative SK-BR-3
and MDA-MB-231 cells [115].

Cells 2021,10, 2966 10 of 25
Figure 4.
Schematic structural organization of human ESR1 promoters and alternative transcripts. The structure of the
human ESR1 gene is presented along the main axis, boxes corresponding to exons. Alternative upstream exons are colored
and labeled with letters according to the nomenclature proposed by Kos et al. [
49
]. White boxes correspond to exons
downstream of the common acceptor splice site and are numbered according to the same nomenclature. Alternative
promoters are represented by a colored thick line matching the color of the corresponding 5
0
exon. The arrows define
transcriptional initiation sites. The numbers under the main axis are exon start/end distances from the transcription start site
originally defined as +1 (transcript A), calculated based on mapped transcripts in the hg19 genome version. The numbers
between the exons represent the size of the introns in kilobase pairs. The common acceptor splice site is represented by a
vertical bar before exon 1 and the ATG start codon is indicated by a black arrow. The upper part of the diagram details the
promoters of exons A, C and F. Reported half-estrogen response elements (1/2 EREs) in these promoters are indicated by
arrows. Transcription factors bound in these regions based on published ChIP-Seq data are shown in bubbles and factors
directly binding DNA (motif predicted by HOMER) are in bold. The different transcripts produced after transcription and
splicing from each of the seven regulated upstream exons to exon 1 are also presented in the lower part of the diagram.
ENCODE ChIP-Seq data and literature reports suggest binding of multiple TFs to this
promoter in MCF-7 cells. The activity of the promoter C in luminal cell lines was observed
to depend on the binding of estrogen receptor associated factor 1 (ERF-1), corresponding
to the transcription factor AP-2 gamma (TFAP2C) [
121
,
122
]. This factor binds promoter
C at
−
1877 bp in a complex with Jun/Fos and plays an important role in the expression
of the ESR1 gene [
123
–
125
]. Sodium bisulfite genomic sequencing revealed that promoter
C comprises nine CpG sites [
115
,
121
]. The ninth methylation site is located in a TFAP2C
binding motif [
115
,
121
]. TFAP2C is expressed in both the luminal and myoepithelial
cells in the adult mammary gland and is expressed in all BC subtypes [
126
]. MMTV-Cre
induced knockout in mouse models leads to delayed migration through the mammary
fat pad without deleterious impact on mammary gland function, to an increase in the
CD24midCD49fhi population, an increase in basal CK5 staining and a decrease in the
normalized luminal:basal ratio. Down-regulation of TFAP2C in ER+ BC cells leads to
epithelial to mesenchymal transition (EMT) characterized by increased VIM and CD44
expression, with decreased CDH1 and CD24 expression and decreased luminal gene (ESR1,
FOXA1,GATA3) expression [
127
]. TFAP2C also regulates FOXM1 expression and its down-
regulation suppresses estrogen-dependent proliferation [
128
]. The related TFAP2A on the
other hand specifically induces CDKN1A and is inactive on luminal genes [129].
Specific methylation of the fourth CpG site in promoter C was responsible for a de-
creased expression of ESR1 and resistance to endocrine therapy; the absence of methylation

Cells 2021,10, 2966 11 of 25
at this specific CpG site allows the recruitment of the methylation-sensitive transcription
factor Ets-2, which leads to ESR1 expression in normal MCF-7 cells [
115
,
121
]. In the MCF-7
cell line, GATA3 and the negative regulator FOXC1 were reported to compete for binding
to DNA at overlapping recognition motifs [
120
]. Induced expression of FOXC1 in MCF-7
cells leads to a loss of GATA3 recruitment to promoter C, indicating that FOXC1 can abolish
the positive regulation of that promoter [
120
]. More distally, CTCF also binds promoter C
in the MCF-7 cell line. Finally, p53 is recruited at the C promoter in ChIP-qPCR assays in
the DNA region between
−
2094 to
−
1941 bp from the main TSS, which contains seven of
the nine methylation sites described previously [118,124,130].
Promoter D:
Promoter D was shown to be fully methylated in the MCF-7 cell line.
DNA methylation studies identified seven CpG sites methylated in that promoter, but the
expression level of the D-transcript did not correlate with the methylation status [
115
].
Moreover, an AP-1 binding site has been described as an enhancer active in MCF-7 cells
and inactive in ER-negative MDA-231 cells [125].
