New Results

G protein-coupled estrogen receptor regulates heart rate in zebrafish embryos

Shannon N Romano, Hailey E Edwards, J Paige Souder, Xiangqin Cui, Daniel A Gorelick
doi: https://doi.org/10.1101/088955

Abstract

Estrogens act by binding to estrogen receptors alpha and beta (ERα, ERβ), ligand-dependent transcription factors that play crucial roles in sex differentiation, tumor growth and cardiovascular physiology. Estrogens also activate the G protein-coupled estrogen receptor (GPER), however the function of GPER in vivo is less well understood. Here we find that GPER is required to maintain normal heart rate in zebrafish embryos. Acute exposure to estrogens increased heart rate in wildtype and in ERα and ERβ mutant embryos but not in GPER mutants. Nuclear estrogen receptor signaling remained normal in GPER mutant embryos, however GPER mutant embryos exhibited reduced basal heart rate while heart rate was normal in ERα and ERβ mutants. We detected GPER transcript in discrete regions of the brain but not in the heart. In the brain, we observed gper expression in cells lacking nuclear estrogen receptor activity, suggesting that GPER acts in the brain to regulate heart rate independently of nuclear estrogen receptor signaling. Our results demonstrate that estradiol plays a previously unappreciated role in the acute modulation of heart rate during zebrafish embryonic development and that GPER functions as an autonomous estrogen receptor in vivo to regulate basal heart rate.

INTRODUCTION

Zebrafish are an established model for human cardiovascular development and function (1) with conserved estrogen signaling (2-4). While studying the function of ERa (esr1) in zebrafish embryonic heart valves (5,6), we serendipitously observed that estrogen receptor modulators caused acute changes in heart rate. Estrogens bind two classes of receptors: nuclear hormone receptors (ERα, ERβ) that are ligand-dependent transcription factors (7), and the G protein-coupled estrogen receptor (GPER, also known as GPR30), an integral membrane protein (8,9). It has been difficult to tease apart to what degree ERα and/or ERβ are involved in regulating GPER function in vivo. The observations that ERα can directly activate G proteins in cultured cells (10-13) and that GPER coimmunoprecipitated with ERα in tumor cells (14) has been used to argue that either GPER is dispensable for estrogen-dependent signaling or that GPER mediates interactions between ERα and G proteins (15). Studies using GPER-deficient mice implicate GPER in ventricular hypertrophy (16), regulation of blood pressure and vascular tone (17,18) and atherosclerosis progression (19), but whether nuclear ER signaling is required for GPER function in these contexts is unknown. Additionally, these studies examined GPER function in adult mice, while the role of GPER during embryonic development is not well understood.

Estrogen signaling often differs between males and females. However, zebrafish embryos and larvae are bipotential hermaphrodites that have not begun to sexually differentiate before approximately 10 days post fertilization (dpf) (20), meaning that estrogen levels are uniform between age-matched embryos. Additionally, zebrafish embryos develop outside of the mother and not within a confined space, such as the uterus. Therefore, zebrafish embryos are not subject to local estrogen concentration gradients, as has been reported to occur in rodents depending upon their position in utero and their proximity to embryos of the same or opposite sex (21,22). These developmental traits make zebrafish a powerful model to study how sex hormone signaling influences the formation and function of non-gonadal tissues. Using complementary genetic and pharmacologic approaches, we sought to characterize how estradiol regulates heart rate and determine to what extent each estrogen receptor mediates estradiol-dependent changes in heart rate in zebrafish embryos.

RESULTS

We exposed 49 hour post fertilization (hpf) embryos to 17β-estradiol (estradiol) and assayed heart rate following one hour exposure. We found that estradiol exposure caused an approximately 20% increase in heart rate (Fig. 1, mean difference in heart rate between estradiol and vehicle exposed embryos was 26.51 ± 3.36 (standard error) beats per minute (bpm)). Exposure to progesterone, a structurally similar steroid sex hormone, had no effect on heart rate (Fig. 1, mean difference in heart rate 2.31 ± 6.54), suggesting that the effects on heart rate were specific to estrogens.

Figure 1.
Figure 1. Estradiol and GPER agonists increased heart rate in zebrafish embryos.

Wildtype embryos were incubated in water containing vehicle (0.1% DMSO), estradiol (3.67 μM, ER/GPER agonist), progesterone (1 μM), ICI (10 μM ICI182,780, ER antagonist/GPER agonist), G1 (1 μM, GPER agonist), G36 (1 μM, GPER antagonist) or two chemicals in combination at 49 hours post fertilization and heart rates were measured 1 hour following treatment. ***, p<0.0001 compared to vehicle, ANOVA with Dunnett’s test. Each black circle represents the mean heart rate from a single clutch of embryos (3-16 embryos per clutch). Clutches in the same treatment group were assayed on different days. Horizontal blue lines are the mean of each treatment.

To explore whether heart rate was influenced by nuclear estrogen receptor or GPER signaling pathways, we employed a pharmacological approach. We exposed embryos to ICI182,780 (fulvestrant), a well-characterized ERα and ERβ antagonist (23) that also acts as a GPER agonist (8). Following one-hour exposure to ICI182,780, heart rate was significantly increased (Fig. 1, mean difference in heart rate 29.81 ± 4.75 bpm). This effect was blocked by co-administration of G36, a specific GPER antagonist (24) (Fig. 1, mean difference in heart rate 4.10 ± 6.23 bpm), suggesting that estradiol increases heart rate via GPER. We also exposed embryos to G1, a specific GPER agonist with no detectable agonist activity against nuclear estrogen receptors (25), and found that heart rate increased significantly (Fig. 1, mean difference in heart rate 40.98 ± 6.35 bpm). Together, our pharmacological results suggest that GPER regulates heart rate acutely.

