RETRACTED ARTICLE: SP1-induced lncRNA AGAP2-AS1 expression promotes chemoresistance of breast cancer by epigenetic regulation of MyD88

Primary cancer tissue and adjacent noncancerous tissue samples were collected from a cohort of 42 patients with breast cancer (male/female: 0/42, range of age (median): 38–61 (47)), and another independent cohort of 67 HER2⁺ breast cancer patients who received trastuzumab treatment (male/female: 0/67, range of age (median): 47–82 (55)). The diagnoses of all the patients were pathologically confirmed, and the clinical tissue samples were collected before chemotherapy was started at Hainan General Hospital and The Second Affiliated Hospital of Chongqing Medical University. The samples were obtained during operation and immediately frozen at − 80 °C until RNA extraction. The written informed consents obtained from all patients were approved according to the guidelines revised by the Hainan General Hospital.

The human breast cancer cell lines SKBR-3 and BT474, which harbor HER2-activating mutations, were purchased from Chinese Type Culture Collection, Chinese Academy of Sciences (Shanghai, China). Both the cell lines were cultured in DMEM (BioWhittaker, Lonza, USA) supplemented with 1 mM L-glutamine, 100 U/ml penicillin/streptomycin (BioWittaker, Lonza) and heat-inactivated 10% fetal bovine serum (FBS, Gibco) at 37 °C in a humidified incubator with 5% CO₂. Trastuzumab (Herceptin) was obtained from Roche (Basel, Switzerland) and dissolved in the enclosed sterile water. The trastuzumab-resistant SKBR-3/Tr and BT474/Tr cells were obtained by continuous culture with 5 μg/mL trastuzumab for 6 months as previously reported [16‐18].

The full-length of lncRNA AGAP2-AS1, and the coding sequence of MyD88 and CREB-binding protein (CBP) were amplified and cloned into the lentivirus vector (Lv-AGAP2-AS1, Lv-MyD88, and Lv-CBP, respectively) for retrovirus production using BT474 cells by GeneChem (Shanghai, China). The negative control vectors were also generated. The lentivirus vector containing small hairpin RNA (shRNA) sequence targeting MyD88 (sh-MyD88), AGAP2-AS1 (sh-AGAP2-AS1) or negative control vector (sh-NC) was also amplified and cloned by GeneChem. All the vectors were labeled with green fluorescence protein (GFP). Transfection was carried out using Lipofectamine 3000 (Invitrogen, Carlsbad, CA, USA) following the manufacturer’s instructions. Transfection efficiency was evaluated in every experiment by RT-qPCR after 24 h to ensure that cells were transfected. Functional experiments were then performed after sufficient transfection for 48 h. The sequences of shRNAs are shown in the Additional file 1: Table S1.

The RNA was reverse transcribed using the SuperScript III® (Invitrogen) and then the obtained cDNAs were quantified using RT-qPCR assay labeled with SYBR (Takara Bio Company, Dalian, China) on BioRad CFX96 Sequence Detection System (BioRad company, Berkeley, CA). The gene expression levels were normalized using GAPDH expression. The RT-qPCR results were analyzed and expressed relative to the CT (threshold cycle) values, and then converted to fold changes. All the primer sequences were synthesized by RiboBio (Guangzhou, China), and their sequences are shown in Additional file 1: Table S1.

The altered cell viability after transfection was assayed using the MTT Kit (Dojindo, Rockville, MD, USA). In brief, cells were seeded into a 96-well plate and then treated with silencing or overexpressing vectors for 48 h. Next, the cells were treated with the MTT reagent and further cultured for 2 h. The optical density at 450 nm was measured with a spectrophotometer (Thermo Electron Corporation, MA, USA). The percentage of the control samples for each cell line was calculated thereafter.