Promoter F:
The F-transcript, produced from the splicing of exons F and E1, is the
predominant variant expressed in primary osteoblasts but is also highly expressed in
breast cancer tissue [
114
,
131
]. As mentioned above, promoter F was found to be regulated
by BARX2 in breast normal and tumoral tissue [
104
]. FOXA1, GATA3 and p300 also
interact with this promoter in the MCF-7 and T-47D cell lines, correlating with the levels of
F-transcripts
in ER-positive breast cancer cells [
132
]. The existence of a half ERE in promoter
F (arrow, Figure 4) was suggested to contribute to the recruitment of ER
α
[
114
,
116
]. E2F1
is also associated with promoter F in ENCODE MCF-7 ChIP-Seq data. DNA methylation
analysis in this promoter has shown the presence of 10 CpGs, unmethylated in MCF-7 but
all methylated in the MDA-MB-231 cell line [132].
Promoter E:
The E-transcript is produced by the transcription of exons E2 (upstream
of promoter F) and E1 (downstream of promoter F) and is expressed mainly in the liver
and at very low levels in breast cancer cells [114].
Promoter T:
This promoter is the only one not to be active in mammary glands or
in breast tumors. The T-transcript is predominantly expressed in testis and epididymal
tissues. The T1 and T2 exons are separated by a 101 bp intron and are located about 16 kb
upstream of the first exon [
49
,
133
]. Transcription from the T promoter can form either a
longer variant from the splicing of exon T1 to exon T2 and finally to exon 1, or a shorter
variant from the splicing of the T1 exon directly to exon 1.
3.3. Upstream Enhancers
Distal enhancers can regulate transcription at up to several hundred kilobases away
from target genes via chromatin looping [
134
]. ChIP experiments have revealed binding
of luminal TFs FOXA1, GATA3, and ER
α
itself to far upstream regulatory sequences in
the ESR1 gene [
19
,
135
–
137
]. All regions interacting with these luminal TFs are associated
with enhancer chromatin marks H3K4Me1 and H3K27Ac in ENCODE datasets generated
in MCF-7 cells, supporting the existence of at least five “luminal” enhancers located at
distances greater than 120 kb upstream of exon A (Figure 5; enhancers 1–3 are upstream of
exon E2, enhancers 4 and 5 are found between exons E2 and F). FOXA1 and GATA3 bind
to enhancers 1–5 and ER
α
to enhancers 1–4. Thus, ER
α
, GATA-3 and FOXA1 appear to
share cis regulatory sites in the ESR1 gene as well as upstream of many estrogen target
genes [
19
,
135
–
137
]. Enhancers 1 and 4 contain motifs related to the consensus ERE se-
quence 5
0
-(A/G)GGTCANNNTGACC(T/C)-3
0
[
138
–
140
]. Predicted FOXA1 motifs related
to the consensus 5
0
-A(A/T)TRTT(G/T)RYTY-3
0
[
141
] are found in enhancers 2, 3 and 4.
Sequences matching the consensus GATA binding motif 5
0
-(A/T)GATA(A/G)-3
0
[
142
–
144
]
are predicted by HOMER in enhancers 4 and 5. A reported GATA3 site [98] not predicted
by HOMER is located in enhancer 2. Thus, one or more predicted binding motifs for ER
α
,
FOXA1 and GATA3 are found in each of the five enhancers. While ER
α
, FOXA1 and
GATA3 are detected at regions that do not contain mapped binding motifs, lower affinity
sequences may be accessed through DNA binding cooperativity and/or pioneer activity.

Cells 2021,10, 2966 12 of 25
Alternatively, these TFs may be tethered to other TFs directly recruited to DNA. Other
transcription factors are also present at these sites as annotated in ENCODE ChIP-Seq
datasets in MCF-7 cells, suggesting that they correspond to active enhancers [
30
]. The
coactivator p300 (EP300), which acts as a general transcriptional coregulator with histone
acetyltransferase activity [
145
], and is present at 51% of the ER
α
binding sites after E2
treatment [
146
], is strongly recruited to enhancers 1–4, as well as RNApolII. While these
enhancers are not clustered in super-enhancers, we note that enhancers 3 and 4 are asso-
ciated with CTCF and components of the cohesin complex, corresponding to the profile
of “hub” enhancers [
32
], which play major roles in local chromatin organization. Finally,
chromatin loops were observed to form between enhancers 1 and 2/3, 1 and 4, 2–3 and 4,
and 1 and promoter F by ChIA-PET with ER
α
in MCF-7 cells [
147
]. Looping with other
regions not strongly associated with luminal factors in MCF-7 cells was also observed,
suggesting that other regulatory sequences play roles in ESR1 transcription, in keeping
also with the existence of several other regions associated with H3K4Me1/H3K27Ac and
DNAse hypersensitivity.