Curiously, the GPER inhibitor G36 by itself did not change heart rate significantly (Fig. 1, mean difference in heart rate 8.23 ± 5.13 bpm). Additionally, while G36 inhibited ICI182,780-dependent increase in heart rate, G36 failed to inhibit estradiol-dependent increase in heart rate (Fig. 1, mean difference in heart rate 41.56 ± 6.27 bpm). This could be due to off-target effects, a limitation of pharmacologic approaches, or due to differences in how G36, which was designed to inhibit human GPER, binds to zebrafish GPER. To definitively test the hypothesis that estradiol regulates heart rate via GPER, we generated GPER mutant embryos, exposed them to estrogen receptor modulators and assayed heart rate. Using CRISPR-Cas technology (26), we generated embryos with a 131 basepair deletion in the gper open reading frame (Fig. 2A, B). Embryos were viable and grossly normal, allowing us to measure heart rate (Fig. 2C, D). We exposed homozygous maternal zygotic gper mutant embryos (MZgper−/−) to estradiol or to ICI182,780 and found no increase in heart rate compared to embryos exposed to vehicle (Fig. 2E). Our results demonstrate that estradiol increases heart rate in a GPER-dependent manner. Note that zygotic gper mutants exhibited increased heart rate in response to estradiol (Fig. S1, mean difference in heart rate 29.11 ± 3.56 bpm), indicating that GPER is maternally deposited into oocytes and expressed in embryos. This is consistent with previously published results that detected gper transcript in zebrafish embryos at 1 hour post fertilization, suggesting the presence of maternally loaded gper mRNA (27).

Figure 2.
Figure 2. Abnormal heart rate in gper mutant zebrafish.

(A) Genomic DNA of gperuab102 zebrafish contains a 131 basepair deletion in the gper coding region between CRISPR guide RNA targets 1 and 2, resulting in a premature stop codon in the GPER protein. Red dashes indicate DNA deletions, mutated amino acids are shown in red. (B) Genomic DNA was harvested from individual embryos, gper was PCR amplified and separated on an agarose gel to identify deletion mutations. (C-D) 3 day post fertilization wildtype and maternal zygotic gperuab102 homozygous larvae (MZgper−/−) exhibit similar gross morphology. Images are lateral views, anterior to the left, dorsal to the top. Scale bar, 500 μm. (E) Neither estradiol (ER/GPER agonist, 3.67 μM) nor ICI182,780 (ER antagonist/GPER agonist, 10 μM) changed heart rate significantly compared to vehicle (0.1% DMSO) in MZgper−/−, two-way ANOVA followed by F test, p=0.27. (F) MZgper−/− exhibited lower basal heart rate than age-matched wildtype embryos. *, p<0.05 compared to wildtype, paired t test. Each black circle represents the mean heart rate from a single clutch of embryos (≥ 7 embryos per clutch). Clutches in the same treatment group or genotype were assayed on different days. Horizontal blue lines are the mean of each treatment.

To test whether endogenous estrogens regulate heart rate during embryonic development, we examined basal heart rate in GPER mutant embryos reared in untreated water, reasoning that if heart rate was reduced, then that would suggest that endogenous estradiol regulates heart rate via GPER. We compared heart rate in wildtype versus MZgper−/− embryos at 50 hpf and found that MZgper−/− embryos had reduced heart rate compared to wildtype (Fig. 2F, mean difference in heart rate between wildtype and mutant -30.80 ± 7.07 bpm). These results demonstrate that GPER is required for normal basal heart rate in embryos and strongly suggest that endogenous estrogens influence heart rate via GPER.

Whether GPER acts as an autonomous estrogen receptor in vivo is controversial. Previous reports suggest that GPER activity might require interaction with nuclear estrogen receptors at the membrane or that estrogens activate GPER indirectly, by binding to nuclear receptors in the cytosol that then activate downstream proteins, including GPER (15,28). To determine whether nuclear estrogen receptors influence heart rate, we generated zebrafish with loss-of-function mutations in each nuclear estrogen receptor gene: esr1 (ERα), esr2a (ERβ1) and esr2b (ERβ2) (Fig. S2-S4). All mutant embryos were viable and grossly normal, allowing us to measure heart rate (Fig. S2-S4). To test whether estradiol increases heart rate via nuclear estrogen receptors, we exposed 49 hpf esr1−/−, esr2a−/− and esr2b−/− embryos to estradiol or vehicle for one hour and assayed heart rate. Following estradiol exposure, heart rate was increased in all mutants compared to vehicle control (Fig. 3A, mean difference in heart rate between estradiol and vehicle 25.04 ± 5.68 bpm for esr1-/-, 37.23 ± 7.66 bpm for esr2a−/−, 32.48 ± 1.92 bpm for esr2b−/−), similar to what we observed when wildtype embryos were exposed to estradiol (Fig. 1). These results demonstrate that nuclear estrogen receptors are not necessary for estradiol-dependent increase in heart rate.

Figure 3.
Figure 3. Normal heart rate in nuclear estrogen receptor mutants.

(A) Homozygous mutant embryos at 49 hour post fertilization were incubated in water containing estradiol (ER/GPER agonist, 3.67 μM) or vehicle (0.1% DMSO) and heart rate was measured 1 hour post treatment. Estradiol increased heart rate compared to vehicle in zebrafish with homozygous mutations in ERα (esr1 −/−), ERβ1 (esr2a −/−), ERβ2 (esr2b −/−). *p<0.05 compared to vehicle within genotype, paired t-test. (B) Basal heart rate was measured at 50 hours post fertilization in embryos reared in untreated water. Heart rate was not significantly different in homozygous mutant (−/−) embryos compared to heterozygous (−/+) and wildtype (+/+) siblings for each esr mutant, two-way ANOVA. Each black circle represents the mean heart rate from a single clutch of embryos (4-8 embryos per clutch). Clutches in the same treatment group or genotype were assayed on different days. Horizontal blue lines are the mean of each treatment.

