Pentapeptide PYRAE triggers ER stress-mediated apoptosis of breast cancer cells in mice by targeting RHBDF1-BiP interaction

Abstract

Reinforced cellular responses to endoplasmic reticulum (ER) stress are caused by a variety of pathological conditions including cancers. Human rhomboid family-1 protein (RHBDF1), a multiple transmembrane protein located mainly on the ER, has been shown to promote cancer development, while the binding immunoglobulin protein (BiP) is a key regulator of cellular unfolded protein response (UPR) for the maintenance of ER protein homeostasis. In this study, we investigated the role of RHBDF1 in maintaining ER protein homeostasis in breast cancer cells. We showed that deleting or silencing RHBDF1 in breast cancer cell lines MCF-7 and MDA-MB-231 caused marked aggregation of unfolded proteins in proximity to the ER. We demonstrated that RHBDF1 directly interacted with BiP, and this interaction had a stabilizing effect on the BiP protein. Based on the primary structural motifs of RHBDF1 involved in BiP binding, we found a pentapeptide (PE5) targeted BiP and inhibited BiP ATPase activity. SPR assay revealed a binding affinity of PE5 toward BiP (K_d = 57.7 μM). PE5 (50, 100, 200 μM) dose-dependently promoted ER protein aggregation and ER stress-mediated cell apoptosis in MCF-7 and MDA-MB-231 cells. In mouse 4T1 breast cancer xenograft model, injection of PE5 (10 mg/kg, s.c., every 2 days for 2 weeks) significantly inhibited the tumor growth with markedly increased ER stress and apoptosis-related proteins in tumor tissues. Our results suggest that the ability of RHBDF1 to maintain BiP protein stability is critical to ER protein homeostasis in breast cancer cells, and that the pentapeptide PE5 may serve as a scaffold for the development of a new class of anti-BiP inhibitors.

Introduction

It has been well documented that cancer cells are capable of sustaining active protein synthesis under various conditions of intracellular or extracellular stress, especially endoplasmic reticulum (ER) stress [1–3]. Cancer cells under ER stress can activate the unfolded protein response (UPR), which involves a number of protein machineries, including the binding immunoglobulin protein (BiP; also known as HSPA5 or GRP78) and signaling pathways related to inositol-requiring enzyme 1 (IRE1), PRKR-like ER kinase (PERK), and activating transcription factor 6 (ATF6) [2, 4–9]. BiP, a major molecular chaperone of the UPR, functions in assisting proteins in folding correctly and is important in reducing unfolded protein accumulation and restoring ER homeostasis [10–13]. BiP expression in cancer cells is increased in response to UPR sensor signaling under ER stress [12–14]. An elevated level of BiP has been attributed to the ability of cancer cells to resist stress-induced cell death [15–18]. BiP has also been attributed to poor prognosis and drug resistance in cancer [19–21]. Therefore, BiP has become an important target for anticancer drug development [22]. It is therefore highly desirable to pursue new strategies to develop BiP inhibitors.

The human rhomboid family-1 protein (RHBDF1; also known as inactive rhomboid-1, iRhom1) [23, 24], the expression of which is significantly increased in cancer tissues compared to tumor-adjacent tissues [23, 25, 26], is an ER-resident membrane protein [27]. It has been shown to be critically involved in breast cancer progression, poor prognosis, and resistance to chemotherapy [28–33]. Additionally, RHBDF1 can facilitate the epithelial-to-mesenchymal transition (EMT) and cell proliferation in colorectal cancer [31, 34], thereby acting as an oncogene in cervical cancer [32], mediate G-protein coupled receptor agitator-driven transactivation of EGFR signaling pathways [35], and maintain the stability of hypoxia-inducible factor-1α (HIF-1α) in cancer cells under hypoxic and inflammatory conditions [29]. An increasing amount of evidence indicates that the RHBDF1 protein can directly interact with a variety of proteins, such as RACK (the receptor of activated protein-C kinase-1) [29], JNK (c-Jun N-terminal kinase) [36], Par6a (a component of a protein complex critical to the establishment of epithelial cell apicobasal polarity) [37], and iTAP (iRhom Tail-associated protein) [38], often with a tangible impact on the activities of these proteins. It is thus plausible that RHBDF1, as an ER-resident membrane protein, may function as a molecular chaperone in the modulation of critical cellular processes, including ER stress and the UPR.