Figure 5.
Enhancer regions of the human ESR1 gene bound by luminal TFs. Colored rectangles highlight the luminal
enhancers defined by ChIP-Seq assays with luminal factors ER
α
, FOXA1 and GATA3 and coactivator p300 [
19
,
135
–
137
].
Alternative exons spliced with coding exon 1 are shown as in Figure 4. The promoters are represented by a colored thick
line upstream of the corresponding alternative first exons. Broken arrows define the different transcription initiation sites.
Distances between the beginning or end of each defined feature and the transcription start site in promoter A are indicated.
The distance in kilobase pairs between two highlighted elements is shown under the DNA axis. The common acceptor
splice site is represented by a black line and the ATG start codon is indicated by a black triangle. Transcription factors
bound to each enhancer based on ChIP-Seq data are shown in bubbles above or below each element, and factors for which a
recognition motif is predicted by HOMER or mapped in a referenced study are in bold. DNA sequences from the enhancers
containing one or more binding motifs for GATA3 (Green), FOXA1 (Pink) and/or ER
α
(Blue) are shown in magnification
boxes (motifs for each TF are in bold).
Breast cancer risk-associated single nucleotide variants (SNVs) and somatic mutations
have been mapped to luminal enhancers and other DNAse hypersensitive sites within
1 Mb
of the ESR1 start site [
98
]. The rs9383590 SNV affects the reported GATA3 binding
site in enhancer 2 and is predicted to increase activity of the enhancer. This suggests that
this GATA3 element acts as a negative rather than positive regulator of ESR1 expression.
Of interest, this motif is located within a TCF7L2 ChIP binding region (ENCODE). The
transcription factor TCF7L2 has binding patterns in MCF-7 cells that coincide with those
of GATA3, resulting in repression of their common target genes [
148
] (see also below). In
addition, mutation chr6:151979547:A>G in enhancer 4 affects a nucleotide within a pre-
dicted FOXA1 motif (5
0
ATTGTTTGCTG 3
0
) [
98
]. While it is unclear whether this mutation
affects FOXA1 binding, residues flanking the core FOX motif (5
0
-RTAAAYA-3
0
) can alter
FOX factor binding specificity (see below). Point mutations that led to increased activity in
reporter gene assays were also found in tumors between enhancers 2 and 3 (chr6:151954506
C>A), after enhancer 3 (chr6: 151955192A>G; chr6:151955219:G>T), downstream of the E2

Cells 2021,10, 2966 13 of 25
promoter (chr6:152024472C>G) or upstream of promoter C (chr6: 152125116G>C); these
mutations may suppress binding by transcriptional repressors or create new binding sites
for transcriptional activators, suggesting that a variety of transcription factors bound to
different regulatory regions regulated ESR1 expression. Based on their data, Bailey et al.
predicted that mutations leading to increased enhancer activity may explain the sustained
expression of ESR1 in some (about 7%) luminal breast tumors [98].
4. Luminal ESR1 Transcriptional Regulators
4.1. GATA3
The GATA3 gene, located at 10p14 in the human genome, codes for a 48 kDa protein
composed of two N-terminal transactivation domains, two zinc-finger DNA-binding mo-
tifs followed by two highly conserved and distinct basic regions (basic
region 1 and 2
) and
a
C-terminal
region required for GATA3 transactivation, which contains a conserved YXKX-
HXXXRP motif [
144
,
149
,
150
]. The GATA proteins form a family of six zinc finger DNA binding
proteins that recognize motifs related to the consensus sequence
50-A/TGATAA/G-30[142–144]
.
GATA3 was
shown to bind DNA with its C-terminal zinc-finger region and form ho-
modimers, or heterodimers with other GATA members via its N-terminal zinc-finger
region [
144
,
151
]. On certain specific DNA sequences, two close GATA sites can be bound
simultaneously by the two zinc fingers of one GATA3 molecule with higher affinity [
144
].