To test whether endogenous estrogens regulate heart rate via nuclear estrogen receptors, we bred heterozygous fish to generate embryos homozygous for mutations in either esr1, esr2a or esr2b genes and assayed heart rate in 50 hpf embryos. We observed no significant difference in basal heart rate between homozygotes, heterozygotes or wild type siblings within the same clutch (Fig. 3B, mean difference in heart rate between homozygote and wildtype −4.34 ± 1.37 for esr1, −0.46 ± 3.75 for esr2a, −2.37 ± 3.26 for esr2b; between heterozygote and wildtype −3.34 ± 1.02 for esr1, −0.91 ± 1.53 for esr2a, 0.63 ± 1.66 for esr2b). These results demonstrate that nuclear estrogen receptors are not required for the establishment of normal basal heart rate in embryos.

It is possible that the mutations generated in each nuclear estrogen receptor gene do not cause loss of functional estrogen receptor proteins. To exclude this possibility and show that esr mutants exhibit loss of functional ER proteins, we generated esr mutants on the Tg(5xERE:GFP)c262/c262 transgenic background, where green fluorescent protein (GFP) expression occurs in cells with activated nuclear estrogen receptors (5) (referred to as 5xERE:GFP). Previous studies using whole mount in situ hybridization demonstrated that esr1 is expressed in embryonic heart valves while esr2b is expressed in the liver (6), therefore we hypothesized that mutants would fail to upregulate GFP in tissues where the relevant receptor is normally expressed. We exposed 2-3 day post fertilization (dpf) 5xERE:GFP, 5xERE:GFP;esr1−/−, 5xERE:GFP;esr2a−/− and 5xERE:GFP;esr2b−/− embryos to 100 ng/ml 17β-estradiol overnight and assayed fluorescence. Consistent with esr gene expression patterns, 5xERE:GFP;esr1−/− larvae exhibited fluorescence in the liver but not in the heart (Fig. S2), whereas 5xERE:GFP;esr2b−/− larvae exhibited fluorescence in the heart but not in the liver (Fig. S4). esr2a transcript was not detected at these embryonic and larval stages (6) and, as expected, we saw no change in fluorescence between 5xERE:GFP and 5xERE:GFP;esr2a−/− (Fig. S3). We conclude that the zebrafish nuclear estrogen receptor mutants lack estrogen receptor function.

Deleterious mutations can induce genetic compensation (29), however results from the 5xERE:GFP esr mutants suggest that compensatory expression of esr genes is not occurring. For example, it is possible that in the esr1 mutant there is compensatory upregulation of esr2a and/or esr2b that masks a heart rate phenotype. If esr2a or esr2b were upregulated in esr1 mutants, then we would expect to see fluorescence in the heart in 5xERE:GFP;esr1−/− embryos. Instead, we observed no fluorescence in the hearts of 5xERE:GFP;esr1−/− embryos (Fig. S2). Similarly, we observed no ectopic fluorescence in 5xERE:GFP;esr2b−/− embryos (Fig. S4), suggesting that esr genes are not compensating for one another in the multiple zebrafish esr mutants.

To further test whether nuclear estrogen receptor signaling is influenced by GPER, we generated gper mutants on the 5xERE:GFP transgenic background and asked whether estradiol exposure reduced nuclear estrogen receptor activity in mutants compared to wildtype. Following overnight exposure to estradiol, 3 dpf 5xERE:GFP and 5xERE:GFP;MZgper−/− larvae exhibited similar fluorescence (Fig. S5). These results demonstrate that nuclear estrogen receptor transcriptional activity does not require GPER and support the hypothesis that GPER acts as an autonomous estrogen receptor in vivo.

Heart rate can be modulated by cardiomyocytes in the heart, or by cells in the central nervous system, which directly innervates the heart to modulate heart rate and also regulates the release of humoral factors, such as thyroid hormone, that bind to receptors in cardiomyocytes and regulate heart rate (30). To determine whether GPER regulates heart rate tissue autonomously, we performed whole mount in situ hybridization to test whether gper transcripts are expressed in 50 hpf zebrafish embryo hearts. We did not detect transcript in the heart or in the vasculature. In contrast, we detected gper mRNA in three discrete anatomic areas of the brain: the preoptic and olfactory areas and in the ventral hypothalamus (Fig. 4A-C). Thus, gper localization is consistent with the hypothesis that GPER acts in the brain, and not through cells in the heart, to regulate heart rate.

Figure 4.
Figure 4. gper expression in the brain.

(A-C) Whole mount colorimetric in situ hybridization was performed on wildtype embryos at 50 hours post fertilization (hpf). (A) amhc (alpha-myosin heavy chain) antisense RNA labels atrial myocardial cells in the heart (boxed). (B, C) gper antisense RNA labels a bilaterally symmetric cluster of cells in the olfactory area (white arrowheads) and preoptic area (black arrowhead) and a medial cluster of cells in the ventral hypothalamus (arrows). No label was detected in the heart. Lateral views with anterior to the left (A,B), ventral view with anterior to the top (C), scale bars = 100 μm. (D-F) Double fluorescent in situ hybridization performed on 48 hpf Tg(5xERE:GFP)c262 embryos following overnight exposure to 100 ng/ml estradiol. gfp marks cells with active nuclear estrogen receptors. Confocal images of selected Z-slices (0.975 μm) show that gper is expressed in the olfactory area (D) and preoptic area (E) in cells lacking gfp (D’’, E’’, scale bars = 50 μm). In the ventral hypothalamus (F), gper is expressed in a medial cluster of cells lacking gfp (arrows, F, F’’), whereas gper is expressed together with gfp more laterally (arrowheads, F’’, scale bar = 10 μm). In merged images, gper is magenta, gfp is green and areas of colocalization are white. Dorsal views, anterior to the top.