ChIP was performed using the EZ ChIP™ Chromatin Immunoprecipitation Kit (Millipore, Burlington, MA) according to the manufacturer’s protocol. Briefly, cross-linked chromatin was sonicated into 200–1000 bp fragments. The chromatin was immunoprecipitated using anti-H3K27ac (Abcam, ab4729, Cambridge, MA)and anti-CBP antibodies (Abcam, ab2832). Magna RIP™ RNA-Binding Protein Immunoprecipitation Kit (Millipore) was used for RIP and anti-CBP antibody was used to pull down AGAP2-AS1.

The cells were fixed in 4% formaldehyde for 15 min and then washed with PBS. The fixed cells were then treated with pepsin and dehydrated through ethanol, and further permeabilized in Triton X100 (Sigma-Aldrich) for 20 min. Goat serum was used for blocking, and then cells were incubated with the anti-Ki67 antibody (Abcam, ab15580, 1:500, Cambridge, MA) overnight at 4 °C. The primary antibody was washed off, and then the cells were incubated with the appropriate rhodamine-conjugated secondary antibody for 1 h. The cells were again washed and incubated with DAPI (Invitrogen) for nuclear staining. The cells were visualized for immunofluorescence with a fluorescence microscope (DMI4000B, Leica).

The nuclear and cytosolic fraction separation was performed using the PARIS kit (Life Technologies), and RNA FISH probes were designed and synthesized according to the manufacturer’s instructions. Briefly, cells were fixed in 4% formaldehyde for 15 min and then washed with PBS. The fixed cells were treated with pepsin and dehydrated through ethanol. The air-dried cells were incubated further with 40 nM of the FISH probe in hybridization buffer (Life Technologies). After hybridization, the slide was washed, dehydrated and mounted with Prolong Gold Antifade Reagent with DAPI for detection. The slides were visualized for immunofluorescence with a fluorescence microscope (DMI4000B, Leica).

The TUNEL staining was performed to evaluate cell apoptosis. In brief, cells were fixed by using 4% formaldehyde followed by staining with the TUNEL kit according to the manufacturer’s instructions (Vazyme, TUNEL Bright-Red Apoptosis Detection Kit, A113). The TUNEL-positive cells were counted under the fluorescence microscope (DMI4000B, Leica).

Male BALB/C nude mice (6 weeks of age) were purchased from Shanghai SIPPR-BK Laboratory Animal Co. Ltd. (Shanghai, China) and maintained in microisolator cages. Mice were housed in a facility-controlled, pathogen-free conditions under 28 °C, 50% humidity and were fed ad libitum with sterile chow food and water. All efforts were made to minimize suffering and the research protocol was approved by Hainan General Hospital of Jinan University based on Ethics in the Care and Use of Laboratory Animals. 1 × 10⁷ BT474 cells stably transfected with Lv-NC/Lv-AGAP2-AS1, or co-transfected with sh-MyD88/sh-NC and Lv-AGAP2-AS1 were suspended in 110 μL of serum-free DMEM, and then injected subcutaneously in the flank. When tumors were palpable, mice were treated by administering 5 mg/kg trastuzumab or PBS (in case of control) intraperitoneally once every 2 days for 3 weeks. Herein, six mice xenograft treatment groups were established: (Lv-NC-transfected cells + PBS, (Lv-AGAP2-AS1-transfected cells + PBS), (Lv-NC-transfected cells + trastuzumab treatment), (Lv-AGAP2-AS1-transfected cells + trastuzumab treatment), (Lv-AGAP2-AS1/sh-NC-cotransfected cells + trastuzumab treatment) and (Lv-AGAP2-AS1/sh-MyD88-cotransfected cells + trastuzumab treatment). Six mice were included in each group and more than three mice remained at the end of the study, excluding mice that were dead or had complications, such as skin necrosis due to infection. At the end of the treatment that lasts for 3 weeks, the xenograft tumor was stripped, and the mass was calculated. The tumor size was evaluated using a standard caliper measuring tumor length and width in a blinded manner and the tumor volume was calculated using the formula: length × width² × 0.52.

