**Annexin A1 Regulates NLRP3 Inflammasome Activation and Modifies Lipid Release Profile in Isolated Peritoneal Macrophages**

**José Marcos Sanches 1,2, Laura Migliari Branco 3, Gustavo Henrique Bueno Duarte 4, Sonia Maria Oliani 5, Karina Ramalho Bortoluci 3, Vanessa Moreira <sup>6</sup> and Cristiane Damas Gil 1,5,\***


Received: 14 February 2020; Accepted: 6 April 2020; Published: 9 April 2020

**Abstract:** Annexin A1 (AnxA1) is a potent anti-inflammatory protein that downregulates proinflammatory cytokine release. This study evaluated the role of AnxA1 in the regulation of NLRP3 inflammasome activation and lipid release by starch-elicited murine peritoneal macrophages. C57bl/6 wild-type (WT) and AnxA1-null (AnxA1-/- ) mice received an intraperitoneal injection of 1.5% starch solution for macrophage recruitment. NLRP3 was activated by priming cells with lipopolysaccharide for 3 h, followed by nigericin (1 h) or ATP (30 min) incubation. As expected, nigericin and ATP administration decreased elicited peritoneal macrophage viability and induced IL-1β release, more pronounced in the AnxA1-/- cells than in the control peritoneal macrophages. In addition, nigericin-activated AnxA1-/- macrophages showed increased levels of NLRP3, while points of co-localization of the AnxA1 protein and NLRP3 inflammasome were detected in WT cells, as demonstrated by ultrastructural analysis. The lipidomic analysis showed a pronounced release of prostaglandins in nigericin-stimulated WT peritoneal macrophages, while ceramides were detected in AnxA1-/- cell supernatants. Different eicosanoid profiles were detected for both genotypes, and our results suggest that endogenous AnxA1 regulates the NLRP3-derived IL-1β and lipid mediator release in macrophages.

**Keywords:** inflammation; nigericin; pyroptosis; mass spectrometry; lipidomics

### **1. Introduction**

Annexin A1 (AnxA1) is a 37-kDa protein that can mimic the anti-inflammatory action of glucocorticoids by inhibiting eicosanoid and phospholipase A2 synthesis, affecting components of inflammatory reaction and arachidonic acid release [1,2]. The ability of AnxA1 to down-modulate cellular and molecular processes of inflammation contributes to tissue homeostasis and reprogramming of macrophages [3]. The development of the AnxA1-null mice (AnxA1-/- ) strain has allowed for a better understanding of the role of endogenous AnxA1 protein in leukocyte biology and the inflammatory

process. In models of inflammation induced by carrageenan or zymosan, AnxA1-/- animals exhibit an exacerbated response characterized by a prominent leukocyte influx and IL-1β release [4]. In addition, macrophages from AnxA1-/- mice demonstrate reduced ability to phagocytose non-opsonized zymosan particles [5], and show higher TLR4 mRNA expression and IL-1β production after lipopolysaccharide (LPS) stimulation than wild-type cells [6]. These data demonstrate that AnxA1 exerts a negative inflammatory response through its down-modulation effects on macrophage cells, which are important leukocytes in an innate response.

In addition, the stimulation of a P2 × 7 receptor (P2X7R) in resting and M2 macrophages, but not in M1 cells, provokes the rapid release of AnxA1 through its exposure with phosphatidylserine to the outer plasma membrane leaflet [7]. Also, the release of AnxA1 after P2X7R activation is not affected in inflammasome knockout macrophages, suggesting that its release is independent of caspase-1 activation. Considering that P2X7R activation is necessary to promote the assembly of the NLRP3 inflammasome and cytokine release [8], the release of AnxA1 represents another P2X7R macrophage signaling pathway for the resolution of the inflammation.

Nucleotide-binding oligomerization domain (NOD)-like receptor family pyrin domain containing 3 (NLRP3 or NALP3) is a cytoplasmic sensor that oligomerizes to form a platform known as inflammasome, a protein complex that controls the release of IL-1β and IL-18 by activating caspase-1 [9]. In macrophages, NLRP3 inflammasome activation can be triggered by the pore-forming ionophore nigericin, extracellular ATP, and crystalline substances that induce pyroptosis, a type of cell death [10]. In addition, previous research has shown that some fatty acid-derived lipids, such as the prostaglandin E2 (PGE2), regulates NLRP3 and the maturation of IL-1β. PGE2 can be associated with inhibition of the NLRP3 activation in human macrophages through the EP4 receptor and by the EP2 receptor in murine macrophages, decreasing IL-1β maturation [11,12]. NLRP3 activation driven by damage-associated molecular patterns is also associated with the production and release of several lipidic mediators, such as eicosanoids derived from arachidonic acids and ceramides, which cause cellular and systemic damages to the organism [13]. Eicosanoids, such as prostaglandins and leukotrienes, are lipid mediators that play a crucial role in initiating the acute inflammatory response [14]. Systemic activation of inflammasomes leads to the production of large amounts of eicosanoids in several minutes, contributing to rapid initiation of inflammation characterized by increased vascular permeability, culminating in pathological inflammatory effects [15].

Inflammasome activation is vital for the control of infections and the regulation of metabolic processes and immune responses [16]. However, altered functions of these platforms are implicated in the pathogenesis of several human diseases. Therefore, investigations that highlight novel signaling components that regulate inflammasome activation are crucial to prevent or treat human infections/inflammatory diseases. Considering that AnxA1 is a potent anti-inflammatory protein that down-regulates proinflammatory mediator release, phospholipase A2 and, consequently, the critical cascade pathways of eicosanoid production such as that of cyclooxygenase 2 (COX-2) [17,18], this study evaluated its role in regulating the NLRP3 inflammasome and lipid release by macrophages.

### **2. Materials and Methods**

### *2.1. Animals*

Male C57BL/6 wild-type (WT) and AnxA1-null (AnxA1-/- ) mice aged 7–8 weeks and weighing 20–25 g were kept in cages (*n* = 4) in a temperature-controlled environment (22–25 ◦C) with a 12-h light-dark cycle. They received water and food ad libitum. All animal procedures were approved by the Ethics Committee in Animal Experimentation of the Federal University of São Paulo-UNIFESP (CEUA agreement number: N◦ 6493130318) and by the Internal Biosafety Commission (CIBio).

### *2.2. Cell Culture and Treatments*

Lipopolysaccharide (LPS), nigericin, and ATP were obtained from InvivoGen (San Diego, CA, USA). LPS and ATP were reconstituted in endotoxin-free water and nigericin in 100% ethanol. The stock solutions were diluted in endotoxin-free water to prepare intermediate concentration solutions, stored at −20 ◦C.

WT and AnxA1-/- peritoneal macrophages were obtained by the intraperitoneal injection of a 1.5% starch solution (Sigma Aldrich, St. Louis, MO, USA) in sterile PBS, and after four days, cells were collected by peritoneal wash. Differential cell counts were made on Diff–Quick-stained cell smears prepared by cytocentrifugation. The macrophage population obtained was more than 85% pure and at least 90% viable, as examined by trypan blue exclusion. Additionally, macrophage morphology was confirmed by ultrastructural analysis using transmission electron microscopy. Peritoneal cells (1 <sup>×</sup> 106 cells/well) were cultured in Opti-MEM (Thermo Fisher Scientific, Waltham, MA, USA) overnight at 37 ◦C under an atmosphere of 5% CO2. Experiments were performed in triplicate in 24-well plates. WT and AnxA1-/- cells were primed with LPS (500 ng/mL for 3 h) followed by stimulation with nigericin (10 μM for 1 h) or ATP (5 mM, 30 min) to activate the NLRP3 inflammasome.

### *2.3. Analysis of Cell Viability and IL-1*β *Release*

Cell viability was determined by a 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. After the treatment process, the supernatants were discarded, and the RPMI medium (Invitrogen, Gibco, Portland, OR, USA) with 10% MTT solution (5 mg/mL) was added to the cells. After incubating the cells for 4 h at 37 ◦C under a 5% CO2 atmosphere, 300 μL of dimethyl sulfoxide (DMSO; Sigma Aldrich, St. Louis, MO, USA) was added to each well (24-well plate), and 100 μL triplicates of the same sample were transferred to a 96-well plate. The spectrophotometric absorbance values at 490 nm were determined. The percentage of viable cells was calculated by optical density normalization for LPS-stimulated cells only.

IL-1β levels were tested in culture supernatants by an enzyme-linked immunosorbent assay (ELISA) using a commercially available immunoassay kit (BioLegend, San Diego, CA, USA), according to the manufacturer's instructions. All experiments were conducted in duplicate, and the data were expressed as the mean ± standard error of the mean (SEM) of protein (pg/mL).

### *2.4. Western Blot Analysis*

After nigericin and ATP stimulation, the supernatant was removed, and cells were washed three times with sterile PBS, and to each well, 50 μL of lysis buffer was added for cell lysis and protein extraction. Equal amounts of supernatants and cell extracts were loaded onto a 15% sodium dodecyl sulphate-polyacrylamide gel with appropriate molecular weight markers (Bio-Rad Life Science, Hercules, CA, USA) for electrophoresis and transferred to ECL Hybond nitrocellulose membranes. Reversible protein staining of the membranes with 0.1% Ponceau-S in 5% acetic acid (Santa Cruz Biotechnology, Dallas, TX, USA) was used to verify protein transfer. Membranes were incubated 30 min in 5% milk in Tris-buffered saline (TBS) prior to incubation with the antibodies. Primary antibodies were rabbit polyclonal anti-AnxA1 (Invitrogen-Thermo Fisher Scientific, Waltham, MA, USA; 1:1000), goat polyclonal anti-IL-1β (R&E Systems, MN, USA; 1:500), mouse monoclonal anti-caspase-1 (Santa Cruz Biotechnology Dallas, TX, USA; 1:200), and polyclonal rabbit anti-β-actin (Cell Signaling Technology, Beverly, MA, USA; 1:1000), all diluted in TBS. The membranes were then incubated with the appropriate peroxidase-conjugated secondary antibodies (Millipore Corporation, Burlington, MA, USA; 1:2500). Finally, membranes were washed for 15 min with TBS, and immunoreactive proteins were detected (Clarity™ Western ECL Substrate; Bio-Rad, Hercules, CA, USA) using a GeneGnome5 chemiluminescence detection system (SynGene, Cambridge, UK).

