All BC cells and normal mammary cells (MCF-10A) were purchased from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China) and cultured according to the supplier’s instructions. Fresh BC tissues, adjacent noncancerous tissues, and corresponding paraffin-embedded cancer tissues (n = 120) were obtained from Harbin Medical University Cancer Hospital between 2012 and 2013. All the patients enrolled in our study had complete clinicopathological information, and patients who received neoadjuvant chemotherapy, radiotherapy, and immunotherapy and those with recurrent tumors, bilateral BC, or other previous tumors were excluded. Our study was approved by the Research Ethics Committee of Harbin Medical University, and all the patients enrolled in our study signed informed consent forms.

Total RNA was extracted from fresh BC tissues and cells with TRIzol reagent (Invitrogen, United States) according to the manufacturer’s instructions. The Prime Script RT Reagent Kit with gDNA Eraser (Takara, Kyoto, Japan) was used to synthesize cDNA, and qRT-PCR was conducted using SYBR Premix Ex Taq™ II (Takara) on a CFX96 Touch Detection System (Bio-Rad, United States). The primers used in our study are listed in Supplementary Table S1. GAPDH was used as an internal control for detecting mRNA expression; LncRNA (HCG18) expression was normalized by 18S rRNA and miRNA expression were normalized by U6. The relative RNA abundance was calculated by the standard 2^−∆∆CT method.

Western blotting assays were performed as previously reported (Liu et al., 2020b). Briefly, the indicated cells were lysed with RIPA (Beyotime, China) buffer containing phenylmethylsulfonyl fluoride (Beyotime). Then, the lysate was collected, and the proteins were separated by 10% sodium dodecyl sulfate–polyacrylamide gel electrophoresis and transferred to polyvinylidene fluoride membranes. After that, the membranes were incubated with primary antibodies and secondary antibodies. Finally, the protein bands were detected using a ChemiDoc Touch imaging system (Bio-Rad). The antibodies used are listed in Supplementary Table S2.

For the Cell Counting Kit-8 (CCK-8) assays, the indicated cells were seeded into 96-well plates at a density of 2 × 10³ cells per well. At each preset time, 10 μL CCK-8 reagent (Beyotime) was added to each well containing 90 μL culture medium, and the plates were incubated at 37°C for 2 h. After that, the cell viability was detected by measuring the absorbance at 450 nm.

For the colony formation assays, the indicated cells were seeded into six-well plates (5 × 10² cells/well) and incubated with culture medium containing 10% fetal bovine serum (FBS) at 37°C for 2 weeks. After that, the cells were fixed with formalin for 30 min and stained with crystal violet. A FluorChem M system was used to take the photographs.

5-Ethynyl-2′-deoxyuridine (EdU) staining assays were performed as follows: the indicated cells were seeded into 96-well plates and cultured with 10% FBS medium at 37°C. After the confluence reached 60–70%, the cells were subjected to EdU staining (Beyotime) according to the manufacturer’s instructions. After fixation and permeabilization, the cells were stained with EdU and DAPI solutions. The samples were photographed and analyzed under a fluorescence microscope.

For wound scratch assays, the indicated cells were seeded into six-well plates and cultured at 37°C. After the cells reached 95% confluence, sterile micropipette tips were used to make scratch wounds. Then, the cells were washed with PBS and cultured with serum-free medium continuously. Images were taken under a microscope at preset times, and the migration rate was subsequently analyzed. The percentage of wound closure (%) = (width on 0 h—with on 24 or 48 h)/width on 0 h × 100%.

For cell invasion assays, the upper chambers of 24-well Transwell plates (8.0 µm) (Corning, United States) were coated with Matrigel and incubated at 37°C for 4 h. Then, 200 μL serum-free medium containing the indicated cells (1 × 10⁵) was added to the upper chambers of the Transwell plates, and 600 μl 10% FBS medium was added to the lower chambers of the Transwell plates. Then, the cells were cultured at 37°C for 24 h. Afterward, the noninvaded cells were scraped from the upper side of the chambers, and invaded cells on the lower side were fixed with 4% paraformaldehyde and stained with 1% crystal violet. Finally, the images were captured, and the invaded cells were counted under a microscope.

Hematoxylin–eosin (HE) staining assays were performed according to a standard HE staining technique as previously reported (Chong and Murphy, 2004). Briefly, mouse lung tissues were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned (5-mm thickness). After that, the lung sections were subjected to HE staining to evaluate pathological changes and photographed under an optical microscope.