Immunohistochemical staining was performed on 4-μm-thick sections. Briefly, the slides were deparaffinized and antigen retrieval was performed in a steam cooker for 1.5 min in 1 mM EDTA. Rabbit anti-MyD88 antibody (Abcam, ab2064) at 1:150 dilution was added and incubated overnight at 4 °C. The universal secondary antibody (DAKO) was applied for 15 min at room temperature. Diaminobenzidine or 3-amino-9-ethylcarbazole was used as chromogens and slides were counterstained with hematoxylin before mounting.

The cell lysates were prepared with RIPA buffer containing protease inhibitors (Sigma-Aldrich). The membranes were incubated overnight at 4 °C with of the primary antibodies at a dilution of 1:1000. A secondary antibody was then used for immunostaining for 1 h at room temperature. The primary antibodies used in this study are anti-MyD88 (Abcam, ab2064), anti-CBP (Abcam, ab50702), anti-SP1 (Abcam, ab124804) anti-NF-κB p65 (Abcam, ab16502), and anti-β-actin (Abcam, ab6276) antibody.

The RT-qPCR analysis was performed to detect the expression of AGAP2-AS1 in the breast cancer cells. As shown in Fig. 1a, AGAP2-AS1 was upregulated in the breast cancer cell lines when compared to the normal breast epithelium MCF-10A cells. To investigate whether AGAP2-AS1 regulates trastuzumab resistance, we established two trastuzumab-resistant sub-lines derived from HER2⁺ SKBR-3 and BT474 cells (SKBR-3/Tr and BT474/Tr, respectively). Compared with the parental cells, the established resistant cells showed less response to trastuzumab treatment, as evidenced by increased IC₅₀ values and elevated cell viability (Fig. 1b, c). Moreover, AGAP2-AS1 was upregulated in trastuzumab-resistant cells than in the respective parental cells (Fig. 1d). This indicates that AGAP2-AS1 may be critical for breast cancer progression.

There have been evidence indicating that transcription factors (TFs) play an important role in lncRNA dysregulation, and hence, we searched for transcription factors that might be linked to lncRNA dysregulation. By using the online transcription factor prediction software JASPAR (http://​jaspar.​genereg.​net/), we found that there are 11 SP1 binding sites in the promoter region of AGAP2-AS1 (Fig. 1e). Previously, Qi et al. demonstrated that AGAP2-AS1 is activated by SP1 in gastric cancer [19]. Therefore, we investigated whether this interaction also applies to breast cancer. We identified that SP1 was upregulated in the trastuzumab-resistant cells when compared to the parental cells at both transcript and protein levels (Fig. 1f). Transfection of SP1-overexpression vector dramatically increased AGAP2-AS1 expression levels (Fig. 1g). Consistently, the immunofluorescence assay showed that SP1 enrichment significantly increased in the nucleus of the SKBR-3 resistant cells when compared to the parental cells (Fig. 1h). We also performed ChIP assay to further verify the enrichment of SP1 at the promoter region of AGAP2-AS1. As expected, SP1 was enriched and the enrichment significantly increased in the resistant cells in contrast to the parental cells (Fig. 1i). In addition, the AGAP2-AS1 promoter region including 2 binding sites of SP1 with a high score was inserted into a PGL3 vector (Fig. 1j), and dual-luciferase reporter assay indicated that SP1 significantly promoted the luciferase activity (Fig. 1k). These results indicated that the upregulation of AGAP2-AS1 in breast cancer cells may be induced by SP1.