### *2.5. Ultrastructural Immunocytochemical Analysis*

WT and AnxA1-/- nigericin-stimulated macrophages were fixed in 4% paraformaldehyde and 0.5% glutaraldehyde, 0.1% sodium cacodylate buffer (pH 7.4) for 24 h at 4 ◦C. Samples were washed in sodium cacodylate, dehydrated through a graded methanol series, and embedded in LR Gold (Sigma Aldrich Corp., St. Louis, MO, USA).

To detect AnxA1 and NLRP3, ultrathin macrophage sections (~90 nm) were submitted for immunocytochemistry, as previously described [19]. To detect the proteins, the sheep polyclonal antibody anti-AnxA1 (1:200) and rabbit polyclonal antibody anti-NLRP3 (1:200; Cusabio, Houston, TX, USA), following a donkey anti-sheep IgG and goat anti-rabbit IgG antibody (1:50) conjugated to 10-nm and 20-nm colloidal gold (British Biocell, Cardiff, UK), respectively, were used. Ultrathin sections were stained with uranyl acetate and lead citrate and examined using a ZEISS EM900 electron microscope (Carl Zeiss, Jena, Germany). Randomly photographed sections of macrophages were analyzed using Axiovision software. The density of immunogold (number of gold particles/μm2) was calculated and reported as the mean ± SEM of 20–40 cells per experimental condition.

### *2.6. Lipidomic Analysis*

After treatments, WT and AnxA1-/- cell supernatants (1 <sup>×</sup> 106 cells/well) were collected and stored at −80 ◦C until sample processing. For lipid extraction, each sample was randomized and resuspended in 1 mL of 1:2 CHCl3: MeOH solution (Sigma Aldrich, Basel, Switzerland), followed by the addition of 0.33 mL CHCl3 and 0.33 mL deionized water. The solution was stirred for 5 min, then centrifuged at 13,000 rpm for 5 min. Derived-organic fractions with lipids were collected from the bottom layer of the tubes and transferred to 1.5-mL glass tubes. These fractions were dried in a SpeedVac Savant SPD131DDA concentrator (Thermo Scientific) for 30 min at 30 ◦C and stored at −80 ◦C. Mass spectrometric analysis was performed in an ultra-high-performance liquid chromatography (UHPLC) Agilent 1290 Infinity system (Agilent, Santa Clara, CA, USA) and chromatographic elution in a Kinetex C18 column (4.6 mm × 50 mm × 2.6 μm) (Phenomenex, Torrance, CA, USA). All samples were randomized before injection and analyzed by the positive and negative mode in a hybrid mass spectrometer with QTOF 6550 mass analyzer (Agilent, Santa Clara, CA, USA). The mass spectra were acquired in centroid mode, and the mass range used for the acquisition was 50–1700 Da. The raw data were converted by the MassHunter Qualitative software (Agilent, Santa Clara, California, USA) and then imported to XCMS online software (Version 3.7.1, Scripps Center for Metabolomics, La Jolla, CA, USA). For the final statistical analysis, the Metaboanalyst 3.0 platform was used (McGill University, Montreal, Quebec, Canada), as well as the potential lipid biomarkers annotation by the measurement of their exact mass, retention time, and elution profile in METLIN (Scripps Center for Metabolomics, La Jolla, CA, USA), Human Metabolome Database (HMDB) (http://www.hmdb.ca/metabolites), and Lipid Maps databases (http://www.lipidmaps.org/).

### *2.7. Statistical Analyses*

The data were analyzed using GraphPad Prism 5.0 software. Results were confirmed to follow a normal distribution using the Kolmogorov–Smirnov test of normality with Dallal–Wilkinson–Lillie for corrected *P*-value. Data that passed the normality assumption were analyzed using analysis of variance (ANOVA) with a Bonferroni post hoc test. Data failing the normality assumption were analyzed using the non-parametric Kruskal–Wallis test followed by Dunn's post-test, and differences were considered statistically significant at a value of *p* < 0.05.

### **3. Results**

### *3.1. The Lack of Endogenous AnxA1 Exacerbates the IL-1*β *Release and Increases NLRP3 Levels after Inflammasome Activation*

First, we verified the endogenous effect of AnxA1 on the activation and regulation of NLRP3 inflammasome in macrophages. As expected, the administration of nigericin caused a significant reduction in cell viability (Figure 1A) without a difference between the two genotypes. Both nigericin and ATP induced IL-1β release by macrophages, which was more pronounced in nigericin-stimulated AnxA1-/- cells than in their respective controls (Figure 1B,C). This latter result was corroborated by the presence of mature IL-1β observed in AnxA1-/- cells (Figure 1C), and pro caspase 1 (Figure 1D) in the same experimental condition. ATP-stimulated WT macrophages presented decreased levels of AnxA1 compared with nigericin-stimulated cells (Figure 1D). In addition, pro caspase 1 levels were similar between only primed and ATP-stimulated AnxA1-/- cells (Figure 1D).

**Figure 1.** Lack of endogenous Annexin A1 (AnxA1) produced a marked release of IL-1β in macrophages. (**A**) 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. Macrophages of both genotypes showed a marked decrease in cell viability under nigericin and ATP exposure. The percentage of viable cells was calculated by optical density normalization for only lipopolysaccharide (LPS)-stimulated cells. Data are shown as mean ± S.E.M. of cell ratio (%). \*\* *p* < 0.01; \*\*\* *p* < 0.001 vs. LPS-stimulated cells of corresponding genotype (ANOVA, Bonferroni post-test). (**B**,**C**) IL-1β levels. Treatment with nigericin produced a marked release of IL-1β in AnxA1-null (AnxA1-/- ) macrophages compared with the other groups. Increased levels of pro-IL-1β were detected in the LPS-stimulated wild-type (WT) and AnxA1-/- cell extracts. Values are expressed as mean <sup>±</sup> SEM of IL-1<sup>β</sup> levels (pg/mL). \*\*\* *p* < 0.001, \*\* *p* < 0.01 vs. LPS-stimulated cells of respective genotype; ### *p* < 0.001 vs. WT nigericin-treated cells (ANOVA, Bonferroni post-test). (**D**) Pro caspase 1 and AnxA1 levels in the cell extracts under different experimental conditions. β-actin was used as an endogenous control (representative image of three experiments performed).

After detecting that nigericin-stimulated AnxA1-/- cells exhibited exacerbated IL-1β production, the AnxA1 and NLRP3 levels were analyzed using ultrastructural immunocytochemistry. The lack of endogenous AnxA1 was associated with increased levels of NLRP3 in nigericin-stimulated macrophages

compared with the primed AnxA1-/- cells and WT cells (Figure 2). In contrast, WT nigericin-stimulated macrophages showed a marked increase of AnxA1 levels compared with the only primed cells (LPS) (Figure 3A,B,E). In addition, points of co-localization between AnxA1 and NLRP3 were detected in the cytoplasm of nigericin-stimulated cells (Figure 3C). No immunogold labelling was detected in the negative control of the reaction (Figure 3D).

**Figure 2.** Lack of endogenous AnxA1 increased NLRP3 levels in nigericin-stimulated macrophages. (**A**) NLRP3 expression detected in the cytoplasm (arrows) of cells. Insets: details of cytoplasmic gold labelling of NLRP3. (**B**) Density of NLRP3 immunogold particles in macrophages. Data are mean ± SEM of distinct cells analyzed for each condition. \* *p* < 0.05 vs. LPS of corresponding genotype (ANOVA, Bonferroni post-test).

**Figure 3.** Nigericin-stimulated WT macrophages increase AnxA1 endogenous levels. (**A**,**B**) AnxA1 expression detected in the cytoplasm (arrows) and plasma membrane (white arrow) of cells. (**C**) Points of co-localization of AnxA1 (arrowheads) and NLRP3 (arrow) were detected in the cytoplasm of nigericin-stimulated cells. (**D**) Negative control. (**E**) Density of AnxA1 immunogold particles in macrophages. Data are mean ± SEM of distinct cells analyzed for each condition (*t*-test). \*\*\* *p* < 0.001 vs. LPS.

### *3.2. NLRP3 Activation Induces Di*ff*erent Lipid Release by WT and AnxA1-*/*- Macrophages*

To investigate whether the lack of endogenous AnxA1 also alters lipidomic profiling of macrophages under NLRP3 activation, lipidomic analysis of cell supernatants was performed. Figure 4 shows the score plots of principal component analysis (PCA) and partial least squares discriminant analysis (PLS-DA) obtained in positive and negative ion modes. As expected, the lipid profiles from the WT and AnxA1-/- control groups (LPS-stimulated cells) were different from each other, as shown by the separated dark blue and red ellipses in PCA and PLS-DA, either in the positive and negative ion modes. In the positive mode of the PCA analysis, the first two components of the score plot described 63.6% of explained variation in which the WT LPS separation from the WT treated with nigericin is more pronounced (Figure 4A) in PCA, whereas PLS-DA shows overlapped ellipses (Figure 4B). The negative mode in PCA and PLS-DA analysis showed similar results between groups, as evidenced by the overlapping of ellipses (Figure 4C,D).