Genetic evidence using esr mutants suggests that GPER acts independently of nuclear estrogen receptors to regulate heart rate (Fig. 3). To further test the hypothesis that GPER acts as an autonomous estrogen receptor in vivo, we asked whether GPER and nuclear estrogen receptors are expressed in the same cells in the brain, reasoning that if GPER and nuclear estrogen receptors fail to colocalize, this would support the idea that GPER acts as an autonomous estrogen receptor in vivo. We took 5xERE:GFP embryos at 1 day post fertilization and exposed them overnight to 100 ng/ml estradiol. When the embryos reached 48 hpf, we used two color fluorescent in situ hybridization to detect gfp and gper transcripts simultaneously. Since all three nuclear estrogen receptor genes activate the 5xERE:GFP transgene, detecting gfp allows us to monitor activity of all three estrogen receptors using a single RNA probe. In the olfactory and preoptic areas, we found no colocalization between gfp and gper (Fig. 4D, E). In the ventral hypothalamus, we found a cluster of cells at the midline expressing gper but not gfp. Surrounding this region of gper-positive cells was a bilaterally symmetric ‘U’-shaped labeling pattern of cells expressing both gper and gfp (Fig. 4F). Thus, GPER and nuclear estrogen receptors are expressed in unique and overlapping cells in the brain, supporting the hypothesis that GPER can act independently of nuclear estrogen receptors in vivo.

Because MZgper−/− fish are viable, we asked whether loss of GPER and reduced basal heart rate in embryos might have an effect on adult heart morphology. We dissected hearts from 8-9 month old adult wildtype and MZgper−/− zebrafish. Female mutants had a mean 47% reduction in antrioventricular (AV) valve width compared to wildtype (Fig. 5E; MZgper−/− 33.5 ± 6.35 μm (mean ± standard deviation), n=10; wildtype 62.49 ± 18.53, n=9, p=0.0002 Student’s t test), while male mutants had a mean 40% reduction in AV valve width compared to wildtype (Fig. 5F; MZgper−/− 29.39 ± 6.32, n=7; wildtype 49.14 ± 9.68, n=5, p=0.0016 Student’s t test). We also analyzed the single thickest part of AV valve leaflets in each heart and found that mutant valves were on average 45% thinner at their thickest point than wildtype valves from fish of the same sex (female MZgper−/− 42.93 ± 10.38 vs wildtype 77.97 ± 24.32 μm, p=0.0006 Student’s t test; male MZgper−/− 38.17 ± 11.62 vs wildtype 67.19 ± 24.77, p=0.02 Student’s t test). These results are consistent with previous observations that eliminating blood flow in embryos impairs heart valve formation (31). Together, these findings demonstrate a correlation between reduced blood flow in embryos and reduced valve thickness in adults and suggest that GPER regulation of heart rate is important for proper heart valve maturation.

Figure 5.
Figure 5. GPER mutants exhibit decreased atrioventricular (AV) valve width in adulthood.

(A-L) Representative images of MZ gper −/− and wildtype (wt) hearts. (A-D) 5 μm coronal sections through the heart, stained with H&E. Atrium (A), ventricle (V), and bulbus arteriosus (BA) indicated with black arrows, AV valve leaflets are indicated with red arrows. Black boxes show the portion of the image that is digitally enlarged in A’-D’ to highlight AV valve morphology. Scale bars in A-D represent 200μm; scale bars in A’-D’ represent 50μm. (E,F) Graphs comparing average and maximum AV valve width between MZgper −/− and wildtype hearts in females (E) and males (F). Width was measured at the widest part of the valve over multiple coronal sections from the middle of the heart, perpendicular to the long axis of the valve, and corrected for standard length of each fish. Each circle respresents measurements from a single fish, horizontal black lines are the mean width from each genotype. *p < 0.05; ** p < 0.01; *** p < 0.001, unpaired Student’s t test.

DISCUSSION

Our results support the hypothesis that GPER acts as an autonomous estrogen receptor in vivo. Previous reports using cultured cells demonstrated that fluorescently labeled or isotopic estradiol specifically binds membranes from cells expressing GPER (8,9). Additionally, estradiol exposure increased cyclic AMP and calcium levels in HEK293 and COS7 cells in a GPER-dependent manner (8,9), while estradiol exposure increased phosphoinositide 3-kinase activity in SKBR3 breast cancer cell line in a GPER-dependent manner (8). However, because these studies utilized cells that either express artificially high levels of GPER or are tumorigenic, the findings do not address whether GPER acts as an estrogen receptor in vivo under normal physiologic conditions. Our genetic and pharmacologic results strongly suggest that GPER is an estrogen receptor in vivo. If estradiol was binding to ERα or ERβ, and these receptors activated GPER, then we would expect to see no increase in heart rate in esr1, esr2a or esr2b mutants following exposure to estradiol. Instead, all esr mutants responded normally to estradiol exposure (Fig. 3), suggesting that ER and GPER signaling pathways are distinct in this context. Consistent with these results, we found gper transcript expressed in cells in the brain that lack nuclear estrogen receptor activity (Fig. 4), further supporting the hypothesis that GPER responds to estrogens in the absence of nuclear estrogen receptors in vivo. The extent to which ER and GPER signaling pathways interact likely depends on cell type, developmental stage, sex and/or pathology. We argue that studying the influence of estrogens on heart rate in zebrafish embryos is a powerful in vivo system where GPER activity is dissociated from classical nuclear estrogen receptor signaling.