Immunohistochemistry (IHC) staining was performed on paraffin-embedded specimens from BC patients following a standard streptavidin–peroxidase complex procedure. The staining results were evaluated and calculated as histochemistry scores (H scores) by three senior pathologists independently. The following antibodies were used: anti-UBE2O (catalog no. 15812-1-AP; Proteintech Group, China) and anti–Ki-67 (catalog no. 9449; CST, United States).

For sphere formation assays, the indicated cells (1 × 10³) were trypsinized and resuspended in stem cell medium consisting of DMEM/F-12 (Gibco, United States), 1 × B27 (Invitrogen), 20 ng/ml epidermal growth factor (Invitrogen), 20 ng/mL basic fibroblast growth factor (Invitrogen), and 2 mM L-glutamine (Invitrogen) and seeded into 24-well ultralow attachment plates. Then, the cells were cultured at 37°C, and the medium was refreshed every 48 h. Two weeks later, the stemness spheres were observed and counted under an optical microscope.

Lentiviral production and cell transfection were performed as we previously described (Liu et al., 2020b). Briefly, recombinant lentivirus for overexpression of HCG18 or UBE2O in BC cells and corresponding negative controls were designed and synthesized by GenePharma (Shanghai, China). The recombinant lentivirus was transfected into the BC cells with polybrene (8 μg/ml). Twenty-four hours after transfection, the cells were selected with puromycin, and qRT-PCR was applied to detect the transfection efficiency.

To establish stable HCG18- or UBE2O-reducing BC cells, shRNAs targeting HCG18 or UBE2O were designed by Sigma and transfected into the indicated cells according to the manufacturer’s instructions. The shRNA sequences are listed in Supplementary Table S3. qRT-PCR was used to detect the transfection efficiency.

Small interfering RNAs (siRNAs) targeting HIF-1α and control siRNAs were designed by GenePharma. The miRNA mimics and inhibitor were synthesized by Sigma. For transfection, the indicated cells were seeded into six-well plates (5 × 10⁵ per well) and cultured at 37°C overnight. Then, the cells were transfected with the corresponding vectors using Lipofectamine 2,000 transfection reagent (Invitrogen) according to the manufacturer’s instructions. Forty-eight hours later, the transfected cells were harvested for follow-up experiments. The siRNA sequences are listed in Supplementary Table S3.

Nuclear and cytoplasmic RNA separation was conducted using a PARIS™ Kit (Invitrogen) according to the manufacturer’s instructions. qRT-PCR was used to analyze the expression of HCG18 in different fragments of BC cells.

RNA immunoprecipitation (RIP) assays were performed to validate the interaction of miR-103a-3p with HCG18, UBE2O, or AGO2 using the Magna RIP™ RNA-Binding Protein Immunoprecipitation Kit (Millipore, United States). To validate the interaction between miR-103a-3p and HCG18 or UBE2O, HEK293T cells were cotransfected with MS2bs vectors cloned with related DNA sequences (MS2bs, MS2bs-HCG18-WT, MS2bs-HCG18-Mut, MS2bs-UBE2O-3′UTR-WT, or MS2bs-UBE2O-3′UTR-Mut) and MS2bp-GFP overexpression vector (GenePharma, China). To validate the interaction between miR-103a-3p and AGO2, the indicated cells (MDA-MB-231 ^NC/shHCG18 and MCF-7 ^NC/HCG18 ) were transfected with miR-103a-3p mimics. Then, the cells were collected and lysed with RIPA. After that, the lysate was subjected to RIP with an anti-GFP or anti-AGO2 antibody (Abcam, United States) according to the manufacturer’s instructions. Finally, the samples were treated with proteinase K (Merck, United States), and qRT-PCR was used to analyze the binding of target RNA and protein.

To assess the regulatory effects of HCG18 on miR-103a-3p and miR-103a-3p on UBE2O mRNA, dual-luciferase reporter assays were performed using a double-luciferase assay system (Promega, United States). In brief, 3′ untranslated region (UTR) of UBE2O (wild/mutant type) or HCG18 (wild/mutant type) luciferase reporter plasmids and miR-103a-3p mimics were cotransfected into HEK-293T cells. Then, the cells were lysed, and luciferase assays were performed according to the manufacturer’s instructions. Firefly luciferase activity normalized to Renilla luciferase activity was used as an internal control.