We demonstrate in this study that RHBDF1 plays a crucial role in maintaining ER protein homeostasis in breast cancer cells. This effect is achieved at least partly through RHBDF1 binding to the BiP protein, enhancing the protein stability of the latter. In addition, we identified segments of the primary structure of the RHBDF1 protein involved in BiP interaction. We show that a pentapeptide, PE5, which resembles a BiP-binding motif in RHBDF1, can effectively inhibit the ATPase activity of BiP, resulting in greatly increased aggregation of ER proteins and a higher ER stress-mediated apoptosis rate. Our findings not only provide new insights into the molecular mechanism underlying cancer cell responses to ER stress but also highlight the utility of targeting RHBDF1-BiP interactions as a plausible strategy in anticancer drug development.

Materials and methods

Reagents and antibodies

The following reagents were used: DMSO (Sigma, D2438, Stamford, USA), DIOC6(3) (MKBio, MX4009, Shanghai, China), MG132 (Selleckchem, S2619, Houston, USA), Cycloheximide (Sigma, C4859, Stamford, USA), Thapsigargin (GLPBIO, GC11482, Montclair, CA, USA), and Leupeptin hemisulfate (MCE, HY-18234A, Nashville, TN 37212, USA). The following antibodies were used: RHBDF1 (Abcam, ab81342, Cambridge, MA, USA), BiP (Proteintech, 11587-1-AP, Rosemont, IL 60018, USA), ER marker PDI (Proteintech, 11245-1-AP and 66422-1-Ig, Rosemont, IL 60018, USA), ATF4 (Beyotime, AF2560, Shanghai, China), CHOP (CST, 2895, Boston, USA), cleaved Caspase-3 (CST, 9661, Boston, USA), cleaved PARP (CST, 5625, Boston, USA), Ubiquitin (Abcam, 134953, Cambridge, MA, USA), p62 (Proteintech, 66184-1-Ig, Rosemont, IL 60018, USA), LC3A/B (Affinity, AF5402, Cincinnati, OH, USA), LC3B (Affinity, AF4650, Cincinnati, OH, USA), HA (Invitrogen, 26183, California, USA), MYC (Proteintech, 16286-1-AP, Rosemont, IL 60018, USA), and FLAG (Sigma, F7425, Stamford, USA). The anti-MYC, anti-HA, and anti-FLAG magnetic beads were purchased from Bimake (Houston, TX77014, USA). The Streptavidin magnetic beads were purchased from MedchemExpress (MCE, HY-K0208, Nashville, TN 37212, USA).

Animals

The female BALB/c mice (20–22 g, 6–8 weeks old) were purchased from Taconic Bioscience (Shanghai, China). They were housed in the Laboratory Animal Center, Nankai University, Tianjin, China. All animal experimental procedures were approved by the Animal Ethics Committee of Nankai University and strictly followed by the Guidelines for the Care and Use of Laboratory Animals (National Institutes of Health, USA). All animals were placed in pathogen-free conditions with a temperature of 24–25 °C and a relative humidity of 50%–55%, a light/dark cycle for 12 h, and free access to standard rodent feed and tap water.

Cell culture and transfection

Human breast cancer cell lines MCF-7 (RRID: CVCL_0031) and MDA-MB-231 (RRID: CVCL_0062), mouse breast cancer 4T1 cells (RRID: CVCL_0125), and Human embryonic kidney AD293 cells (RRID: CVCL_9804) and 293 T cells (RRID: CVCL_RU09) were purchased from the American Type Culture Collection (ATCC). The MCF-7 cells with the stable knockout of RHBDF1 used in this paper are the cells constructed by our group before [36]. MCF-7 and AD293 cells were grown in Dulbecco’s Modified Eagle’s Medium (DMEM), MDA-MB-231 cells were grown in Roswell Park Memorial Institute 1640 Medium (RPMI-1640), and 4T1 cells were grown in DMEM/F12 with 10% fetal bovine serum. Cells were cultured in cell culture dishes or multi-well plates with the indicated medium supplemented with 10% FBS and penicillin/streptomycin (100 U/mL, 50 mg/mL) at 37 °C in mixed gas consisting of 20% O₂, 5% CO₂, and 75% N₂. Cells were transfected with indicated plasmids using Lipofectamine2000 (Thermo Fisher Scientific, Massachusetts, USA) with reduced serum medium Opti-MEM (Life Technologies, California, USA) following the manufacturer’s instructions.