**Figure 4.** Principal component analysis (PCA) and partial least squares discriminant analysis (PLS-DA) score plots of the lipid fraction. Each spot represents one supernatant sample from control (LPS-stimulated cells - WT: dark blue; AnxA1-/- : red) and nigericin-stimulated cells (WT: light blue; AnxA1-/- : green). The ellipses show the differences and similarities between groups. (**A**,**C**), PCA in positive and negative ion mode, respectively. (**B**,**D**), PLS-DA in positive and negative ion mode, respectively.

Heatmaps and dendrograms highlight the normalized concentrations of different lipids in the supernatants of WT and AnxA1-/- macrophages. In the positive mode (Figure 5A), the majority of potential lipid biomarkers were produced by nigericin-stimulated AnxA1-/- cells, if compared with the other experimental conditions. These cells showed increased production of sphingolipids, especially ceramides Cer(t18:0/18:O(2OH)), Cer(d14:1/26:0), PE-Cer(d14:2(4E,6E)/16:0), and PE-Cer(d14:2(4E,6E)/19:0) (Figure 5A). In addition, supernatants

from nigericin-stimulated AnxA1-/- exhibited a higher concentration of the eicosanoid 11S-15S-dihydroxy-14R-(S-glutathionyl)-5Z,8Z,12E-eicosatrienoic acid and the neutral lipid PI(P-18:0/00). In contrast, heptanoic acid was identified as a potential lipid biomarker in the supernatants of nigericin-stimulated WT cells, with a higher concentration than control WT cells (LPS), while arachidonoyl ethanolamide and the plasma membrane compound phosphatidylglycerol PG (16:1(9Z)/17:2(9Z,12Z)) were lower. PS (15:1(9Z)/16:1(9Z)) is a phosphatidylserine which only appeared in the supernatants of AnxA1-/- control cells and those treated with nigericin (Figure 5A).

**Figure 5.** Lipidomic analysis of WT and AnxA1-/- macrophage supernatants. Heatmaps and dendrograms show the hierarchical clustering of potential lipid biomarkers of the control (WT: dark blue; AnxA1-/- : red) and nigericin-stimulated cells (WT: light blue; AnxA1-/- : green). (**A**) Positive mode. (**B**) Negative mode. Lipidomic analysis demonstrated a completely different lipid profile between WT and AnxA1-/- supernatant cells. In WT cells, nigericin induced a pronounced release of eicosanoids and prostaglandins, while AnxA1-/- cells showed precursors of prostaglandin and some ceramides. The right bar in Figure B represents the blue–red code (−1.5 to 1.5) of the lipid concentrations. Unknown: noncharacterized lipids.

Figure 5B shows the potential lipid biomarkers in the negative mode. In the supernatants of nigericin-stimulated WT cells, the more concentrated lipids were associated with the arachidonic acid metabolism, such as the 10-hydroxyeicosatetraenoic acid (10-HETE), prostaglandins D1 and E1 (PGD1, PGE1), as well as palmitic acid, S-acetyldihydrolipoamide, N-palmitoyl serine, and PE(P-16:0/0:0). Curiously, there are different lipid profiles in the supernatants of WT and AnxA1-/- control cells, as characterized by a higher concentration of the eicosanoid 20-hydroxy-leukotriene E4 and fatty acid amides (13Z-docosenamide and 9,12Z-octadecadienamide) in the WT cells.

### **4. Discussion**

Monocytes and macrophages express NOD-like receptors, which are very important for the immune response during inflammation [20]. By NLRP3 inflammasome activation in these cells, pro-IL-1β and pro-IL-18 are cleaved and released, increasing the pro-inflammatory response [21]. Once macrophages are activated by the NLRP3 inflammasome agonists, such as nigericin and ATP, there is an IL-1β production and release, and the K<sup>+</sup> concentration is reduced [22]. IL-1β is a proinflammatory cytokine, and once it is released by macrophages, IL-6, TNF, nitric oxide, and prostaglandin E2 (PGE2) are produced [23]. Our data provide previously unknown details regarding the interplay between AnxA1 and NLRP3-derived IL-1β in macrophages and its relationship with lipid-mediator release.

Under normal conditions, the AnxA1 protein is present in high levels in the cytoplasm of human and rodent leukocytes, such as neutrophils, monocytes, and macrophages, and once these cells are activated, the AnxA1 moves to the cell membrane to be released and act as an autocrine or paracrine mediator [24,25]. The ATP-binding cassette transporter is responsible for AnxA1 secretion in macrophages [26], and under conditions of cellular stress, AnxA1 is rapidly released [27].

Our results show that the lack of AnxA1 exacerbates the IL-1β production after NLRP3 activation. These findings were supported by increased levels of NLRP3 in AnxA1-/- macrophages, as observed by ultrastructural immunocytochemistry. Considering that peritoneal AnxA1-/- macrophages presented increased levels of TLR4 [6], the lack of AnxA1 could favor LPS "over-priming" and consequent increase in NLRP3 inflammasome-derived IL-1β secretion. In addition, nigericin stimulation increased endogenous AnxA1 that presents points of co-localization with NLRP3 in WT macrophages, supporting a role of AnxA1 in the activation and regulation of the NLRP3 inflammasome. Regarding ATP stimulation, western blotting detected more decreased levels of AnxA1 in WT macrophages than in the priming LPS and nigericin-stimulated cells. In fact, activation of the P2 × 7 receptor by extracellular ATP in macrophages has been widely studied as a trigger of the NLRP3 inflammasome and is associated with the release of AnxA1 [7]. However, this study also demonstrated that nigericin did not induce AnxA1 release, suggesting pathways within P2 × 7R signaling in addition to K+ efflux in macrophages for the release of this protein.

Despite our findings, recent analyses show that bone-marrow-derived AnxA1-/- macrophages produced significantly lower secretion of IL-1β when activated with monosodium urate (MSU) crystals and ATP, but not NLRC4 or AIM2 activators (*Legionella pneumophila* or poly(dA:dT)) [28]. Notably, during the setting of MSU crystal-induced inflammation, the peak of the neutrophil influx was greater, and the resolution was slower in AnxA1-/- mice than in WT animals [29]. These findings are consistent with many studies that have shown an exacerbated inflammatory response in AnxA1-/ mice characterized by a marked leukocyte influx and production of proinflammatory mediators, such as IL-1β and IL-6 [4,6,30–32]. Additionally, the administration of the AnxA1 peptide (Ac2–26) 1 h before and 12 h after challenge with MSU crystals induced decreased levels of IL-1β in periarticular tissue, showing the important anti-inflammatory and proresolving activity of this protein on the course of MSU crystal-induced inflammation in mice [29]. Thus, the opposite effects described for the role of AnxA1 on NLRP3 inflammasome-derived IL-1β secretion could be a result of testing different macrophage populations, bone-marrow versus peritoneal-derived, and different strains, BALB/c versus C57BL/6, an important factor in the mouse immunology response [33,34].

The NLRP3 inflammasome is a cytosolic platform formed by a multi-protein complex containing a nucleotide-binding oligomerization domain-like receptor and the adaptor apoptosis-associated spec-like protein (ASC) containing an amino-terminal caspase-recruitment domain (CARD) [9]. The interaction of the NLRP3 with ASC promotes the recruitment of the procaspase-1 and its autoproteolysis, driving IL-1β and IL-18 maturation, membrane pore formation by the gasdermin D action, and then cytokine secretion and pyroptosis [35,36]. Besides the cleavage and release of the IL-1β, the activation of the NLRP3 inflammasome is directly related to the production of lipid mediators, including eicosanoids and ceramides. These lipids can be involved in metabolic and immunological

pathways, and the deregulation of NLRP3 activation causes an increase of the lipidic mediators released after pyroptosis, and it is possible to conduct metabolic damage on a systemic level [13].

The current lipidomics approach is a great tool to understand biological systems and many diseases. The major lipid classes can be categorized as fatty acyls, glycerolipids, glycerophospholipids, sphingolipids, sterol lipids, prenol lipids, saccharolipids, and polyketides [37]. In addition, cells can synthesize lipids, such as the eicosanoids, which are derived by the arachidonic acid oxidation and represented by prostaglandins, leukotrienes, thromboxanes, lipoxins, and epoxyeicosatrienoic acids [38]. By providing the exact mass, retention time, and elution profile, our study presents new data about the lipidomic profile of the supernatant of macrophages after NLRP3 activation by nigericin.

In our study, we showed the ceramides PE-Cer(d14:2(4E,6E)/16:0), Cer(t18:0/18:O(2OH)), PE-Cer(d14:2(4E,6E)/19:0), and Cer(d14:1/26:0) as potential lipid biomarkers released by AnxA1-/ macrophages after nigericin stimulation. Ceramides are bioactive sphingolipids present in the plasma membrane and they mediate cell signaling, with a close relationship to many pathophysiological processes associated with inflammation [39]. Previous studies showed that the exposure of macrophages to ceramides causes activation of caspase-1, and this effect is prevented by the absence of NLRP3 [40]. Ceramides have also been related to the activation of TLR4, augmenting LPS-induced pro-inflammatory response [41]. In this regard, it is reasonable to infer that increased levels of TLR4 in AnxA1-/ macrophages [6] contribute to the more pronounced ceramide concentration in their supernatants and also in NLRP3 activation.