To analyze whether HIF-1α could bind to the promoter region of HCG18, HEK-293T cells were cotransfected with luciferase reporter plasmids encoding the HCG18 promoter region (wild/mutant type) and with HIF-1α plasmids. Forty-eight hours later, the cells were harvested, and dual-luciferase assays were performed. Firefly luciferase activity was normalized to Renilla luciferase activity (used as an internal control).

Chromatin immunoprecipitation (ChIP) assays were performed as previously reported (Liang et al., 2020). Briefly, the indicated cells were treated with formaldehyde to crosslink the target protein and chromatin and sonicated to an average length of 200 to 500 bp on ice. Then, a DNA depuration kit (Beyotime) was used to extract and clean the DNA fragments. After that, ChIP assays were conducted with an anti–HIF-1α antibody (catalog no. 36169S; CST) or immunoglobulin G (IgG) control. qRT-PCR was used to detect the promoter fragments of HCG18. The primers used in our experiment are listed in the Supplementary Material.

All animal studies were approved by the Medical Experimental Animal Care Commission of Harbin Medical University and performed in the Second Affiliated Hospital of Harbin Medical University Laboratory. Animals used in our study were raised in specific pathogen-free animal facilities (temperature was maintained at 25°C with a 12-h light–dark cycle) and provided free access to clean water and food. The animals were anaesthetized with 1–3% isoflurane and humanely sacrificed by CO₂ inhalation as previously reported (Liu et al., 2020c). For the tumorigenesis assay, 6-week-old BALB/c nude mice (Vital River, Beijing, China) were randomly assigned into two groups (n = 6), and then MDA-MB-231 ^NC or MDA-MB-231 ^HCG18-sh (5 × 10⁵) was injected into the mammary fat pads of mice. Tumor growth was measured every week and 7 weeks after injection. The mice were humanely sacrificed, and the tumors were extracted and fixed in formalin solution. The tumor volumes were evaluated by the following formula: 1/2 [length × width (Wang et al., 2019)]. For the lung metastasis model, MDA-MB-231 ^NC or MDA-MB-231 ^HCG18-sh (5 × 10⁵) cells were injected into the tail vein of 6-week-old BALB/c nude mice. Seven weeks after injection, the mice were humanely sacrificed, and the lungs were collected to count metastatic nodules and for HE staining.

All the data are presented as the means ± SD from at least three independent experiments. A normality test was used to analyze the distribution of all datasets. Differences between two groups were analyzed by Student t tests, and differences among multiple groups were analyzed by one-way analysis of variance. The χ ² test was used to analyze the relationship between HCG18 expression and the clinicopathological features of BC patients. The Kaplan–Meier method and log-rank test were applied to generate and assess survival curves. p < 0.05 indicated statistical significance, which was evaluated by GraphPad Prism 8.0 software.

A previous study reported that HCG18 was upregulated in BC tissues compared with normal tissues in the TCGA database (Xu et al., 2017). However, the biological function of HCG18 in BC is largely unknown. To investigate this, we first detected the HCG18 expression profile in BC tissues and corresponding normal breast tissues. The results showed that HCG18 was significantly upregulated in BC tissues compared with normal mammary tissues (Figures 1A,B). qRT-PCR was applied to detect HCG18 expression in a panel of BC cell lines and the mammary epithelial cell line MCF-10A. Figure 1C shows that MDA-MB-231 cells, which are highly invasive BC cells, had the highest HCG18 expression level, and HCG18 expression was lower in MCF-7 cells with low malignant potential. However, MCF-10A cells had the lowest HCG18 expression among all assessed BC cells. Then, we sought to assess the associations between HCG18 expression and clinicopathologic features in BC patients. We divided the samples into a HCG18 high-expression group and HCG18 low-expression group according to the average value of HCG18 expression in BC tissues. The results in Figures 1D,E and Table 1 show that HCG18 was positively correlated with the histological and clinical stage, tumor size, and axillary lymph node metastasis in BC patients. However, there were no differences in HCG18 expression in different subtypes of BC (Figure 1E), which was further confirmed by data from the lnCAR database (Supplementary Figures S1A–C). Figures 1F,G show that BC patients with high HCG18 expression had poor distant metastasis-free survival (DMFS) and overall survival (OS), which was also supported by survival information in the Kaplan–Meier Plotter database (Figure 1H). Collectively, these results revealed that HCG18 was commonly overexpressed in BC patients and that patients with high HCG18 expression tended to have a higher risk of metastasis and a poorer prognosis than those with low expression.