Like ER
α
, GATA3 interacts with the acetyltransferase CREBBP/CBP, which acetylates it
at K119, contributing to its capacity to downregulate mesenchymal transcription factors
ZEB1/2 and Slug [152].
In breast tumors and cell lines, expression of GATA3 is strongly associated with those
of ESR1 and FOXA1 [
153
]. On the other hand, GATA1, 2, 4 and 5are only mildly or
not correlated, while GATA6, the only GATA gene whose expression is biased toward
basal-like tumors, is mildly anti-correlated with ESR1 in the TCGA dataset [
99
]. GATA3
was identified as a regulator of mammary branching morphogenesis by genome-wide
transcript analysis [
154
] and shown to maintain the differentiation of luminal cells in
mouse models [
155
–
157
]. High levels of expression of GATA3 in ER-positive cells are
consistent with the reported positive cross-regulation feedback loop between the GATA3
and ESR1 genes [
158
], which also likely involves the FOXA1 luminal transcription factor. As
mentioned above, GATA3 interacts with the ESR1 promoters A and C but also with the five
enhancers associated with FOXA1 in ChIP-Seq datasets [
120
,
136
] (Figure 5). Conversely,
an enhancer recruiting ER
α
, FOXA1 and GATA3 itself, together with the coactivator
p300, was described approximately 10 kb downstream of the GATA3 gene (about 30 kb
downstream of the TSS) [
158
]. Another region located 53.4 kb downstream of the gene
(
73.4 kb
downstream of the TSS) is even more strongly associated with all factors in relevant
ChIP-Seq/Exo datasets [
136
,
137
] and contains a predicted ERE and several FOX and GATA
motifs, supporting the existence of regulatory loops between these factors.
Consistent with the role of GATA3 in the regulation ESR1 expression, GATA3 silencing
by two different siRNAs was observed to reduce ESR1 expression in T-47D and MCF-7 cells,
to blunt induction of transcription of estrogen target genes, and to inhibit estrogen-induced
proliferation [
158
]. On the other hand, expression of GATA3 in MDA-MB-231 cells results
in reprogramming of these cells from a mesenchymal to a luminal subtype, associated with
a reduction of tumorigenesis and metastasis in implanted xenograft assays; GATA3 also
induces a growth inhibitory response to TGFβ[159,160].
Moreover, GATA3 may act as a pioneer factor by binding target sites within nucleo-
somes and making neighboring cis-regulatory elements accessible for the recruitment of
additional transcription factors [
161
–
163
]. During the mesenchymal-to-epithelial transition
induced by GATA3, cellular reprogramming involves in part binding to closed chromatin
regions, nucleosome eviction and chromatin remodeling in a transcriptional activation
domain-dependent manner [
162
]. In ER-positive breast cancer cell lines, enrichment in
GATA motifs was observed in ER
α
and FOXA1 ChIP-Seq/Exo peaks [
136
,
137
] and GATA3
silencing decreased or even abolished the recruitment of ER
α
at distal regulatory regions

Cells 2021,10, 2966 14 of 25
normally co-occupied with GATA3 [
161
]. Recruitment of common cofactors may contribute
to complex formation between GATA3 and other luminal TFs. GATA3 may also recruit
other transcription factors by tethering. GATA3 expression was shown to be required for
the high mobility group box-containing factor TCF7L2 to bind to about 50% of TCF7L2 sites
in the ER-positive breast cancer cell line MCF-7, and these sites were enriched in GATA3
but not TCF7L2 motifs [
148
]. GATA3 and TCF7L2 bound to shared sites simultaneously
and tagged TCF7L2 could co-immunoprecipitate with endogenous GATA3 in transfected
MCF-7 cells. TCF7L2 functioned mainly as a transcriptional repressor at shared sites [
148
].
TCF7L2 is recruited at the level of enhancers 1, 2 and 4 of the ESR1 gene; however, whether
TCF7L2 represses ESR1 expression needs to be confirmed.