Next, we investigated the functional role of AGAP2-AS1 in proliferation and apoptosis of breast cancer cell lines. The expression studies revealed that endogenous expression of AGAP2-AS1 in SKBR-3 breast cancer cell line was high while BT474 showed a low endogenous expression. Therefore, we constructed the AGAP2-AS1 overexpression model by using BT474 cells and the AGAP2-AS1 knockdown model by using SKBR-3 cells (Fig. 2a, b). The results from the MTT assay, revealed that BT474 cells overexpressed with AGAP2-AS1 showed significantly elevated levels of cell proliferation compared to negative controls, while knockdown of AGAP2-AS1 in SKBR-3 cells decreased cell proliferation (Fig. 2c). Further in the colony formation assay, the number of colonies formed was much higher in Lv-AGAP2-AS1-BT474 cells than Lv-NC-BT474 cells. However, a suppressed colony formation ability was identified in sh-AGAP2-AS1-SKBR-3 cells when compared to sh-NC-SKBR-3 cells (Fig. 2d). To confirm the effect of AGAP2-AS1 on cell growth, we investigated the Ki-67 expression level by immunofluorescence assay. As expected, an enhanced AGAP2-AS1 promoted Ki-67 expression was observed, whereas the AGAP2-AS1 knockdown dramatically silenced the level of Ki-67 (Fig. 2e). In addition, FACS apoptosis assay showed that overexpression of AGAP2-AS1 suppressed apoptosis whereas knockdown of AGAP2-AS1 exerted an opposite effect (Fig. 2f). To conclude, we demonstrated that AGAP2-AS1 played an oncogenic role in breast cancer.

We also investigated whether AGAP2-AS1 regulates trastuzumab resistance. As expected, knockdown of AGAP2-AS1 promoted the trastuzumab-induced (0.5 μg/mL) cell cytotoxicity in the SKBR-3/Tr cells (Fig. 2g). In addition, overexpression of AGAP2-AS1 partially abrogated the effects of trastuzumab on cell viability in the BT474 parental cells (Fig. 2h). The TUNEL assay was then performed to analyze whether AGAP2-AS1 influenced the nuclear apoptosis induced by trastuzumab. We found that the overexpression of AGAP2-AS1 suppressed the trastuzumab-induced cell apoptosis in the BT474 parental cells whereas knockdown of AGAP2-AS1 enhanced the apoptosis caused by trastuzumab in the SKBR-3/Tr cells (Fig. 2i).

Based on the understanding of the pathologic role of AGAP2-AS1, we continued to explore the underlying regulatory mechanisms. RNA-pull down experiments were performed to search for the AGAP2-AS1-associated downstream proteins, and we found that MyD88 was enriched (Fig. 3a). MyD88 is an adaptor molecule for toll-like receptor (TLR) and interleukin 1 receptor (IL-1R) signaling implicated in tumorigenesis and resistance to chemotherapeutic drugs through proinflammatory mechanisms (Fig. 3b). Therefore, we assume that MyD88 could be important for trastuzumab resistance. In this study, we verified that the overexpression of AGAP2-AS1 upregulated the expression of MyD88 at both mRNA and protein levels (Fig. 3c), whereas knockdown of AGAP2-AS1 downregulated its expression (Fig. 3d). In addition, MyD88 was also upregulated in the trastuzumab-resistant cells when compared to their respective parental cells (Fig. 3e), strongly indicating that MyD88 may serve as a direct target of AGAP2-AS1.

To investigate whether MyD88 is a functional target of AGAP2-AS1, we constructed MyD88-overexpression vector and MyD88-knockdown vector (Fig. 3f). Then, we modulated the expression of endogenous AGAP2-AS1 and MyD88 simultaneously. The MTT assay revealed that silencing MyD88 significantly abrogated the effects of Lv-AGAP2-AS1 on cell growth, while overexpression of MyD88 reversed the effect induced by sh-AGAP2-AS1 (Fig. 3g). More importantly, Lv-MyD88 abrogated the sh-AGAP2-AS1-induced trastuzumab response in the SKBR-3/Tr cells whereas co-transfection of sh-MyD88 reversed the trastuzumab resistance induced by Lv-AGAP2-AS1 in the BT474 parental cells (Fig. 3h). Taken together, the lncRNA AGAP2-AS1 may promote trastuzumab resistance via binding to MyD88.