To investigate the biological roles of HCG18 in BC, we used MDA-MB-231, which has high HCG18 expression, and MCF-7 cells, which have low HCG18 expression, for further experiments. Three sets of specific shRNAs targeting HCG18 were transfected into MDA-MB-231 cells to generate HCG18 reducing cells (MDA-MB-231 ^HCG18-sh1 , MDA-MB-231 ^HCG18-sh2 , and MDA-MB-231 ^HCG18-sh3 ), and HCG18 was stably overexpressed in MCF-7 cells (MCF-7 ^HCG18 ). The HCG18 expression profile in these newly generated cells was detected by qRT-PCR. Supplementary Figure S1D shows that HCG18-sh1 and HCG18-sh2 yielded a better reducing efficiency than HCG18-sh3 in MDA-MB-231 cells, and HCG18 was significantly upregulated in MCF-7 ^HCG18 cells, so MDA-MB-231 ^HCG18-sh1 , MDA-MB-231 ^HCG18-sh2 , and MCF-7 ^HCG18 cells were used in the following experiments. The CCK-8 assay is a viability assay that provides a readout, that is, proportional to cell number; this approach is often applied to detect the proliferative capacity of cancer cells. We used CCK-8 assays to evaluate the effect of HCG18 alterations on proliferation. Figure 2A shows that the proliferation capability was significantly decreased after reducing HCG18 in MDA-MB-231 cells, whereas MCF-7 ^HCG18 cells exhibited a better proliferation ability than the control cells. Colony formation assays (Figure 2B) and EdU staining assays (Figure 2C) further confirmed this conclusion. Then, the expression of Ki-67 (the nuclear expression of Ki-67 can be evaluated to assess tumor proliferation capability; Ki-67 >20% was regarded as high Ki-67 expression profile) was detected in BC tissues by IHC, and the χ ² test was used to analyze the relationship between HCG18 and Ki-67 expression. Figure 2D shows the positive correlation between HCG18 and Ki-67 expression in clinical samples of BC patients. To verify the effect of HCG18 on tumorigenesis in vivo, MDA-MB-231 ^NC and MDA-MB-231 ^HCG18-sh1 cells were orthotopically injected into the mammary fat pads of BALB/c nude mice. As shown in Figure 2E, reducing HCG18 significantly suppressed tumor growth and prolonged tumor-free survival compared with that in the control group. Taken together, these results demonstrated that HCG18 could promote BC cell proliferation both in vitro and in vivo.

The results in Table 1 show that there was a close relationship between HCG18 expression and the status of axillary lymph node metastasis in BC patients, so we focused on the prometastatic function of HCG18 in BC. Wound healing assays showed that the migration ability was extremely decreased after reducing HCG18 in MDA-MB-231 cells, and the migration ability of MCF-7 ^HCG18 cells was increased in comparison with that of the control group (Figure 3A). Matrigel invasion assays also showed similar results (Figure 3B). Western blotting assays revealed that the epithelial-related protein E-cadherin was upregulated, but the mesenchymal marker vimentin and the metastatic markers MMP-2 and MMP-9 were downregulated in MDA-MB-231 ^HCG18-sh cells compared with control cells. The opposite results were observed in MCF-7 ^HCG18 cells (Figure 3D and Supplementary Figure S1E).

In recent years, a growing number of studies have reported that CSPs play important roles in tumor survival, proliferation, invasion, immune escape, and drug resistance. Thus, the following experiments explored the relationship between HCG18 expression and CSPs in BC cells. Sphere formation assays revealed that the sphere formation ability of MDA-MB-231 ^HCG18-sh cells was obviously decreased compared with that of the control cells; however, the CSPs were remarkably enhanced after overexpressing HCG18 in MCF-7 cells (Figure 3C). Western blotting assays showed that the CSP markers CD44, ABCG2, and OCT4 were markedly downregulated in MDA-MB-231 ^HCG18-sh cells and increased in MCF-7 ^HCG18 cells compared with the control cells (Figure 3D and Supplementary Figure S1E). Finally, lung metastasis mouse models were used to investigate the prometastatic function of HCG18 in vivo. The results in Figures 3E–G show that mice injected with MDA-MB-231 ^HCG18-sh cells exhibited fewer metastatic nodes than the mice injected with control cells, which indicates that HCG18 could promote BC cell lung metastasis in vivo. Collectively, these results confirmed that HCG18 could endow BC cells with CSPs in vitro and promote BC cell metastasis both in vitro and in vivo.