Between 2 and 5 dpf, zebrafish heart rate normally increases (32,33). Our results support the idea that endogenous estradiol regulates this increase in heart rate. The fact that GPER mutant embryos have lower basal heart rate compared to wildtype embryos implicates endogenous estradiol. Additionally, a recent HPLC analysis of endogenous estradiol concentration in zebrafish embryos found that estradiol concentrations increased from 137 pg/embryo at 48 hpf to 170 pg/embryo at 72 hpf (34). Taken together, these results support the hypothesis that endogenous estradiol regulates heart rate in zebrafish embryos and larvae. The source of embryonic estradiol, whether synthesized by the embryo or maternally deposited in the yolk, is not clear.

Blood flow in embryos is a critical determinant of cardiovascular development (35). Zebrafish embryos in which blood flow is drastically reduced or eliminated exhibited a valve dysgenesis phenotype (31), yet because of the drastic reduction in blood flow these embryos are not viable and were not examined at later stages of development. MZgper−/− embryos exhibit a ~20% reduced heart rate compared to wildtype, yet are viable, allowing us to examine the adult heart following reduced basal heart rate in embryos. Based on the heart rate phenotype in MZgper−/− embryos and the detection of gper transcripts in the brain but not the heart, we propose that the reduced heart rate in GPER mutant embryos causes a reduction in blood flow that prevents heart valves from maturing properly and manifests in adults as thinner atrioventricular valves compared to wildtype. It is not known whether gper is expressed in the heart at later stages of development, where it could directly influence valve cell division and/or survival. How might GPER activity in the brain regulate heart rate? Previous work suggests that GPER influences neurotransmitter release (36), meaning that GPER activity could trigger neuronal activity that leads to release of humoral factors that regulate heart rate. Alternatively, neurons that express GPER could be part of a circuit that ultimately terminates in motor neurons that innervate the heart and regulate contractions. It is not known whether the zebrafish heart is innervated by 50 hpf.

While our results illuminate GPER signaling in the context of embryonic heart rate, it is not clear to what extent GPER influences heart rate at later stages of development. In adult mice with mutations in GPER, there was no significant difference in basal heart rate between mutant and wildtype mice of either sex (16, 17, 37). It is possible that GPER regulates heart rate in embryos but not in adults. Additionally, heart rate in GPER mutant mice was assayed using general anesthesia. which is known to depress heart rate compared to conscious mice (38). Anesthesia may mask the effect of GPER on basal heart that we observe in conscious animals. We also cannot exclude the possibility that the effects of GPER on heart rate are specific for zebrafish.

In summary, this study identified a role for GPER in the regulation of embryonic heart rate. The zebrafish estrogen receptor mutants we developed enable experiments to rapidly and conclusively identify the causative estrogen receptor associated with any estrogen signaling phenotype, as demonstrated with the estradiol-dependent increase in heart rate reported here. This has significant implications for studies of estrogenic environmental endocrine disruptors, which are frequently tested on zebrafish to identify effects on embryonic development, organ formation and function (39). Zebrafish estrogen receptor mutants can now be used to determine whether such effects are specific for estrogen receptors and to identify the precise receptor target. Our results also establish a need to consider the impact on cardiac function when considering the toxicity of estrogenic environmental endocrine disruptors.

Figure S1:
Figure S1: Zygotic gper mutant embryos are sensitive to estradiol.

Zygotic homozygous gper mutant embryos were incubated in water containing estradiol (ER/GPER agonist, 3.67 μM) or vehicle control (0.1% DMSO) at 49 hours post fertilization and heart rates were measured 1 hour post treatment. *, p<0.05 compared to vehicle, paired t test. Each black circle represents the mean heart rate from a single clutch of embryos (≥ 6 embryos per clutch). Clutches in the same treatment group were assayed on different days. Horizontal blue lines are the mean of each treatment.

Figure S2:
Figure S2: Generation and validation of esr1 mutant zebrafish.

(A) Genomic DNA of esr1uab118 zebrafish contains a 4 basepair deletion in the esr1 coding region, resulting in a premature stop codon in the Esr1 (ERα) protein. Nucleotide deletions are shown as red dashes, amino acid mutations are in red. Map indicates site of frameshift mutation and premature stop codon (AF-1, activating function 1 domain; DBD, DNA binding domain; LBD, ligand binding domain; AF-2, activating function 2 domain). (B) High resolution melting curve analysis was used to distinguish mutants from wildtype. Curves represents DNA amplified from a wildtype AB (black) or esr1uab118 mutant zebrafish (cyan). (C) Strategy for validating zebrafish estrogen receptor mutants using transgenic 5xERE:GFP zebrafish. Mutants were generated on a transgenic background where estrogen receptor (ER) transcriptional activity is marked by green fluorescent protein (GFP) expression. Following exposure to estradiol, loss-of-function mutants should exhibit reduced fluorescence in cells expressing esr1. (D-K) 2-day post fertilization embryos were exposed to 367 nM (100 ng/mL) estradiol, live fluorescent images (D, F, H, J) and corresponding brightfield images (E, G, I, K) were taken at 3 d. 5xERE:GFPc262;esr1uab118 homozygous larvae (esr1 −/−) exhibit normal morphology, but lack fluorescence in heart valves, wheres heterozygotes (esr1 −/+) exhibit fluorescent heart valves. High magnification images of the heart are shown in H-K. Arrows indicate heart valves, arrow head indicates liver. Images are lateral views, anterior to the left, dorsal to the top. Scale bars, 500 μm (D-G), 100 μm (H-K).