Supernatants from nigericin-stimulated AnxA1-/- macrophages also exhibited a higher concentration of the 11S-15S-dihydroxy-14R-(S-glutathionyl)-5Z,8Z,12E-eicosatrienoic acid, a type of eicosanoid, and PI(P-18:0/0:0). Some studies have shown that 12-hydroxyeicosatrienoic acid (12-HETE) plays an important role as a paracrine mediator of inflammation, as well as in the regulation of neutrophil infiltration in damaged tissue [42,43]. PI(P-18:0/0:0) is an important phosphatidylinositol in cell membranes and for metabolic processes, such as being the primary source of arachidonic acid metabolism for eicosanoid synthesis and intracellular signals in animal tissues [44]. The lack of AnxA1 in macrophages increased the arachidonic acid metabolism and eicosanoid production, as evidenced by the high concentration of 11S-15S-dihydroxy-14R-(S-glutathionyl)-5Z,8Z,12E-eicosatrienoic acid and PI(P-18:0/0:0) after NLRP3 activation and pyroptosis induction. The 11S-15S-dihydroxy-14R-(S-glutathionyl)-5Z,8Z,12E-eicosatrienoic acid is present in leukocytes and red blood cells, and earlier studies have demonstrated that this fatty acid can be converted by arachidonic acid by the lipoxygenase pathway [45,46], which is basically synthesized during the inflammatory process [47].

In the supernatants of nigericin-stimulated WT macrophages, more concentrated lipids are also associated with the arachidonic acid metabolism, such as the 10-HETE, PGD1, PGE1, as well as palmitic acid, S-acetyldihydrolipoamide, N-palmitoyl serine, PE(P-16:0/0:0), and heptanoic acid, indicating a completely different lipid profile of AnxA1-/- supernatants; 10-HETE is a hydroxyeicosatrienoic acid with proinflammatory action, increasing TNF-α and IL-6 production in macrophages [48]. In contrast, PGE1 and PGD1 are anti-inflammatory lipid mediators that inhibit leukocyte migration and adhesion and mast cell activation [49–51]. Palmitic acid (PA), also called hexadecanoic acid, is one of the most common saturated fatty acids in animals. PA acts as a lipid mediator in inflammation and its derived-metabolic products accumulate in the endoplasmic reticulum (ER) and increase reactive oxygen species (ROS) generation, leading to cell death [52]. Additionally, the inflammatory response caused by ER stress and ROS generation from high concentrations of PA drives NF-κB and NLRP3 activation and, consequently, proinflammatory cytokine release by monocytes/macrophages [53–56]. S-Acetyldihydrolipoamide, N-palmitoyl serine and PE(P-16:0/00) (2-Hexadecanoyl-1-(1Z-hexadecenyl)-sn-glycero-3-phosphoethanolamine) are associated with cell metabolism [57], membrane receptor [58], and cell membrane compounds [59], respectively. However, there is little information about heptanoic acid in biological systems. Metabolomic studies have shown a high concentration of heptanoic acid in the faeces of autistic children [60], while in patients with

Crohn's disease, ulcerative colitis, and pouchitis, lower levels of this lipid were found [61]. Altogether, our data show that macrophages can release potential lipid biomarkers after NLRP3 activation that can regulate the inflammatory responses in the damaged tissue.

This study also detected a different lipid profile in supernatants from WT and AnxA1-/- LPS-stimulated macrophages. Higher concentrations of the eicosanoid 20-hydroxy-leukotriene E4 and fatty acid amides (13Z-docosenamide and 9,12Z-octadecadienamide) were found in the WT samples, while PS(15:1(9Z)/16:1(9Z)) was found in AnxA1-/- samples. In addition, 20-hydroxy-leukotriene E4 is an eicosanoid metabolite originating from the lipid oxidation of leukotriene E4 that plays an essential role in cell proliferation, differentiation, and immunoregulation [62]. Moreover, the leukotriene E4 contributes to prolonged intracellular signaling, increasing intracellular Ca2<sup>+</sup> and ERK phosphorylation [63], confirming signal pathway triggering by LPS on WT macrophages. Biological functions for both fatty acid amides detected in this study still need to be addressed.

Finally, PS(15:1(9Z)/16:1(9Z)) is a phosphatidylserine (PS) involved in cell signaling, including an important role in cell death, either by apoptosis, necroptosis, or pyroptosis, by its exposure on the outer plasma membrane layer [64]. The detection of this potential lipid biomarker in both AnxA1-/- supernatants (LPS and nigerin) can be related to the release of extracellular vesicles by macrophages. Macrophages can release extracellular vesicles after *Mycobacterium tuberculosis* infection, or spontaneously with a high concentration of phosphatidylserine [65].

### **5. Conclusions**

The lack of AnxA1 favors LPS "over-priming" and the release of lipid mediators (e.g., ceramides) that produce exacerbated NLRP3 activation under nigericin stimulation. Although more detailed investigations are warranted, this study identifies AnxA1 as a novel signaling component of inflammasome activation and a potential therapeutic target to treat inflammatory diseases.

**Author Contributions:** Conceptualization, J.M.S. and C.D.G.; Methodology, Validation, J.M.S., L.M.B., G.H.B.D., K.R.B., V.M. and C.D.G.; Formal analysis, J.M.S., G.H.B.D., V.M. and C.D.G.; Investigation, J.M.S. and C.D.G.; Resources, S.M.O., K.R.B., V.M. and C.D.G.; Writing—original draft preparation, J.M.S. and C.D.G.; Writing—review and editing, J.M.S., G.H.B.D., S.M.O., K.R.B., V.M. and C.D.G.; Funding acquisition, C.D.G. and S.M.O. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP), grant number 2016/02012-4 (S.M.O.) and 2017/26872-5 (C.D.G.). J.M.S. was supported by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior-CAPES (Finance Code 001) scholarship.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


© 2020 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 (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Annexin A1**/**Formyl Peptide Receptor Pathway Controls Uterine Receptivity to the Blastocyst**

### **Cristina B. Hebeda 1, Silvana Sandri 1, Cláudia M. Benis 1, Marina de Paula-Silva 1, Rodrigo A. Loiola 1, Chris Reutelingsperger 2, Mauro Perretti <sup>3</sup> and Sandra H. P. Farsky 1,\***


Received: 29 March 2020; Accepted: 28 April 2020; Published: 11 May 2020

**Abstract:** Embryo implantation into the uterine wall is a highly modulated, complex process. We previously demonstrated that Annexin A1 (AnxA1), which is a protein secreted by epithelial and inflammatory cells in the uterine microenvironment, controls embryo implantation in vivo. Here, we decipher the effects of recombinant AnxA1 in this phenomenon by using human trophoblast cell (BeWo) spheroids and uterine epithelial cells (Ishikawa; IK). AnxA1-treated IK cells demonstrated greater levels of spheroid adherence and upregulation of the tight junction molecules claudin-1 and *zona occludens-1*, as well as the glycoprotein mucin-1 (Muc-1). The latter effect of AnxA1 was not mediated through IL-6 secreted from IK cells, a known inducer of Muc-1 expression. Rather, these effects of AnxA1 involved activation of the formyl peptide receptors FPR1 and FPR2, as pharmacological blockade of FPR1 or FPR1/FPR2 abrogated such responses. The downstream actions of AnxA1 were mediated through the ERK1/2 phosphorylation pathway and F-actin polymerization in IK cells, as blockade of ERK1/2 phosphorylation reversed AnxA1-induced Muc-1 and claudin-1 expression. Moreover, FPR2 activation by AnxA1 induced vascular endothelial growth factor (VEGF) secretion by IK cells, and the supernatant of AnxA1-treated IK cells evoked angiogenesis in vitro. In conclusion, these data highlight the role of the AnxA1/FPR1/FPR2 pathway in uterine epithelial control of blastocyst implantation.

**Keywords:** mucin-1; claudin-1; *zona occludens*; ERK1/2 pathway; angiogenesis; F-actin polymerization; BeWo spheroids; Ishikawa cells

### **1. Introduction**

The endometrium is a critical tissue for the establishment and maintenance of pregnancy, during which it undergoes extensive physiological changes and demonstrates extraordinary plasticity. Cyclic changes in its tissues enable the endometrium to convert to a receptive state, allowing implantation, attachment, and invasion by the embryo through the epithelium into the underlying stromal compartment [1,2]. Embryo implantation is a highly-organized process that involves a receptive epithelium as well as a competent embryo for attachment, which is finely controlled by soluble and membrane-bound factors such as cytokines, prostaglandins, growth factors, and matrix-degrading enzymes as well as adhesion molecules, and transcription factors [2,3]. In order to achieve successful implantation, crosstalk between a receptive uterus and a competent blastocyst can only occur during a limited time span, known as the "window of implantation" [4,5]. During this short period of time,

which occurs approximately 6 to 10 days after ovulation in humans [1,6], the uterus undergoes structural and functional remodeling, mainly via the modulation of estradiol and progesterone. Basically, uterine receptivity is improved when estradiol levels decrease and high levels of progesterone are present [3]. Under these specific conditions, glycoproteins are expressed and highly secreted to prepare the endothelium for embryo attachment, tight junctions are reinforced, and angiogenesis occurs [2,7].