GATA3 is the third most mutated gene in luminal breast cancers with a prevalence
of approximately 14% and 15% in luminal A and B tumors, respectively (cbioportal.org,
TCGA Firehose Legacy dataset). GATA3 is also amplified in breast tumors (Figure 2),
mostly in ER-negative tumors where amplification correlates with decreased rather than
increased GATA3 expression, suggesting the existence of different driver genes in this 10p14
amplicon [
164
]. On the other hand, GATA3 is frequently affected by mutations in ER+ BC,
with a bias for invasive ductal carcinoma, including splicing mutations (mostly X308_splice)
and truncating frameshift mutations (in or after the second zinc finger) [
165
,
166
]. GATA3
mutations can lead to active forms of the GATA3 protein that contribute to tumor growth
in BC cell xenografts and promote precocious lobuloalveolar development in transgenic
mice [
167
]. In addition, ER-positive breast tumors with GATA3 mutations were reported
to have a worse prognosis compared to wt tumors [
168
]. In the MCF7 cell line, GATA3
is affected by a heterozygous D336fs mutation in the second zinc finger that leads to a
truncated protein with reduced affinity for DNA. Both wt and mt GATA3 have increased
stability in MCF-7 vs. wt GATA3 in T-47D cells, leading to increased genomic occupancy
by GATA3; genomic distribution was similar as in wt T47D but with gene-specific loss
of binding, such as at the progesterone receptor gene [
169
]. Mutation R330fs was on
the other hand reported to alter genomic distribution of ER
α
and FOXA1, leading to
changes in gene regulation affecting genes involved in mammary gland development
and epithelial cell biology [
170
]. In contrast, the X308_splice mutation, which generates a
GATA3 protein lacking the second zinc finger but containing a novel 44 aa C-terminal end,
downregulates ER genomic signaling and is associated with improved overall survival in
ER+ patients [171].
4.2. FOXA1
The forkhead box (FOX) family of transcription factors comprises 50 human mem-
bers that share a characteristic ‘winged-helix’ DBD composed of a helix-turn-helix motif
with two C-terminal loops or “wings” [
172
] and regulate gene expression spatially and
temporally during development [
173
]. These transcription factors can be divided into
19 subgroups (FOXA to FOXS) based on the sequence of their DBDs. Although all FOX
proteins recognize core motifs related to the consensus 5
0
-RTAAAYA-3
0
, flanking sequences
contribute to DNA binding specificity within the family [
172
,
174
]. Further, divergence in
FOX factor primary sequence outside of the DBD results in a wide range of biological func-
tions. In particular, FOXA1 plays an essential role in mammary ductal morphogenesis [
175
],
and its expression is strongly correlated with that of ER
α
in breast luminal tumors [
153
].
FOXA1 has been identified as an upstream regulator of ESR1 expression in mouse and
human breast cancer cells [
175
,
176
] and as a repressor of basal-like genes [
177
]. Notably,
FOXA1 is co-associated with GATA3 at several enhancers in the ESR1 gene (Figure 5) and
may thus cooperate with it for regulation of ESR1 expression. In addition, FOXA1 can act as
a pioneer transcription factor, directly binding and opening condensed chromatin [
178
,
179
].
FOXA1 can enable recruitment of ER
α
and the androgen receptor (AR) to their respective
response elements in breast and prostate cancer, respectively [
19
,
26
,
180
–
182
]. Beyond facili-
tation of hormone receptor signaling, FOXA1 overexpression has been shown to contribute

Cells 2021,10, 2966 15 of 25
to tamoxifen insensitivity of breast cancer cells in a mechanism that involves induction of
interleukin-8 [183].
The FOXA1 gene is infrequently amplified (Figure 2), and copy number gain does
not associate with markedly higher expression levels, while shallow deletion and DNA
methylation may account for low FOXA1 expression (cbioportal.org). Mutations in FOXA1
are less frequent than those in GATA3 and are mostly missense mutations associated
with high FOXA1 mRNA levels. Contrary to GATA3 mutations, they are frequent in
invasive lobular carcinoma [
166
]. These mutations affect mostly the forkhead domain, the
most common in primary tumors being I176M/V, D226G/N and S250F (cbioportal.org,
Breast invasive carcinoma, TCGA Firehose Legacy dataset). Mutations are enriched in
metastatic tumors and correlate with resistance to aromatase inhibitors. Mutations in
the
Wing 2 region
increase cooperativity with ER while mutation SY242CS, located in
the third beta strand, leads to altered DNA binding patterns [
184
]. In addition, a G>A
mutational hotspot was discovered in ER+ breast tumors at position -81 relative to the
FOXA1 TSS, resulting in increased binding of E2F1 and FOXA1 overexpression [
185
]. Thus,
enhanced expression or activity of FOXA1 could in turn result in increased ER
α
levels via
direct transcriptional regulation of ESR1 expression, in addition to the role of FOXA1 in
cooperating with ERαfor transcriptional regulation.