E2s play important roles in regulating cellular posttranslational protein modifications and are involved in a wide range of key physiological or pathologic processes. Multiple studies have reported that aberrant E2 expression is commonly observed in a variety of human cancers. Some cancer-related E2s could facilitate DNA repair and cell cycle progression and activate oncogenic signaling pathways, which promote cancer progression and metastasis (Voutsadakis, 2013). Our previous study reported that UBE2O could promote proliferation and EMT and endow BC cells with CSPs by degrading AMPKα2, thus activating the mTORC1-MYC axis (Liu et al., 2020b). For this reason, we hypothesized that there may be a potential interaction network between HCG18 and E2s in BC. Then, a panel of cancer-related E2s was detected in MDA-MB-231^NC/MDA-MB-231 ^HCG18-sh and MCF-7 ^NC /MCF-7 ^HCG18 cells. Coincidentally, we found that there was a close positive correlation between HCG18 and UBE2O expression in the indicated cells (Figure 4A and Supplementary Figure S1F). Correlation analysis with the Gene Expression Profiling Interactive Analysis (GEPIA) database showed that there was a positive correlation between HCG18 and UBE2O expression in BC (Figure 4B), which was further verified in clinical samples (Figures 4C,D). Next, we reduced UBE2O in MCF-7 ^HCG18 cells (generating MCF-7 ^{HCG18+UBE2O−sh} cells) and overexpressed UBE2O in MDA-MB-231 ^HCG18-sh1 cells (generating MDA-MB-231 ^{HCG18-sh+UBE2O} cells). Western blotting assays showed that UBE2O was remarkably decreased in MDA-MB-231 ^HCG18-sh cells and that the expression level of UBE2O was increased in MCF-7 ^HCG18 cells (Figure 4E and Supplementary Figure S2A). The results in Figure 4F and Supplementary Figure S2B revealed that the upregulation of E-cadherin and the downregulation of vimentin, MMP-2, MMP-9, CD44, and OCT4 were attenuated in MDA-MB-231 ^{HCG18-sh+UBE2O} cells, and the opposite results were observed in MCF-7 ^{HCG18+UBE2O−sh} cells compared with control cells.

Then, we explored the effect of UBE2O on the protumor function of HCG18 in BC cells in vitro. CCK-8 (Figure 5A), colony formation assays (Figure 5B), and EdU staining assays (Figure 5C) showed that overexpressing UBE2O could significantly reverse the decline in the proliferation ability of MDA-MB-231 ^HCG18-sh cells, and the enhancement of proliferative capacity was attenuated after reducing UBE2O in MCF-7 ^HCG18 cells. Figures 5D–F show that the migration, invasion, and sphere formation capabilities of MDA-MB-231 ^{HCG18-sh+UBE2O} cells were improved compared with those of MDA-MB-231 ^HCG18-sh cells, and reducing UBE2O in MCF-7 ^HCG18 cells remarkably destroyed these capabilities in comparison with those of the control cells. Taken together, these results confirmed that UBE2O plays a vital role in the cancer-promoting function of HCG18 in BC cells.

To further clarify the mechanism by which HCG18 regulates UBE2O in BC cells, we first ascertained the cellular location of HCG18 in BC cells. Subcellular fractionation assays showed that HCG18 was predominantly distributed in the cytoplasm of BC cells (Figure 6A), which was further confirmed by the lncLocator database (Supplementary Figure S2C) and was in accordance with previous studies (Xu et al., 2019; Li et al., 2020a). Recently, many studies confirmed that cytoplasmic lncRNAs could affect the biological process of BC cells by modulating the expression of miRNAs in a ceRNA-dependent manner (Liang et al., 2020; Ni et al., 2020). Therefore, we speculated that HCG18 could influence the stability of UBE2O mRNA via a ceRNA-mediated mechanism. To validate this, TargetScan, miRwalk, and starBase were used to predict the potential target miRNAs, and the results in Figure 6B revealed that there were three miRNAs (miR-30a-5p, miR-34a-5p, and miR-103a-3p) connected with both HCG18 and UBE2O. Then, we reduced HCG18 in MDA-MB-231 and MCF-7 cells, and qRT-PCR was applied to detect the changes in the expression of the three miRNAs above. The results showed that compared with that in the control group, the expression level of miR-103a-3p was remarkably increased after reducing HCG18 in both MDA-MB-231 and MCF-7 cells. However, there were no changes in the expression of the other two miRNAs in the indicated cells (Figure 6C). RIP assays showed that miR-103a-3p could associate with HCG18 via complementary binding sites, whereas the nontargeting miR-NC did not associate with HCG18 (Figures 6D,E). RIP assays also confirmed that miR-103a-3p could specifically bind to the UBE2O 3′UTR (Figure 6F). Dual-luciferase assays revealed that transfection of miR-103a-3p mimics could significantly reduce the luciferase activity of the HCG18-WT group but failed to affect that of the mutant group (Figure 6G). miRNAs degrade targeted mRNA in an Ago2-dependent manner by binding their targets. Next, we performed RIP in MDA-MB-231 and MCF-7 cells transiently overexpressing miR-103a-3p to pull down HCG18 using anti-Ago2 antibodies or control IgG, and qRT-PCR was used to analyze the enrichment of HCG18 in immunoprecipitates. The amount of HCG18 pulled down with anti-Ago2 antibodies was remarkably increased in cells transfected with miR-103a-3p mimics compared with control cells (Figure 7A). Then, the expression level of miR-103a-3p was detected in BC tissues and corresponding normal tissues. The results showed that miR-103a-3p was obviously downregulated in BC tissues compared with normal breast tissues (Figure 7B). Finally, we found that there was a negative relationship between miR-103a-3p and HCG18 in BC tissues (Figure 7C). Collectively, these results revealed that HCG18 acts as a sponge for miR-103a-3p in BC cells.