Annexin A1 (AnxA1) is a 37 KDa protein that belongs to the calcium and phospholipid-binding protein family within the annexin superfamily. A wide range of cells secretes AnxA1, including those of the innate immune system, as well as epithelial and cancer cells. AnxA1 mediates physiological processes in the body [8,9] although its secretion is highly augmented during challenging processes, such as inflammation and cancer [10,11]. The most well-recognized functions of AnxA1 are its potent anti-inflammatory and pro-resolution activities in the context of the innate immune response, during which glucocorticoids and cytokines induce synthesis and secretion of AnxA1 to halt inflammation and induce its resolution [12,13]. The binding of extracellular AnxA1 to G-protein coupled seven-domain transmembrane formylated peptide receptors (FPRs), especially type 2 (FPR2), is the most described and proven anti-inflammatory mechanism of AnxA1 [14–17]. Binding of AnxA1 to FPR2 induces rapid heterotrimeric G protein dissociation into the α and βγ subunits. βγ subunit downstream transduction signals, such as those mediated via phospholipase Cγ (PLCγ), result in activation of Ras family proteins and, in turn, activation of the mitogen-activated protein kinase (MAPK) pathway, particularly that of the extracellular signal-regulated kinases (ERK)-1/2. Activation of these later pathways leads to Ca2<sup>+</sup> mobilization and activation of protein kinase C (PKC) [18,19].

Previous data have correlated high levels of AnxA1 in human uterine tissue during gestation and in the seminal fluid [20,21], while lower amounts of AnxA1 have been found in the amnion and placenta at delivery [22–26]. More recently, our group has linked AnxA1 to pregnancy. Specifically, AnxA1-genetically deficient mice (AnxA1−/−) presented an increased number of blastocysts and implantation sites, resulting in an increased number of pups delivered [27]. Additionally, the uterine microenvironment of AnxA1−/<sup>−</sup> mice displayed an inflammatory profile, including a higher content of neutrophils and M1 macrophages as well as enhanced levels of pro-inflammatory cytokines, especially IL-6 [28]. These findings suggest that AnxA1 may play a crucial role in the maintenance of the uterine microenvironment, particularly in relation to maintenance of a receptive environment during implantation [27]. To further understand this process, in the current study we have elucidated the direct actions of AnxA1 on the initial events of blastocyst implantation in cultured human uterine epithelial cells.

### **2. Materials and Methods**

### *2.1. Cell Lines*

The human uterine epithelial cell line Ishikawa (IK) was purchased from Banco de Células do Rio de Janeiro. IK cells were maintained in Dulbecco's Modified Eagle Medium (DMEM; #12100046, Gibco, Carlsbad, CA, USA) supplemented with 10% heat-inactivated fetal bovine serum (FBS; #2024-06, Gibco), 2 mM L-glutamine (#25030081, Gibco) 1 mM pyruvate (#11360-070, Gibco) and 1% antibiotic solution containing streptomycin and penicillin (#15140-122, Gibco). Human umbilical vein endothelial cells (HUVECs) were donated by Dr. Ricardo José Giordano from the Chemical Institute of the University of Sao Paulo. HUVEC cells were maintained in Roswell Park Memorial Institute (RPMI) 1640 medium (#31800089, Gibco) containing 10% FBS and 1% antibiotic solution containing streptomycin and penicillin. BeWo cells were kindly donated by Professor Ana Campa from the Faculty of Pharmaceutic Sciences, University of Sao Paulo, and maintained in DMEM/F12 medium (#12500-062, Gibco) supplemented with 10% FBS and 1% antibiotic solution containing streptomycin and penicillin. All cells were maintained in an atmosphere of 5% CO2 at 37 ◦C and sub-cultured every 3 days by trypsinization, if necessary.

### *2.2. Cell Treatments*

Uterine epithelial cells were seeded in 24-well plates (Corning, New York, NY, USA) and cultured for adhesion over 18 h. Once cells had adhered, the medium was replaced and the cells were either pre-incubated with the culture medium (non-treated [NT], i.e., control) or medium supplemented with Boc-2 (1 μM; #SKU 0215276005, MP Biomedicals, Santa Ana, CA, USA), cyclosporine H (1 μM; #AG CN2 0447-M005, Adipo Gen Life Sciences, San Diego, CA, USA) or WRW4 (1 μM; #2262, Tocris Bioscience, Bristol, UK) for 30 min. Following the pre-incubation, AnxA1 (1.35 nM; donated by Professor Chris Reutelingsperger from Faculty of Health, Medicine and Life Sciences, Maastricht University) was added to the cell culture, either in the absence or presence of inhibitors, and incubated for a time period according to the specific assay performed.

For tube formation assay, uterine epithelial cells were washed three times with warm PBS and pre-incubated with FPR inhibitors (30 min) followed by addition of AnxA1, and cultured for 18 h in RPMI 1640 medium supplemented with 1% bovine serum albumin (BSA; #A9418-10G, Sigma-Aldrich). Afterwards, the supernatant was collected in sterile conditions and used to perform the assay.

### *2.3. Cell Viability Assay*

Uterine epithelial cells were seeded at 2.5 <sup>×</sup> 104 cells/well in 24-well plates and incubated in the absence or presence of different concentrations of AnxA1, Boc-2, cyclosporine H, or WRW4 over either 24 or 48 h. Following the incubation period, the medium was carefully removed and 300 μL of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT, 0.5 mg/mL; #M5655, Sigma-Aldrich) was added in each well. Cells were maintained at 37 ◦C for 3 h, after which the supernatant was removed and 200 μL of dimethyl sulfoxide (DMSO; #276855, Sigma-Aldrich) was added into each well and homogenized for 15 min. Absorbance was determined using a spectrophotometer at 575 nm (SpectraMax M Series, Molecular Devices, San Jose, CA, USA). Results were expressed as the percentage of viable cells relative to NT cells (control).

### *2.4. Flow Cytometry*

Flow cytometry experiments were performed to characterize the expression levels of AnxA1, FPR1, and FPR2 in uterine epithelial cells, as well as to investigate the role of AnxA1 on CD61, signal transducer and activator of transcription (STAT)1α, nuclear factor (NF)-κB, ERK1/2, homeobox A-10 (HOXA10), progesterone, and estrogen receptor expressions. Briefly, uterine epithelial cells were seeded at 5 <sup>×</sup> 104 cells/well and treated as mentioned above. Cells were trypsinized (#T1757, Vitrocell, Campinas, SP, BRA), washed twice in PBS containing 1% BSA (collectively referred to as PBS/BSA). To investigate the expression of AnxA1, HOXA10, progesterone, and estrogen receptors, the cells were fixed overnight at 4 ◦C using FACS lysing solution (#349202, BD Biosciences, San Jose, CA, USA), then washed with PBS containing 1% glycine (#01A1021.01.AG, Synth, Diadema, SP, BRA), permeabilized with Triton-X (0.001%; #T8787, Sigma-Aldrich), washed with PBS/BSA, and incubated with primary anti-human rabbit antibodies to AnxA1 (1:100; #713400, Thermo Fisher, Waltham, MA, USA), HOXA10 (1:100; #720220, Thermo Fisher), the progesterone receptor (1:500; #IM-0558, Rhea Biotech, Campinas, SP, BRA), or the estrogen receptor (1:250; #IM-0557, Rhea Biotech) overnight at 4 ◦C. Next, cells were washed with PBS/BSA and incubated with secondary goat anti-rabbit antibodies conjugated to Alexa Fluor 488 (1:500; #A11008 Invitrogen) for 40 min in the dark at room temperature (RT). To investigate STAT1α and ERK1/2 expressions, the cells were fixed in cold methanol for 30 min at RT, permeabilized in 0.1% Triton-X for 20 min at RT, and then incubated with primary anti-total STAT1α (1:50; #9172, Cell Signaling, Boston, MA, USA), anti-phospho-STAT1α (1:50; #9167, Cell Signaling), anti-total ERK1/2 (1:200; #ab54230, Abcam, Burlingame, CA, USA) and/or anti-phospho ERK1/2 (1:200; #ab214036, Abcam) antibodies overnight at 4 ◦C. After this incubation, the cells were blocked with PBS containing 2% FBS, and incubated with secondary anti-rabbit-phycoerythrin (1:200; #ab97070, Abcam) or anti-mouse-fluorescein isothiocyanate (FITC, 1:200; #ab6785, Abcam) goat antibodies for 1 h

in the dark at RT. In order to analyze FPR1, FPR2, CD61 and NF-kB expression, the cells were washed twice in PBS/BSA and incubated with specific antibodies as follows: FPR1-PE (1:100; #FAB3744P BD Biosciences, Minneapolis, MN, USA), FPR2-FITC (1:100; #bs3654R FITC; Bioss, Boston, MA, USA), CD61-FITC (1:50; #555753; BD Biosciences) or NF-kB (1:100; #0465R, Biolegend, San Diego, CA, USA) for 40 min in the dark at RT. The cells were then washed and resuspended in PBS at the end of each protocol. Samples were subjected to flow cytometric analysis in a BD Accuri C6 flow cytometer taking 10,000 events into consideration and using CSampler software (BD Pharmingen, CA, USA).

### *2.5. Proliferation Assay*

The proliferation assay was performed using a Live/Dead Viability/Cytotoxicity Kit for mammalian cells (#L3224, Thermo Fisher). Briefly, uterine epithelial cells were seeded in 24-well plates at a concentration of 1 <sup>×</sup> <sup>10</sup><sup>4</sup> cells/well and incubated in culture medium containing 0.5% BSA for 24 h prior to treatment. Following the incubation, the cells were washed and treated with medium containing either 0.5%, 2%, or 10% BSA in the absence or presence of AnxA1 (1.35 nM) for 24 or 48 h. Next, the cells were washed with PBS, trypsinized, and then incubated using the Live/Dead Viability/Cytotoxicity Kit according to the manufacturer's instructions. Samples were analyzed using a BD Accuri C6 flow cytometer and 10,000 events were considered in the analysis using CSampler software (BD Pharmingen).