Construct expression plasmids

Full-length human RHBDF1 gene (NM_022450) and Full-length human HSPA5 gene (NM_005347) were amplified from complementary DNA of MCF-7 cells. Then full-length human RHBDF1 tagged with C-terminal HA was cloned into the pLVX-EF1α-IRES-Puro (LMAI Bio, LM-2015, Shanghai, China) and full-length human BiP was cloned into pcDNA3.1 MYC (RRID: Addgene_176045, Cambridge, MA, USA). The different coding sequences (for 1–411 aa, 411–655 aa, 655–855 aa, 655–828 aa, and 800–855 aa) of RHBDF1 were further amplified from full-length human RHBDF1 and cloned as described above. The 70–125 aa of human ubiquitin was amplified from the complementary DNA of MCF-7 cells and cloned into pLVX-EF1α-IRES-Puro/6 × His.

Construct cell lines with stable knockdown of RHBDF1 or BiP

To construct cell lines with stable knockdown of the indicated gene, the pLKO.1 puro vector (RRID: Addgene_8453, Cambridge, MA, USA) containing the shRNA for indicated mRNA was constructed and then transfected into 293 T cells with psPAX2 (RRID: Addgene_12260, Cambridge, MA, USA) and pMD2.G (RRID: Addgene_12259, Cambridge, MA, USA). After 48 h, the cell culture medium was collected and filtered by a 0.45 μm filter to use as a virus crude solution. The obtained lentivirus was infected into MCF-7 or MDA-MB-231 cells with 10 μg/mL Polybrene, followed by screening infected cells with puromycin. The sequences of shRNAs for RHBDF1 are: forward 5′-CCGGCAGTGACAGCACCCAGAAATGCTCGAGCATTTCTGGGTGCTGTCACTGTTTTTTG-3′ and reverse 5′-AATTCAAAAACCGGCAGTGACAGCACCCAGAAATGCTCGAGCATTTCTGGGTGCTGTCACTG-3′.

Western blotting analysis

Cells were cultured in 6 or 12-well tissue culture plates and lysed in RIPA buffer containing 20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1% Triton X-100, and 1% protease inhibitors (Sigma-Aldrich, P8215, Stamford, USA). Cell lysates were separated by SDS-PAGE and transferred onto PVDF membranes (Millipore, IPFL00010, Massachusetts, USA). Membranes were blocked with 5% nonfat dry milk powder in Tris-buffered saline-Tween 20 buffer containing 20 mM Tris-HCl (pH 7.4), 137 mM NaCl, and 0.1% Tween-20 for 1 h at room temperature, incubated overnight at 4 °C with primary antibodies against the target proteins, further incubated with appropriate horseradish peroxidase (HRP)–conjugated secondary antibody for 1 h, and the protein expression is detected by enhanced ECL luminescence solution. The expression is analyzed by Image J software.

Quantitative RT-PCR

To obtain cDNA, total RNA was extracted using TRIzol (Invitrogen, California, USA) from indicated cells and reverse transcribed using SuperScript III First-Strand Synthesis SuperMix (Invitrogen, 11752-50, California, USA). A 20 µL volume reaction consisted of 1 µL reverse transcription product and 250 nM of each primer, and the thermal cycle program was as follows: denaturing for 15 s at 95 °C, annealing, and extension at 60 °C for 30 s. The relative quantity of samples was calculated according to the comparative −ΔΔCt method and normalized to ActB. The primers for BiP are: forward 5′-GAAAGAAGGTTACCCATGCAGT-3′ and reverse 5′-CAGGCCATAAGCAATAGCAGC-3′.

Immunoprecipitation (IP) and pull-down assays

For immunoprecipitation assays, AD293 cells overexpressing the indicated plasmids were lysed for 15 min on ice with NP40 lysis buffer containing 50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1% NP40, 0.5% deoxycholate, and 1% protease-inhibitor cocktail (Sigma-Aldrich, P8215, Stamford, USA). The lysates were centrifuged at 13,000 r/min at 4 °C for 10 min, and the supernatant was rotated lightly with anti-HA, anti-MYC, or anti-FLAG magnetic beads for 5 h at 4 °C. The beads were washed two times for 20 s at 4 °C with rotation using the Triton X-100 lysis buffer containing 300 mM NaCl. Samples were boiled at 100 °C for 10 min with 1 × SDS-PAGE sample buffer and then loaded.