Then, we validated whether miR-103a-3p could mediate the function of HCG18 in promoting UBE2O expression. The results in Figure 7D and Supplementary Figure S2D show that ectopic expression of miR-103a-3p by miRNA mimics obviously reduced UBE2O expression in the indicated cells. Dual-luciferase assays further revealed that overexpression of miR-103a-3p could decrease the luciferase activity of the wild-type UBE2O reporter but not the mutant UBE2O reporter, which indicated that UBE2O was a direct target of miR-103a-3p (Figure 7E). The correlation analysis in Figure 7F identified that UBE2O expression was negatively correlated with miR-103a-3p expression in BC tissues. Finally, we detected the biological functions of miR-103a-3p in BC cells. EdU staining assays (Supplementary Figure S2E), invasion assays (Supplementary Figure S2F), and scratch assays (Supplementary Figure S2G) demonstrated that miR-103a-3p could significantly abolish the proliferation, migration, and invasion capabilities of MCF-7 cells after upregulating HCG18 in MCF-7 cells. The results in Supplementary Figure S2H revealed that the decreased expression of E-cadherin and increased expression of vimentin, MMP-2, MMP-9, CD44, and OCT4 were remarkably reversed by miR-103a-3p mimics. Taken together, these results claimed that HCG18 promoted UBE2O expression by sponging miR-103a-3p in human BC.

As we verified that HCG18 was upregulated in BC, subsequent experiments were carried out to explore the regulatory mechanism resulting in the aberrant expression of HCG18 in BC. The JASPAR database was applied to identify the potential HCG18 transcription factors, and coincidentally, we found that there were four potential binding sites for HIF-1α in the promoter region of HCG18 (Figure 8A). Our previous study demonstrated that UBE2O could activate the mTORC1 signaling pathway by meditating AMPKα2 ubiquitination and that mTORC1 could regulate HIF-1α expression, which in turn promoted cell growth and anabolism (Laplante and Sabatini, 2012; Vila et al., 2017). Therefore, HIF-1α was reduced in MDA-MB-231 and MCF-7 cells. The efficiency of HIF-1α silencing and the relative HCG18 expression were detected by qRT-PCR. The results in Figure 8B revealed that HCG18 expression was significantly decreased after reducing HIF-1α in the indicated cells. Correlation analysis showed that there was a positive relationship between HIF-1α and HCG18 expression in BC tissues (Figure 8C), which was further confirmed with the GEPIA database (Figure 8D). Then, four sets of luciferase reporter plasmids including the wild-type or mutant HCG18 promoter region were constructed, and dual-luciferase assays were performed to validate the potential HIF-1α binding sites. For site 2, the transfection of HIF-1α plasmids could significantly increase the luciferase activity of the WT group but not the mutant group. For the other three sites, however, there were no differences in luciferase activity between the WT and mutant groups after HIF-1α plasmid transfection (Figure 8E). These results indicated that HIF-1α could bind to the promoter region of HCG18 (site 2) and facilitate HCG18 transcription. This conclusion was further confirmed by ChIP assays (Figure 8F). Taken together, these results demonstrate that HIF-1α could promote HCG18 expression, thus forming a positive feedback loop in BC cells (Figure 8G).