Figure S3:
Figure S3: Generation of esr2a mutant zebrafish.

(A) Genomic DNA of esr2auab134 zebrafish contains an 2 basepair deletion (red dashes) in the esr2a coding region, resulting in a premature stop codon in the Esr2a (ERβ1) protein. Amino acid mutations are in red. Map indicates frameshift mutation and premature stop codon in the Esr2a protein. AF-1, activating function 1 domain; DBD, DNA binding domain; LBD, ligand binding domain; AF-2, activating function 2 domain. (B) High resolution melting curve analysis was used to distinguish mutants from wildtype. Curves represent DNA amplified from a wildtype AB (black) or esr2auab134 mutant zebrafish (cyan). (C-F) 5xERE:GFPc262 and 5xERE:GFPc262;esr2auab134 (esr2a −/−) 3-day post fertilization (d) larvae were exposed to 367 nM (100 ng/mL) estradiol. Live fluorescent images (C, E) and corresponding brightfield images (D, F) were captured at 4 d. esr2a −/− larvae exhibit normal morphology and fluorescence, consistent with data demonstrating that esr2a is not expressed during these developmental stages. Arrows indicate heart valves, arrow head indicates liver. Images are lateral views, anterior to the left, dorsal to the top. Scale bar, 500 μm

Figure S4:
Figure S4: Generation and validation of esr2b mutant zebrafish.

(A) Genomic DNA of esr2buab127 zebrafish contains a 5 basepair deletion (red) in the esr2b coding region, resulting in a premature stop codon in the Esr2b (ERβ2) protein. Amino acid mutations are in red. Map indicates frameshift mutation and premature stop codon in the Esr2b protein. AF-1, activating function 1 domain; DBD, DNA binding domain; LBD, ligand binding domain; AF-2, activating function 2 domain. (B) High resolution melting curve analysis was used to distinguish mutants from wildtype. Curves represents DNA amplified from a wildtype AB (black) or esr2buab127 mutant zebrafish (cyan). (C-F) 5xERE:GFPc262;esr2buab127 3-day post fertilization (d) larvae were exposed to 367 nM (100 ng/mL) estradiol. Live fluorescent images (C, E) and corresponding brightfield images (D, F) were captured at 4 d. 5xERE:GFPc262;esr2buab127 homozygous larvae (esr2b −/−) exhibit normal morphology, but lack fluorescence in the liver. Arrows indicate heart valves, arrow head indicates liver. Images are lateral views, anterior to the left, dorsal to the top. Scale bar = 100 μm.

Figure S5:
Figure S5: Nuclear estrogen receptor transcriptional activityis normal in gper mutant zebrafish.

(A-H)Maternal zygotic gperuab 102 homozygous larve on the (5xERE:GFP)c262 transgenic background (MZgper−/−) were exposed to 367 nM (100 ng/ml)estradiol at 2-days post fertilization (2 d). flurescence (A, C, E, G)and corresponding brightfield images (B, D, F, H)were taken at 3 d. Flurescence in the heart valves (arrows) and liver (arrow heads)is similar between MZgper−/− and wildtype larvae.C, D, H, G High magnification images of heart. Images are lateral views, anterior to the left, dorsal to the top. Scale bars, 500μm (C-F), 100μM (G-J).

Acknowledgements:

We thank J.L. King for technical assistance and S. Farmer and the staff of the UAB zebrafish facility for animal care. Funding: This work was supported by start-up funds from UAB (to D.A.G.) and by funds from NIH T32GM008361 (to J.P.S.) and T32GM008111 (to S.N.R.). Author contributions: S.N.R., J.P.S, H.E.E. and D.A.G. performed experiments and analyzed experimental data. X.C. performed statistical analyses of heart rate data. S.N.R. and D.A.G. conceived the project, D.A.G. supervised the project. All authors contributed to writing the paper and read and approved the final manuscript. Competing interests: The authors declare that they have no competing interests.