### *2.6. Immunofluorescence*

Uterine epithelial cells were seeded at a concentration of 5 <sup>×</sup> 104 cells/well on glass coverslips inside the wells of 24-well plates and treated as mentioned above. Cells were fixed in cold methanol for 30 min at −20 ◦C, after which the methanol was removed, and the cells were maintained at −20 ◦C until the assay was performed. Briefly, cells were washed in PBS and incubated overnight at 4 ◦C in the presence of anti-Muc-1 (rabbit anti-human; #bs-4763R, Bioss), anti-claudin-1 (rabbit anti-human; #ab15098, Abcam), anti-*zona occludens*-1 (ZO-1; goat anti-human; #PA5-19090, Thermo Fischer), or anti-AnxA1 primary antibodies. Following this incubation, the cells were washed with PBS/BSA and incubated with donkey anti-goat and goat anti-rabbit secondary antibodies conjugated to either Alexa Fluor 568 (#A11011, Thermo Fisher) or FITC (#A11008, Thermo Fisher), respectively. The coverslips containing the cells were removed from the 24-well plates, inverted, and mounted on glass slides in 5 μL of Vectashield (#H-1200, Vectorlabs, Burlingame, CA, USA). The slides were maintained at 4 ◦C and images were acquired using an Axio Zeiss microscope and analyzed with ImageJ software (NIH, Bethesda, MD, USA).

### *2.7. ELISA*

Uterine epithelial cells were seeded at 3–5 <sup>×</sup> 104 cells/well and treated as mentioned above. The supernatant from these preparations was used to quantify the expression of IL-6 (#555220 BD OptEIA, BD Biosciences Pharmingen), AnxA1 (#MBS704042, MyBiosource, San Diego, CA, USA) and vascular endothelial growth factor (VEGF; #KHG0111, Thermo Fisher) through ELISA, according to the manufacturer's instructions. Expression level results from the ELISA were expressed in terms of pg/mL.

### *2.8. Trophoblast Spheroid*

The method of trophoblast spheroid growth was adapted from a previous study [29]. Briefly, in order to obtain spheroids, 50 μL agarose solution (1.5%; #A9539, Sigma-Aldrich) was added to 96-well plates. After solidifying, BeWo cells were seeded at a concentration of 1 x 104 cells/well and maintained in an atmosphere of 5% CO2 at 37 ◦C for 72 h. Following the incubation, individual spheroids were visualized through optic microscopy. The viability of spheroids was determined using the Live/Dead Viability/Cytotoxicity Kit (Thermo Fisher).

### *2.9. Implantation Assay*

The BeWo spheroid implantation model used here was adapted from previous studies [29–31]. Briefly, uterine epithelial cells were plated in 96-well plates at a concentration of 2.5 <sup>×</sup> 104 cells/well and, after adherence, the cells were incubated for 1 h in either the control medium or media containing Boc-2 (1 μM), cyclosporine H (1 μM), or WRW4 (1 μM), and then incubated with AnxA1 (1.35 nM) throughout the implantation time. The spheroids (one spheroid/well) were gently transferred onto adhered uterine epithelial cells and this co-culture was maintained in a humid atmosphere at 5% CO2 and 37 ◦C for 2 h. Following this incubation period, the wells were filled up to the brim with culture medium and the plates were sealed with an adhesive film for microplates, inverted, and then centrifuged at 30× *g* at RT for 5 min. After centrifugation, the plates were kept inverted while they were taken from the centrifuge and examined under a Leica DMi1 inverted microscope (Leica, Shinagawa, Tokyo, Japan) for the presence of the spheroids. The spheroids that disappeared during the centrifugation process were considered to be unattached, and the results were expressed as the percentage of attached spheroids.

### *2.10. Confocal Microscopy*

### 2.10.1. F-Actin Expression

Uterine epithelial cells were seeded at a concentration of 5 <sup>×</sup> 104 cells/well on a 24-well plate and then treated as mentioned above. Next, the cells were washed in PBS and fixed in 2% paraformaldehyde for 20 min at RT. The cells were then washed in PBS, incubated with rhodamine phalloidin (#R415, Thermo Fisher) for 20 min in the dark at RT, and then washed in PBS. The intensity of fluorescence was detected using high-content imaging with a GE IN Cell Analyzer 2200 (GE Healthcare Life Sciences, Chicago, IL, USA) and quantified with IN CartaTM image analysis software (GE Healthcare Life Sciences).

### 2.10.2. AnxA1 Expression

AnxA1 expression at the implantation site was evaluated in C57bl/6 mice of 5 to 6 weeks of age. For this purpose, female mice were caged overnight with male mice (3:1) and successful mating was verified the following morning. The presence of a vaginal plug was designated as day 0.5 of gestation. The animals were maintained and bred at the Animal House at the School of Pharmaceutical Sciences, University of Sao Paulo (Brazil). Chow (Quimtia, Colombo, PR, Brazil) and water were made available to the mice *ad libitum*. All animals were housed in a temperature-controlled room (22–25 ◦C and 70% humidity) with a 12-h light–dark cycle. All procedures were performed according to the Brazilian Society for the Science of Laboratory Animals (SBCAL) and approved by the Institutional Animal Care and Use Committee from the Faculty of Pharmaceutical Sciences of the University of Sao Paulo (Protocol number 557).

For confocal microscopy, the uterus was removed at day 5.5 of gestation following euthanasia of the mice via isoflurane overdose. Samples of the implantation sites were fixed in 4% buffered paraformaldehyde for 72 h at 4 ◦C, washed in Tris-buffered saline (TBS) and incubated with AnxA1 polyclonal antibody (#713400, Thermo Fisher) or only TBS (negative control) for 24 h. Next, these tissues were incubated with goat anti-rabbit antibodies conjugated with Alexa Fluor 488 (#A11008, Thermo Fisher) and DAPI (10 μg/mL; #D9542, Sigma-Aldrich) for 4 h, at RT in the dark, and then analyzed using a Confocal Zeiss LSM-780-NLO microscope (Carl Zeiss, Jena, Germany).

### *2.11. Tube Formation*

The tube formation assay was performed as detailed in previous studies [32,33]. Briefly, HUVEC cells were serum-starved for 24 h in RPMI 1640 medium with 1% BSA. Next, the cells were trypsinized, harvested, and plated on a 96-well plate at a density of 2.5 <sup>×</sup> 104 cells/well on 100 <sup>μ</sup>L Matrigel coating (#356237, Corning). The cells were incubated for 4 h with uterine epithelial cell-free supernatant that was previously obtained according to item 2.2. HUVEC cells, maintained in RPMI 1640 containing

10% FBS, were used as the positive control (data not shown). Photomicrographs (5X magnification) were obtained using a Leica DMI1 optical microscope (Shinagawa, Tokyo, Japan), and closed units (polygons) were considered in the count. Twelve fields were count per well.

### *2.12. Statistical Analyses*

The data were expressed as mean ± standard error of the mean (SEM) and comparisons were made between the experimental groups using a one-way ANOVA followed by either the Tukey test or Bonferroni test for multiple comparisons using GraphPad software version 5. A *p* value < 0.05 was used to denote statistically significant differences.

### **3. Results**

### *3.1. Uterine Epithelial Cells Express FPRs 1 and 2 and Secrete AnxA1*

To validate our study, we first confirmed that uterine epithelial cells express and secrete AnxA1, and express its receptors, FPR1 and FPR2 (Figure S1). The secretion of AnxA1 was not detected from other epithelial cell lineages, such as Caski and Siha cells, and low levels were detected for HeLa (Figure S1B). Additionally, the concentration-response curves demonstrated that AnxA1, Boc-2, cyclosporine H, and WRW4 did not affect the cellular viability under any of the concentrations employed in our studies following either 24 or 48 h of incubation (Figure S2A,C–E). Moreover, AnxA1 did not alter the cellular proliferation (Figure S2B). Using these data, effective concentrations of FPR agonists and antagonists were chosen to proceed with the further investigations, specifically 1 μM of Boc-2, cyclosporine H, and WRW4, and 1.35 nM of AnxA1.

### *3.2. AnxA1 Increased the Number of Implanted Trophoblast Spheroids*

BeWo spheroids were cultured on uterine epithelial cells in order to mimic embryo implantation in vitro (Figure S3A). Of note, BeWo spheroid viability was confirmed by observation of both a higher number of viable (green; Figure S3B,D) and lower number of dead cells (red; Figure S3C,D).

The in vitro implantation assay showed that NT (i.e., control) uterine epithelial cells demonstrated 36.4% spheroid adherence after 2 h of incubation. Similar adherence is observed when cells were treated with Boc-2, cyclosporine H or WRW4. AnxA1 treatment evoked a large increase in spheroid adherence, as 85.4% of the spheroids attached to the uterine epithelial cells following the treatment. This effect was reversed when cells were co-incubated with either cyclosporine H or Boc-2 with AnxA1. WRW4 did not affect the improved adherence evoked by AnxA1 (Figure 1A). A representative image of the in vitro spheroid adhesion assay is shown in Figure 1B.