FOXA2 and FOXA3 are the closest homologs of FOXA1 in the FOX family. RNA
levels of both factors are much lower than those of FOXA1 in breast cancer, and neither are
positively correlated with ESR1 expression. FOXO family members are also expressed in
breast tumors, without a strong bias for luminal tumors. Activity of these factors is under
post-translational regulation. For instance, FOXO protein expression is downregulated by
the Akt pathway [
186
]. FOXO3 was reported to positively regulate ESR1 expression, the
expression of a dominant negative form repressing expression of ESR1 in MCF-7 cells [
187
],
and binding sites were identified in promoters A and B. However, FOXO3 was also reported
to suppress the transcriptional activation properties of ER
α
via protein–protein interactions
and to suppress growth of ER+ breast tumor cells [
188
]. In addition, FOXO3 mediates the
cytostatic and cytotoxic properties of breast cancer therapeutics, such as anthracyclins and
taxanes, and is down-regulated in MCF-7 cells resistant to epirubicin and paclitaxel [189].
A number of other FOX family members have been reported to regulate ER
α
function
and/or expression. For instance, FOXM1 drives breast cancer cell proliferation under
transcriptional control by ER
α
. It also activates ESR1 transcription via binding to several
sites upstream of promoters A, B and C, possibly in interaction with FOXO3, which bound
the same sites and interacted with FOXM1 in co-immunoprecipitation experiments [
190
].
One site identified in promoter A overlaps with a ChIP peak for FOXA1 [
137
]. Despite
these observations, FOXM1 is highly expressed in triple negative tumors [
191
], and in mice
inhibits luminal differentiation via inhibition of GATA3 expression [192].
FOXC1 and FOXC2 expression patterns are likewise elevated mostly in basal-like
tumors [
193
,
194
]. FOXC1 repressed ESR1 expression when overexpressed in three ER+
BC cell lines. ChIP qPCR and pulldown with biotinylated antibodies indicated binding
of GATA3 and FOXC1 to overlapping motifs in promoter C and enhancer 4. FOXC1-
mediated suppression of ESR1 was accompanied by exclusion of GATA3 binding, increased
H3K9me3, and reduced KDM4B and RNApolII interaction at these sites and upstream of
promoter A [120].
Together, these studies suggest a positive role for FOXA1 in ESR1 up-regulation in
ER+ breast cancer cells, with possible cross-talk with several FOX factors.
4.3. Estrogen Receptor Alpha
Published ChIP-Seq/ChIP-Exo datasets [
19
,
135
–
137
,
195
] show that ER
α
binds ESR1
enhancers 1 to 4; in addition, EREs were detected in the center of the ChIP peaks corre-
sponding to enhancers 1 and 4 (Figure 5). Previous studies reported half-ERE motifs in
promoters A [
116
] and F [
114
] (Figure 4), although these motifs do not correlate with strong
recruitment of ER
α
at those sites in the ChIP-Exo study. The presence of ER
α
at distal

Cells 2021,10, 2966 16 of 25
enhancers is in agreement with reports that ER
α
is primarily recruited at enhancers rather
than promoters of estrogen-modulated genes in breast cancer cell lines [19,135].
Consistent with its recruitment on ESR1 regulatory regions, ER
α
exerts feedback
regulation on its own expression in a cell context-dependent manner [
116
,
158
]. Indeed,
ESR1 mRNA levels are decreased by 17-
β
-estradiol in MCF-7 and induced in T-47D [
196
].
The basis for this differential regulation, as well as the reasons dictating the wide variations
in ESR1 expression levels in BC cell lines and tumors, remain unclear. Of note, exogenous
expression of ER
β
in MCF-7 cells identified genomic targets that for the vast majority
overlapped with ER
α
binding regions [
197
]. This likely results from the capacity of both
receptors to recognize the same binding sites and to heterodimerize; however, divergence in
their transactivation regions often result in different effects on transcription, heterodimers
having intermediate profiles [
198
,
199
]. Whether ER
β
can bind ESR1 regulatory regions
and cross-regulate its expression in normal tissue, in which it is expressed to higher levels
than in breast tumors, remains to be clarified.