References

  1. 1.↵
    D. Staudt, D. Stainier, Uncovering the Molecular and Cellular Mechanisms of Heart Development Using the Zebrafish. Annual Review of Genetics 46, 397-418 (2012).
    OpenUrlCrossRefPubMedWeb of Science
  2. 2.↵
    X. Liu et al., Identification of a membrane estrogen receptor in zebrafish with homology to mammalian GPER and its high expression in early germ cells of the testis. Biology of reproduction 80, 1253-1261 (2009).
    OpenUrlCrossRefPubMedWeb of Science
  3. 3.
    A. Menuet et al., Molecular characterization of three estrogen receptor forms in zebrafish: binding characteristics, transactivation properties, and tissue distributions. Biology of reproduction 66, 1881-1892 (2002).
    OpenUrlCrossRefPubMedWeb of Science
  4. 4.↵
    P. Thomas et al., Conserved estrogen binding and signaling functions of the G protein-coupled estrogen receptor 1 (GPER) in mammals and fish. Steroids 75, 595-602 (2010).
    OpenUrlCrossRefPubMedWeb of Science
  5. 5.↵
    D. A. Gorelick, M. E. Halpern, Visualization of Estrogen Receptor Transcriptional Activation in Zebrafish. Endocrinology 152, 2690-2703 (2011).
    OpenUrlCrossRefPubMedWeb of Science
  6. 6.↵
    D. A. Gorelick, L. R. Iwanowicz, A. L. Hung, V. S. Blazer, M. E. Halpern, Transgenic zebrafish reveal tissue-specific differences in estrogen signaling in response to environmental water samples. Environmental health perspectives 122, 356-362 (2014).
    OpenUrlPubMed
  7. 7.↵
    R. Evans, A transcriptional basis for physiology. Nat Med 10, 1022-1026 (2004).
    OpenUrlCrossRefPubMedWeb of Science
  8. 8.↵
    C. M. Revankar, D. F. Cimino, L. A. Sklar, J. B. Arterburn, E. R. Prossnitz, A Transmembrane Intracellular Estrogen Receptor Mediates Rapid Cell Signaling. Science 307, 1625-1630 (2005).
    OpenUrlAbstract/FREE Full Text
  9. 9.↵
    P. Thomas, Y. Pang, E. J. Filardo, J. Dong, Identity of an estrogen membrane receptor coupled to a G protein in human breast cancer cells. Endocrinology 146, 624-632 (2005).
    OpenUrlCrossRefPubMedWeb of Science
  10. 10.↵
    C. E. Navarro et al., Regulation of cyclic adenosine 3’,5’-monophosphate signaling and pulsatile neurosecretion by Gi-coupled plasma membrane estrogen receptors in immortalized gonadotropin-releasing hormone neurons. Mol Endocrinol 17, 1792-1804 (2003).
    OpenUrlCrossRefPubMedWeb of Science
  11. 11.
    Q. Wu et al., Point mutations in the ERalpha Galphai binding domain segregate nonnuclear from nuclear receptor function. Mol Endocrinol 27, 2-11 (2013).
    OpenUrlCrossRefPubMedWeb of Science
  12. 12.
    P. Kumar et al., Direct interactions with G alpha i and G betagamma mediate nongenomic signaling by estrogen receptor alpha. Mol Endocrinol 21, 1370-1380 (2007).
    OpenUrlCrossRefPubMedWeb of Science
  13. 13.↵
    C. S. Watson et al., Estrogen‐ and xenoestrogen-induced ERK signaling in pituitary tumor cells involves estrogen receptor-alpha interactions with G protein‐ alphai and caveolin I. Steroids 77, 424-432 (2012).
    OpenUrlCrossRefPubMedWeb of Science
  14. 14.↵
    A. Vivacqua et al., G protein-coupled receptor 30 expression is up-regulated by EGF and TGF alpha in estrogen receptor alpha-positive cancer cells. Mol Endocrinol 23, 1815-1826 (2009).
    OpenUrlCrossRefPubMedWeb of Science
  15. 15.↵
    E. R. Levin, G Protein-Coupled Receptor 30: Estrogen Receptor or Collaborator? Endocrinology 150, 1563-1565 (2009).
    OpenUrlCrossRefPubMedWeb of Science
  16. 16.↵
    M. Delbeck et al., Impaired left-ventricular cardiac function in male GPR30‐ deficient mice. Molecular medicine reports 4, 37-40 (2011).
    OpenUrl
  17. 17.↵
    U. E. Martensson et al., Deletion of the G protein-coupled receptor 30 impairs glucose tolerance, reduces bone growth, increases blood pressure, and eliminates estradiol-stimulated insulin release in female mice. Endocrinology 150, 687-698 (2009).
    OpenUrlCrossRefPubMedWeb of Science
  18. 18.↵
    E. Haas et al., Regulatory role of G protein-coupled estrogen receptor for vascular function and obesity. Circ Res 104, 288-291 (2009).
    OpenUrlAbstract/FREE Full Text
  19. 19.↵
    M. R. Meyer et al., G protein-coupled estrogen receptor protects from atherosclerosis. Sci Rep 4, 7564 (2014).
    OpenUrlCrossRefPubMed
  20. 20.↵
    T. Takahashi, Juvenile hermaphroditism in the zebrafish, Brachydanio rerio. Bull Fac Fish Hokkaido Univ 28, 57-65 (1977).
    OpenUrl
  21. 21.↵
    F. S. vom Saal, Variation in phenotype due to random intrauterine positioning of male and female fetuses in rodents. Journal of reproduction and fertility 62, 633-650 (1981).
    OpenUrlAbstract/FREE Full Text
  22. 22.↵
    F. S. vom Saal, Sexual differentiation in litter-bearing mammals: influence of sex of adjacent fetuses in utero. Journal of animal science 67, 1824-1840 (1989).
    OpenUrlCrossRefPubMedWeb of Science
  23. 23.↵
    J. F. Robertson, ICI 182,780 (Fulvestrant)‐‐the first oestrogen receptor down‐ regulator‐‐current clinical data. British journal of cancer 85 Suppl 2, 11-14 (2001).
    OpenUrlCrossRefPubMed
  24. 24.↵
    M. K. Dennis et al., Identification of a GPER/GPR30 antagonist with improved estrogen receptor counterselectivity. The Journal of steroid biochemistry and molecular biology 127, 358-366 (2011).