### *3.3. AnxA1 Induced Muc-1 Expression in Uterine Epithelial Cells via FPR1 and FPR2*

Mucins are glycoproteins that line the surfaces of organs exposed to the external environment, including the lung, gut, eyes, and uterus [34]. It has been shown that, in humans, mucin-1 (Muc-1) acts as a scaffold and ligand for selectins present on the blastocyst in order to facilitate attachment [35,36]. The data obtained here show that, in uterine epithelial cells, AnxA1-induced expression of Muc-1 was abrogated by simultaneous incubation with Boc-2 or WRW4 (Figure 2A). Representative images of the immunofluorescence studies are shown in Figure 2B.

Since IL-6 is a key cytokine involved in blastocysts implantation and Muc-1 expression [37,38], we hypothesized that AnxA1 may control Muc-1 expression via IL-6. Indeed, we confirmed IL-6 increased Muc-1 expression by uterine epithelial cells (Figure 2C,D), although IL-6 secretion was impaired in cells treated with AnxA1 (Figure 2E). Taken together, these data show an IL-6-independent mechanism by which AnxA1 impacts on Muc-1 expression, and that such actions are mediated via FPRs.

**Figure 1.** AnxA1 increased BeWo spheroid attachment via FPR1 on uterine epithelial cells. Uterine epithelial cells were treated with FPRs antagonists during 1 h and AnxA1 was added with spheroids. Uterine epithelial and spheroids were co-cultured during 2 h, and the percentage of adhered spheroids were calculated and considered as attached. (−) means absence and (+) means presence of treatments (**A**). Representative image of non-treated (NT) and AnxA1-treated uterine epithelial cells containing or not a spheroid is shown in (**B**). The data are expressed as mean ± standard error of 10 experiments. <sup>a</sup> *p* < 0.05 vs. NT; <sup>b</sup> *p* < 0.05 vs. AnxA1.

**Figure 2.** AnxA1 controlled Muc-1 expression on uterine epithelial cells via FPR1/FPR2, independent of IL-6 secretion. Muc-1 expression on uterine epithelial cells was determined 24 h after incubations (**A**,**B**). Muc-1 expression was determined 24 h after IL-6 treatment (**C**). IL-6 secretion was determined in the supernatant of uterine epithelial cells 24 h after treatments (**D**). (−) means absence and (+) means presence of treatments. The data are expressed as mean ± standard error of mean of three to five independent experiments. <sup>a</sup> *p* < 0.05 vs. NT; b,c *p* < 0.05 vs. AnxA1.

### *3.4. AnxA1 Induced Claudin-1 and Zona Occludens-1 Expression in Uterine Epithelial Cells via FPR1 and FPR2*

Claudin-1 is a member of the junctional complex and is associated with cytoplasmic plaque proteins in the *zona occludens* (ZO), which is crucial for maintaining the integrity of the uterine epithelium [7]. AnxA1 treatment increased claudin-1 and ZO-1 expression in uterine epithelial cells, and this effect was abrogated by co-incubation with Boc-2 or WRW4 (Figure 3A,B, respectively). Representative images of claudin-1 and ZO-1 immunofluorescence are shown in Figure 3.

**Figure 3.** AnxA1 controlled claudin-1 and ZO-1 expressions on uterine epithelial cells via FPR1 and FPR2. Claudin-1 (**A**) and ZO-1 (**B**) expressions on uterine epithelial cells were determined 24 h after incubations. Representative images of claudin-1 and ZO-1 immunofluorescence are shown. (−) means absence and (+) means presence of treatments. The data are expressed as mean ± standard error of mean of three to five independent experiments. <sup>a</sup> *p* < 0.05 vs. NT; <sup>b</sup> *p* < 0.01 and <sup>c</sup> *p* < 0.05 vs. AnxA1.

### *3.5. Increased Muc-1 and Claudin-1 Expression Evoked by AnxA1 Was Supported by the MAPK Pathway Activation in Uterine Epithelial Cells*

ERK1/2, STAT1α, and NF-κB are some of the signaling molecules connected to Muc-1, claudin-1, and ZO-1 expression [31,39–41]. Furthermore, AnxA1 produces its actions mainly through the MAPK, JAK/STAT, and NF-κB signaling transduction pathways [9,42,43]. We observed that AnxA1 treatment increased ERK1/2 phosphorylation in uterine epithelial cells (Figure 4A) but did not modify STAT1α phosphorylation (Figure 4B) or the p65 subunit of NF-κB (Figure 4C) compared to the control cells (NT). Moreover, our findings showed that the pharmacological blockade of ERK1/2 phosphorylation, by pre-incubation with PD98059, abrogated the increment of Muc-1 and claudin-1 expressions induced by AnxA1 in uterine epithelial cells (Figure 4D,F, respectively). In contrast, pre-incubation of uterine epithelial cells with PD98059 did not block the ZO-1 expression evoked by AnxA1 (Figure 4H). Representative images of Muc-1, claudin-1 and ZO-1 immunofluorescence are shown in Figure 4E,G,I, respectively.

**Figure 4.** AnxA1 controlled Muc-1 and Claudin-1 expressions on uterine epithelial cells via ERK1/2 phosphorylation. The effect of AnxA1 on ERK (**A**) and STAT1α (**B**) phosphorylation and NF-κB expression (**C**) were investigated by flow cytometry. The inhibition on ERK1/2, evoked by PD98059 incubation, was investigated on Muc-1 (**D**), Claudin-1 (**F**) and ZO-1 (**H**) expressions using immunofluorescence. Representative images of Muc-1, Claudin-1 and ZO-1 are shown in (**E**), (**G**) and (**I**), respectively. (−) means absence and (+) means presence of treatments. The data are expressed as mean <sup>±</sup> standard error of mean of three to five independent experiments. <sup>a</sup> *p* < 0.05 and <sup>b</sup> *<sup>p</sup>* <sup>&</sup>lt; 0.01 vs. NT; <sup>c</sup> *p* < 0.05 and <sup>d</sup> *p* < 0.01 vs. AnxA1.

### *3.6. AnxA1 Increased F-Actin Polymerization in Uterine Epithelial Cells via FPR1*

F-actin is connected to the ZO-1 and is important in stabilizing the tight junctions [44,45]. AnxA1 treatment increased F-actin polymerization in comparison to the control treatments (NT; dotted line), and this effect was inhibited by co-treatment with Boc-2 or cyclosporine H. Co-incubation of cells with WRW4 and AnxA1 did not modify F-actin polymerization (Figure 5A). Representative images are shown in Figure 5B.

**Figure 5.** AnxA1 induced F-actin polymerization via FPR1. F-actin was quantified on uterine epithelial cells 2 h after incubations, using phalloidin-rhodamine by confocal microscopy. (−) means absence and (+) means presence of treatments (**A**). Representative images of F-actin polymerization are shown (**B**). The data are expressed as mean ± standard error of mean of three to five independent experiments. <sup>a</sup> *p* < 0.05 vs. NT; <sup>b</sup> *p* < 0.05 vs. AnxA1.

### *3.7. AnxA1 Controls Endothelial Tube Formation and VEGF Secretion via FPR2*

Angiogenesis is a fundamental step in the implantation process, and thus, required for pregnancy to progress [46]. Therefore, we investigated the role of substances secreted by uterine epithelial cells after treatment with AnxA1 and/or FPR inhibitors on tube formation of HUVECs, referred to as in vitro angiogenesis. Cyclosporine H or WRW4 did not alter the number of HUVEC tubes *per se*. The supernatant of uterine epithelial cells, which had previously been incubated with AnxA1 or AnxA1 plus the FPR1 antagonist cyclosporine H, did not modify the formation of tubes by HUVECs. In contrast, the supernatant of uterine epithelial cells treated with AnxA1 and WRW4 did markedly reduce the number of tubes formed (Figure 6A). Representative images demonstrating this are depicted in Figure 6B.

**Figure 6.** Supernatant of uterine epithelial cells treated with AnxA1 plus WRW4 presented reduced levels of VEGF and reduced HUVEC tube formation. Supernatant was obtained from uterine epithelial cells previously treated with cyclosporine H or WRW4 in the absence or presence of AnxA1. HUVEC was incubated with supernatants and tube formation was evaluated 4 h later (**A**). Representative images of tube formation are shown in (**B**). The levels of VEGF in the uterine cells supernatant were quantified 4 h after incubations (**C**). (−) means absence and (+) means presence of treatments. The data are expressed as mean ± standard error of mean of at least three to five independent experiments. <sup>a</sup> *p* < 0.05 vs. AnxA1.

VEGF is a fundamental growth factor in angiogenesis. Therefore, to understand the mechanisms linked to the altered tube formation observed, we quantified the levels of VEGF in the supernatant previously obtained from uterine epithelial cells incubated with AnxA1 and FPR blockers. In accordance with the results obtained from the HUVEC tube formation experiments, VEGF levels were not altered by cyclosporine H, WRW4, AnxA1, or co-incubation with cyclosporine H and AnxA1. In contrast, significantly lower levels of VEGF were quantified in the supernatants from uterine epithelial cells treated with both WRW4 and AnxA1 (Figure 6C).

### *3.8. AnxA1 Is Physiologically Expressed on Uterine Epithelial Cells and the Blastocyst during In vivo Implantation*

To corroborate our results on the functional involvement of AnxA1 in uterine receptivity and embryo implantation, the expression of AnxA1 was monitored by confocal microscopy of implantation sites obtained from C57bl/6 mice on gestational day 5.5 (Figure 7A). The data obtained showed that AnxA1 was broadly expressed by both the luminal epithelium and the embryo (Figure 7B). A representative image of the negative control is shown in Figure S5**.**

**Figure 7.** AnxA1 is expressed at embryo-implantation site in vivo. Implantations sites were obtained from C57bl/6 at gestational day 5.5 (**A**). Representative image of AnxA1 expression at implantation site (arrow) and luminal tissue by confocal microscopy is shown (**B**).