Estradiol treatment also induces ER
α
turnover in tumor cells, leading to an attenuation
of ER signaling [
196
,
200
]. The existence of negative feedback mechanisms on ER
α
levels
in the presence of agonists in normal tissue [
201
] may contribute to the transient nature
of the proliferative response to estrogens during the hormonal cycle and may explain in
part the lack of correlation between ER
α
and proliferation markers such as KI67 in the
normal human mammary gland (see above). In tumors or in normal tissue of high-risk
women [
201
], bypass of these control mechanisms and ER
α
overexpression may drive
mammary cell proliferation.
5. Conclusions
The identification of long-range regulatory elements in ESR1 by ChIP-Seq analysis
together with the mapping of alternative promoters outlines a complex mode of ESR1
regulation. Transcription factors bind these regulatory elements directly via the recog-
nition of cognate DNA target motifs but may also interact indirectly with other regions
via long-range chromatin loops, through direct or co-factor-mediated interactions with
other transcription factors. Luminal factors FOXA1 and GATA3 contribute to the positive
regulation of the ESR1 gene, with ER
α
and possibly ER
β
exerting context-dependent
feedback regulation. CpG sites methylated in a cell-specific manner are found across
the entire ESR1 gene and likely modulate expression of alternative promoters. Epige-
netic modulation via molecules inhibiting histone/DNA methyltransferases or histone
deacetylases suggests that ESR1 expression is controlled in a cell-specific manner by
chromatin-modifying/remodeling enzymes potentially recruited to the ESR1 gene by spe-
cific transcription factors [
202
]. Luminal transcription factors described above likely play
an important role in recruiting coactivators such as histone acetyl transferases, and chro-
matin remodelers such as the SWI-SNF complex. Repressive complexes may be recruited
by negative regulators such as FOXC1 or other repressors, including the transcription
factor ELF5, shown to repress ESR1 and a set of ER
α
-associated genes with promotion of
basal-like breast cancer characteristics [
203
]. Mesenchymal transcription factors have also
been observed to repress ESR1 expression [
204
,
205
]. Other regulators not discussed in this
review include miRNAs; several are capable of targeting the ESR1 3
0
UTR and some are
negatively correlated with ESR1 in breast cancer [206].
In spite of the accumulated knowledge about regulators of ESR1 transcription, our
understanding of the mechanisms that control ER
α
expression in normal tissue and how
they are altered during breast tumorigenesis requires further investigation. A better
dissection of the hierarchical nature of enhancers controlling the ESR1 gene is required to
identify which factors are key to enhancing or silencing ESR1 expression. Several epigenetic
regulators have been proposed to induce ESR1 expression in triple negative tumors, but
whether this will result in therapeutic approaches based on subtype conversion remains
unclear. Finally, it is crucially important to better understand reasons for the variable
expression patterns of ER
α
in tumors, the nature of ER-negative cells in heterogeneous

Cells 2021,10, 2966 17 of 25
ER-positive tumors, the impact of hormonal therapies on these tumors and the mechanisms
of the apparent conversion of some ER-positive to ER-negative tumors during cancer
progression. The recent availability of single cell and spatially resolved genomic analyses
should greatly facilitate the study of intra-tumoral heterogeneity in ER
α
expression in the
near future.
Author Contributions:
Writing—original draft preparation, L.P. and H.I.; diagrams—creation and
design, L.P.; writing—review and editing, supervision, S.M. All authors have read and agreed to the
published version of the manuscript.
Funding:
This research was funded by the Canadian Institutes of Health Research, grant number
PJT153178.
Institutional Review Board Statement: Not applicable.
Informed Consent Statement: Not applicable.
Data Availability Statement:
No new data were created or analyzed in this study. Data sharing is
not applicable to this article.
Acknowledgments:
The authors apologize for not being able to cite all relevant articles due to space
limitations and are grateful to John White for manuscript reading and comments.
Conflicts of Interest:
The authors declare no conflict of interest. The funders had no role in the design
of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or
in the decision to publish the results.
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