To further understand the regulation of MyD88 by AGAP2-AS1, we explored the probable mechanisms by analysis of the ENCODE database (http://​genome.​ucsc.​edu/​). As shown in Fig. 4a, there is a high concentration of H3K27ac in the promoter region of MyD88, indicating that MyD88 may be regulated at the transcriptional level via histone modification. To test this hypothesis, we performed a ChIP assay by using breast cancer parental and trastuzumab-resistant cells. As shown in Fig. 4b, an increased enrichment of H3K27ac was verified in breast cancer cells compared to normal cells. Moreover, an increased enrichment of H3K27ac was also verified in trastuzumab-resistant cells in contrast to the parental cells (Fig. 4c). We next treated breast cancer cells with C646, a histone acetyltransferase (HAT) inhibitor, and found a significant reduction in the MyD88 expression in response to the treatment (Fig. 4d). To this end, we demonstrated that MyD88 was upregulated in breast cancer cells due to the histone acetylation in its promoter region.

To further investigate whether lncRNA AGAP2-AS1 participates in the H3K27 acetylation at the MyD88 promoter, we identified the cellular localization of AGAP2-AS1 in breast cancer cells. FISH assay with a specific probe of AGAP2-AS1 confirmed that AGAP2-AS1 was mainly distributed in the nuclear section of both cells (Fig. 4e). Then, we detected whether AGAP2-AS1 regulates the enrichment of H3K27ac at the promoter region of MyD88. As expected, upregulation of AGAP2-AS1 in the BT474 cells increased the enrichment of H3K27ac whereas knockdown of AGAP2-AS1 resulted in a decrease in H3K27ac enrichment in the SKBR-3 cells (Fig. 4f). Therefore, we conclude that the lncRNA AGAP2-AS1 regulates MyD88 expression via the histone H3K27 acetylation.

As CREB-binding protein (CBP) is an important enzyme that participates in chromatin acetylation, we explored whether CBP is essential for the AGAP2-AS1-regulated H3K27 acetylation of MyD88. As expected, overexpression of CBP using specific plasmids (Fig. 5a) dramatically increased the expression of MyD88 at both transcript and protein levels (Fig. 5b). The ChIP analysis showed that CBP was enriched at the MyD88 promoter regions (Fig. 5c), indicating that CBP may be essential for histone acetylation. On account of the above result that both AGAP2-AS1 and CBP could regulate H3K27 acetylation at the promoter of MyD88, we then tested whether AGAP2-AS1 and CBP are functionally linked during the histone acetylation process. We detected the CBP expression after knockdown or overexpression of AGAP2-AS1 and found that AGAP2-AS1 did not affect the expression level of CBP (Fig. 5d), so we hypothesized that AGAP2-AS1 may exert the oncogenic function by recruiting CBP to target genes. To test this assumption, we conducted RIP assay. As shown in Fig. 5e, an enriched AGAP2-AS1 was identified by anti-CBP antibody in contrast to anti-IgG. More importantly, AGAP2-AS1 influenced the enrichment of CBP and H3K27ac in the MyD88 promoter region (Fig. 5f, g). These results indicate that the binding of AGAP2-AS1 with CBP promoted the enrichment of H3K27ac, thereby resulting in the activation of MyD88 transcription.

To validate the in vitro data of lncRNA AGAP2-AS1, we established a model of nude mice bearing the BT474 xenograft. The BT474 cells infected with Lv-AGAP2-AS1 or Lv-NC were planted into the flanks of nude mice. Four treatment groups of mice xenograft were established: Group I (Lv-NC-transfected cells + PBS), Group II (Lv-AGAP2-AS1-transfected cells + PBS), Group III (Lv-NC-transfected cells + trastuzumab treatment) and Group IV (Lv- AGAP2-AS1-transfected cells + trastuzumab treatment). Then, tumors were stripped and presented as in Fig. 6a. The results showed that AGAP2-AS1 promoted tumor growth in xenograft model (Group II vs. Group I). Trastuzumab treatment significantly suppressed the growth of tumor cells when compared to the control groups (Group III vs. Group I). With the treatment of trastuzumab, the tumor cells that were infected with Lv-AGAP2-AS1 grew faster than Lv-NC-transfected cells (Group IV vs. Group III), suggesting that AGAP2-AS1 promoted trastuzumab resistance in vivo (Fig. 6b).