    OpenUrlCrossRefPubMed
  25. 25.↵
    C. G. Bologa et al., Virtual and biomolecular screening converge on a selective agonist for GPR30. Nat Chem Biol 2, 207-212 (2006).
    OpenUrlCrossRefPubMedWeb of Science
  26. 26.↵
    L. E. Jao, S. R. Wente, W. Chen, Efficient multiplex biallelic zebrafish genome editing using a CRISPR nuclease system. Proc Natl Acad Sci U S A 110, 13904-13909 (2013).
    OpenUrlAbstract/FREE Full Text
  27. 27.↵
    B. S. Jayasinghe, D. C. Volz, Aberrant ligand-induced activation of G protein‐ coupled estrogen receptor 1 (GPER) results in developmental malformations during vertebrate embryogenesis. Toxicol Sci 125, 262-273 (2012).
    OpenUrlCrossRefPubMedWeb of Science
  28. 28.↵
    G. Langer et al., A critical review of fundamental controversies in the field of GPR30 research. Steroids 75, 603-610 (2010).
    OpenUrlCrossRefPubMedWeb of Science
  29. 29.↵
    A. Rossi et al., Genetic compensation induced by deleterious mutations but not gene knockdowns. Nature 524, 230-233 (2015).
    OpenUrlCrossRefPubMed
  30. 30.↵
    I. Klein, K. Ojamaa, Thyroid hormone and the cardiovascular system. The New England journal of medicine 344, 501-509 (2001).
    OpenUrlCrossRefPubMedWeb of Science
  31. 31.↵
    J. Vermot et al., Reversing blood flows act through klf2a to ensure normal valvulogenesis in the developing heart. PLoS Biol 7, e1000246 (2009).
    OpenUrlCrossRefPubMed
  32. 32.↵
    E. De Luca et al., ZebraBeat: a flexible platform for the analysis of the cardiac rate in zebrafish embryos. Sci. Rep., (2014).
  33. 33.↵
    E. de Pater et al., Distinct phases of cardiomyocyte differentiation regulate growth of the zebrafish heart. Development 136, 1633-1641 (2009).
    OpenUrlAbstract/FREE Full Text
  34. 34.↵
    K. M. Alharthy, F. F. Albaqami, C. Thornton, J. Corrales, K. L. Willett, Mechanistic evaluation of benzo[a]pyrene’s developmental toxicities mediated by reduced Cyp19a1b activity. Toxicol Sci, (2016).
  35. 35.↵
    F. Boselli, J. B. Freund, J. Vermot, Blood flow mechanics in cardiovascular development. Cellular and molecular life sciences: CMLS 72, 2545-2559 (2015).
    OpenUrl
  36. 36.↵
    E. M. Waters et al., G-protein-coupled estrogen receptor 1 is anatomically positioned to modulate synaptic plasticity in the mouse hippocampus. The Journal of neuroscience: the official journal of the Society for Neuroscience 35, 2384-2397 (2015).
    OpenUrlAbstract/FREE Full Text
  37. 37.
    L. Meoli et al., Sex‐ and age-dependent effects of Gpr30 genetic deletion on the metabolic and cardiovascular profiles of diet-induced obese mice. Gene 540, 210-216 (2014).
    OpenUrlCrossRefPubMed
  38. 38.↵
    B. J. Janssen et al., Effects of anesthetics on systemic hemodynamics in mice. American journal of physiology. Heart and circulatory physiology 287, H1618-1624 (2004).
    OpenUrlCrossRefPubMedWeb of Science
  39. 39.↵
    C. W. McCollum, N. A. Ducharme, M. Bondesson, J. A. Gustafsson, Developmental toxicity screening in zebrafish. Birth defects research. Part C, Embryo today: reviews 93, 67-114 (2011).
    OpenUrl
  40. 40.↵
    M. Westerfield, The Zebrafish Book. A Guide for the Laboratory Use of Zebrafish (Danio Rerio). (University of Oregon Press, Eugene, OR, ed. 4th, 2000).
  41. 41.↵
    J. A. Gagnon et al., Efficient Mutagenesis by Cas9 Protein-Mediated Oligonucleotide Insertion and Large-Scale Assessment of Single-Guide RNAs. PLoS One 9, e98186 (2014).
    OpenUrlCrossRefPubMed
  42. 42.↵
    J. M. Parant, S. A. George, R. Pryor, C. T. Wittwer, H. J. Yost, A rapid and efficient method of genotyping zebrafish mutants. Dev Dyn 238, 3168-3174 (2009).
    OpenUrlCrossRefPubMed
  43. 43.↵
    J. S. Waxman, B. R. Keegan, R. W. Roberts, K. D. Poss, D. Yelon, Hoxb5b acts downstream of retinoic acid signaling in the forelimb field to restrict heart field potential in zebrafish. Dev Cell 15, 923-934 (2008).
    OpenUrlCrossRefPubMed
  44. 44.↵
    C. Thisse, B. Thisse, High-resolution in situ hybridization to whole-mount zebrafish embryos. Nature protocols 3, 59-69 (2008).
    OpenUrl
  45. 45.↵
    G. Lauter, I. Soll, G. Hauptmann, Multicolor fluorescent in situ hybridization to define abutting and overlapping gene expression in the embryonic zebrafish brain. Neural Dev 6, 10 (2011).
    OpenUrl
  46. 46.↵
    P. D. Vize, K. E. McCoy, X. Zhou, Multichannel wholemount fluorescent and fluorescent/chromogenic in situ hybridization in Xenopus embryos. Nature protocols 4, 975-983 (2009).
    OpenUrl
  47. 47.↵
    D. M. Parichy, M. R. Elizondo, M. G. Mills, T. N. Gordon, R. E. Engeszer, Normal table of postembryonic zebrafish development: staging by externally visible anatomy of the living fish. Dev Dyn 238, 2975-3015 (2009).
    OpenUrlCrossRefPubMedWeb of Science
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Posted November 21, 2016.
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G protein-coupled estrogen receptor regulates heart rate in zebrafish embryos
Shannon N Romano, Hailey E Edwards, J Paige Souder, Xiangqin Cui, Daniel A Gorelick
bioRxiv 088955; doi: https://doi.org/10.1101/088955
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