### **4. Discussion**

Failure of the blastocyst to implant in the uterine wall is a putative cause of unsuccessful pregnancy. Although large volumes of data have been published regarding this process, research thus far has failed to produce an effective treatment that supports the uterus' receptivity to implantation. Blastocyst apposition and attachment to the uterus involve the actions of a diverse range of molecules and intracellular signaling processes that are not unique in themselves, but play unique roles in each step of the process [2]. Moreover, ethical concerns limit the ability for human studies, and thus, our current understanding of human embryo–endometrium interactions is limited, although experimental models and in vitro cell systems have been designed and are currently being used to help understand the complexity of this phenomenon [29,30]. About 30% of human pregnancies end in miscarriages, the vast majority of which occur in the early phase of gestation [47], thus emphasizing the crucial importance of further studies in this field. Here, we highlight the role that the AnxA1 protein plays, via activation of G-protein coupled receptors (specifically FPRs), in preparing the uterine epithelium for blastocyst implantation using an in vitro model. We found that, indeed, AnxA1 application favors blastocyst attachment by inducing the expression of proteins by the uterine epithelium that control the paracellular flux, structure, and adhesiveness of the uterine wall, as well as promoting angiogenesis.

Progesterone binds to nuclear receptors in the cells of the uterine epithelium and stroma, and activates the transcription factor HOXA10 to induce cell proliferation in the uterine wall, a process which is required for implantation of the blastocyst [48,49]. It has been shown the HOXA10 gene promoter contains a progesterone-responsive element, implying that HOXA10 is a direct target of the progesterone receptor [50,51]. Moreover, activation of the transcription factor HOXA10 induces the expression of the adhesion molecule CD61, a pivotal beta-3 integrin for blastocyst attachment in the epithelium [52]. In humans, high plasma progesterone levels have been associated with increased levels of CD61 in uterine biopsy samples, suggesting a role for CD61 as a biomarker of uterine receptivity [53]. A role for AnxA1 in the fundamental progesterone/HOXA10/CD61 pathway of blastocyst implantation was initially disregarded after studies demonstrated that AnxA1 treatment failed to induce intracellular

expression of the progesterone receptor or HOXA10 (Figure S4), epithelial proliferation (Figure S2) or expression of CD61 (Figure S4).

Conversely, our data show that AnxA1, via activation of FPRs, induces expression of Muc-1. Muc-1 is a glycoprotein that is highly expressed in the receptive human endometrium and, subsequently, is removed from the apical epithelium at the location of implantation, while continuing to be expressed by cells neighboring the implantation site [2,54–56]. Moreover, abnormal Muc-1 expression has been detected in women suffering from recurrent miscarriages or fertility problems [57]. Muc-1 expression is controlled by various molecules, such as IL-6 [37,38] and high levels of progesterone, either alone or in combination with estradiol in vivo and in vitro [58,59]. Here, we assumed that other pathways are involved in AnxA1-induced Muc-1 expression, as AnxA1 reduced secretion of IL-6 by uterine epithelial cells while no alterations of progesterone and estrogen receptors were observed in this cell line. Data regarding the action(s) of AnxA1 on mucin expression are not currently available, and the results presented here may help further investigations concerning the effects of AnxA1 on Muc-1 expression in other epithelial cells, since Muc-1 is known to be present on the surface of most epithelial cells and to be involved in processes such as microbe invasion, inflammation, and fibrosis [60,61].

Similar to the results observed for Muc-1 expression, we detected increase in claudin-1 and ZO-1 expression following treatment with AnxA1, and these effects were blocked by co-treatment with FPR1 and FPR2 antagonists. Claudin-1 is a transmembrane adhesion protein that binds to cytoplasmatic adapter proteins of the ZO-1 and forms tight junctions [62–64]. Crucial roles for both proteins have been described in regulating transepithelial permeability to small molecules and ions [63,65], as well as cell growth and differentiation under various conditions [42,66–69]. Recently, claudin-1 and ZO-1 have been revealed to be key components involved in uterine receptivity, as tight junctions become reinforced during blastocyst implantation [7,70]. Here, we show that the AnxA1/FPR1/2 pathway contributes to strengthening claudin-1 and ZO-1 between uterine epithelial cells, suggesting a role by AnxA1 in controlling paracellular flow. Accordingly, the reinforcement of tight junctions contributes to the retention of uterine secretions within the lumen of the uterus, which is important for nourishing the embryo and favoring implantation [7,70,71].

The downstream effects of AnxA1 on intracellular pathways in uterine epithelial cells involve MAPK signaling, but not engagement of the transcription factors STAT1α or NF-κB. Increased ERK1/2 phosphorylation was detected following AnxA1 treatment, while inhibition of ERK1/2 phosphorylation reverted the changes in expression of Muc-1 and claudin-1 induced by this AnxA1 treatment. In fact, it has been shown that both Muc-1 and claudin-1 are pre-transcriptionally induced via the MAPK/ERK pathway [72–74]. Furthermore, MAPK signaling is a common pathway implicated in various actions induced by AnxA1 [9,13,75,76]. However, AnxA1 controls ZO-1 expression independently of ERK1/2 phosphorylation, potentially via improving F-actin polymerization as interactions between ZO-1 and the actin cytoskeleton have been reported in epithelial cells [44,77]. The pivotal role of AnxA1 on cytoskeletal reorganization of epithelial cells has been demonstrated under physiological conditions, and a deficiency of AnxA1 expression with consequent destabilization of focal adhesions and tight junctions has been implicated as a possible mechanism of various diseases [78–83].

Angiogenesis is crucial for successful implantation and placentation, and is critical during decidualization [46]. Angiogenesis begins early in the course of implantation and is supported by pro-angiogenic molecules [84]. The role of AnxA1 in angiogenesis has been shown in different models of cancer [85,86]. Moreover, the N-terminal AnxA1 peptide Ac2-26 has been demonstrated to increase endothelial tube formation by increasing proliferation, migration, and actin polymerization in a manner similar to that induced by VEGF-A [83]. Our data show AnxA1, via FPR2, induces the secretion of angiogenesis factors by uterine epithelial cells, resulting in HUVEC tube formation. In our model, VEGF was identified as one of these angiogenic factors as its concentrations were reduced if uterine epithelial cells were treated with FPR2 antagonist in association with AnxA1. This is in line with recent work by Ferraro and co-workers [87], who demonstrated a functional link between AnxA1 and reparative macrophage phenotype in settings of heart failure, consequent to VEGF release from the

immune cell. Therefore, we conclude that AnxA1, via FPR2, induced epithelial secretion of VEGF which, subsequently, modulated angiogenesis in the endothelium, demonstrating a complementary mechanism by which AnxA1 is able to support embryo implantation.

The direct action of AnxA1 on uterus epithelium demonstrated here seems to sound controversial to the high number of pups delivered by AnxA1 knockout mice [27]. In addition to the differences found in epithelium of mice and humans, we infer the exacerbated inflammation detected in the uterus microenvironment in AnxA1-deficient mice may favor the implantation process. It is known that elevated levels of pro-inflammatory cytokines, such as IL-6, favor the implantation of blastocysts [37]. Therefore, we infer AnxA1 is pivotal in the implantation phase, by controlling the inflammation that maintains the microenvironment to support a compatible blastocyst implantation, and also, by inducing the required signaling in the epithelium to trigger the adhesive properties.

In conclusion, this study unveils the existence of an intricate mechanism by which AnxA1 controls embryo implantation through regulating expressions and functions of key molecules linked to uterine receptivity, integrity, and angiogenesis. AnxA1 interacts with FPRs to activate members of MAP kinases and modulate the epithelial cytoskeleton, resulting in a uterine environment conducive for embryo implantation in the epithelium. Moreover, AnxA1 is also connected to the dynamic interplay between the uterine epithelium and endothelium, crucial for embryo implantation, posterior decidualization, and consequently, successful pregnancy.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2073-4409/9/5/1188/s1, Figure S1: AnxA1 expression and secretion and FPR1 and FPR2 expressions by uterine epithelial cell lineage Ishikawa. Figure S2: AnxA1, Boc-2, cyclosporine H and WRW4 did not modify uterine epithelial cells viability. Figure S3: BeWo spheroid viability. Figure S4: AnxA1 did not modify progesterone and estrogen receptors, HOXA10, and CD61 expressions. Figure S5: Representative image of negative control of embryo implantation site.

**Author Contributions:** C.B.H., S.S., R.A.L., M.d.P.-S., C.R., M.P., and S.H.P.F. were involved in the conceptualization of the study; C.B.H., S.S., C.M.B., R.A.L., and S.H.P.F. collected, analyzed and interpreted the data; C.B.H., S.S., and S.H.P.F. wrote the manuscript; S.H.P.F. was the project administrator; S.H.P.F. acquired the funding acquisition. All authors critically reviewed the paper and approved its final version. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by FAPESP (Fundação de Amparo à Pesquisa do Estado de São Paulo), grant number 2014/07328-4; C.B.H. and S.S. are post doctoral fellows of CAPES (Coordenação de Aperfeiçoamento de Pessoal de Nível Superior). S.H.P.F. is a fellow researcher of CNPq (Conselho Nacional de Pesquisa). M.P. is supported by the William Harvey Research Foundation.

**Acknowledgments:** The authors acknowledge Mario Costa Cruz and Iuri Cordeiro Valadão from the Core Facility for Support Research (CEFAP, USP) for contributions on confocal protocol discussions.

**Conflicts of Interest:** The authors declare no conflict of interest.

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