In addition, IHC experiment was conducted to verify whether AGAP2-AS1 affects the expression of MyD88 in xenograft tumor tissues. As shown in Fig. 6c, overexpression of AGAP2-AS1 promoted the expression of MyD88 in tumor tissues (Group II vs. Group I, or Group IV vs. Group III, respectively), indicating that AGAP2-AS1 regulates tumor growth and trastuzumab resistance via targeting MyD88 in vivo. To directly verify whether MyD88 could rescue the effects of AGAP2-AS1 in vivo, we injected the BT474 cells that were co-transfected with sh-MyD88 and Lv-AGAP2-AS1 followed by trastuzumab treatment as described above. As expected, co-transfection of sh-MyD88 dramatically reversed the Lv-AGAP2-AS1-induced trastuzumab resistance in vivo (Fig. 6d), further confirming that MyD88 is essential for the carcinogenic function of AGAP2-AS1.

There is sufficient evidence indicating that MyD88 regulates cancer cell proliferation, apoptosis and chemoresistance by the activation of NF-κB pathway. As we have proved that lncRNA AGAP2-AS1 promotes trastuzumab resistance via enhancing MyD88 expression by modifying histone acetylation, we hypothesized that the NF-κB pathway may be involved in the oncogenic role of AGAP2-AS1 and MyD88. Western blot experiment was performed to test this hypothesis, and we found that NF-κB activity was suppressed in sh-AGAP2-AS1-SKBR-3 cells, while ectopic expression of MyD88 rescued this effect (Fig. 7a). Conversely, NF-κB activity increased in the BT474 cells transfected with Lv-AGAP2-AS1 compared to the negative control cells, while silencing of MyD88 diminished this enhanced activity induced by AGAP2-AS1 overexpression (Fig. 7b). To this end, we demonstrated that AGAP2-AS1 promoted tumor growth via upregulation of MyD88 expression and further activation of NF-κB signaling pathway.

We detected the expression of AGAP2-AS1 in 42 breast cancer tissues and paired adjacent non-tumor tissues. As shown in Fig. 8a, AGAP2-AS1 expression frequently increased in cancerous tissues and 69% of the cancer samples displayed increased AGAP2-AS1 expression in our study. MyD88 was upregulated in 76.2% of breast cancer patients. In addition, correlation analysis by using Spearman testing confirmed a significant positive correlation between the expression of AGAP2-AS1 and MyD88 in these specimens (Fig. 8b).

To verify the clinical role of AGAP2-AS1 in trastuzumab-treated patients, we collected 62 cancer tissues from advanced HER2⁺ breast cancer patients who received trastuzumab treatment. Patients were divided into responding (CR + PR, 33 patients) and non-responding (SD + PD, 29 patients) groups according to the immuno-related Response Evaluation Criteria In Solid Tumors (irRECIST) [20]. RT-qPCR revealed that AGAP2-AS1 was upregulated in the non-responding group (Fig. 8c). We then investigated the value of AGAP2-AS1 in differentiating responding patients from non-responding patients by establishing ROC curve. As shown in Fig. 8d, the area under the curve (AUC), diagnostic sensitivity, and specificity reached 0.753, 78.7 and 63.7% with the established cut-offs (3.78), respectively. Under this stratification criteria (3.78), patients were divided into a low and a high AGAP2-AS1 expression groups, and the proportion of patients not responding to chemotherapy was significantly higher in the high AGAP2-AS1 expression group than in the low expression group (Fig. 8e). Altogether, our clinical results indicate that AGAP2-AS1 may be a promising prognostic marker to discriminate responders and non-responders to trastuzumab therapy.