For endogenous IP analysis, cells were lysed according to the described above and the supernatant was incubated with the indicated primary antibodies overnight at 4 °C and mixed with protein A/G agarose beads (Santa Cruz, sc-2003, Santa Cruz, USA) for 3 h at 4 °C. The beads were washed two times at 4 °C using the same lysis buffer by centrifugation at 2500 × g. The following processes were the same as described above.

For pull-down assay, MCF-7 cells were transiently transfected with pLVX-RHBDF1-HA or control pLVX-HA, and pull-down analysis was performed with anti-HA magnetic beads as described above. The immunoprecipitants were electrophoresed by 10% SDS-PAGE and stained using the Fast Silver Stain Kit (Beyotime Biotechnology, P0017S, Shanghai, China) according to the manufacturer’s instructions. The bands of interest were cut into small pieces, decolorized in 50% acetonitrile with 25 mM ammonium bicarbonate, reduced with dithiothreitol, alkylated with iodoacetamide, and digested with trypsin (working concentration 10 ng/μL) at 37 °C overnight. The digestion solution after trypsin peptide extraction and lyophilization was analyzed by LC-MS/MS. The LC/MS/MS analysis of Pinpoint and Proteome Discoverer1.4 was carried out using Thermo Scientific triple quadruplex ion trap and Orbitrap fusion technology.

For biotin-pull-down analysis, the MCF-7 cells were lysed and centrifuged at 13,000 r/min at 4 °C for 10 min, and the supernatants were rotated lightly with streptavidin beads and biotin-PE5 (100 µM) or biotin-mPE5 for 5 h at 4 °C and then the beads were washed, boiled, and loaded as the described above.

For determination of the effect of PE5 on the RHBDF1-BiP interaction, MCF-7 cells were co-transfected with R1-HA and BiP-MYC, and the lysates were centrifuged at 13,000 r/min at 4 °C for 10 min, and then the supernatants were rotated lightly with anti-HA beads and PE5 (100 µM) at 4 °C for 5 h and the following processes were same as the described above.

Immunofluorescent staining

Cells were attached to the circular glass sheet and the frozen tumor tissue was sectioned into 6–8 mm sections, then were fixed with 4% paraformaldehyde, followed by permeabilization with 0.3% Triton X-100. Then samples were blocked with 5% bovine serum albumin and probed with the primary antibodies overnight at 4 °C. The following samples were washed and incubated with secondary antibodies for 1 h at room temperature, then stained with DAPI (Invitrogen, 33342, California, USA). Samples were sealed by an antifading mounting medium and imaged using LSM 800.

To stain ER using DIOC6(3), the cells were attached to the circular glass sheet. After overnight, the cells were probed with DIOC6(3) (5 μM) for 5 min at room temperature, and then washed and stained with DAPI, followed by imaging using LSM 800.

For the immunofluorescence assay of PE5, MCF-7 cells were attached to a 24-well cell culture dish with a glass slide and cultured at 37 °C in 5% CO₂ for 6–12 h. Then 100 μM Mca-PE5 was added into the medium and cultured for 1 h and then the following processes were the same as described above.

For the immunofluorescence assay of mouse breast cancer tissue, the frozen tumor tissue was sectioned into 6–8 mm sections and then the following processes were the same as described above.

Transmission electron microscopic assay

The cell pellets were collected and fixed with 2.5% glutaraldehyde (Sigma-Aldrich, G5882, Stamford, USA). The samples were washed three times using 100 mM sodium phosphate buffer and were post-fixed in 1% osmium tetroxide for 1 h. After blocking with 1% uranyl acetate, the samples were dyed for 1 h and dehydrated through a graded series of alcohol to 100% and embedded in epoxy resin. The embedded samples were cut using Leica ultramicrotome EM UC6 (Leica). Later, the samples were stained with uranyl acetate and lead citrate in a Leica EM Stainer and imaged under a Spirit Transmission Electron Microscope (FEI Company) operating at 120 kV.

Aggresomal assay

To detect intracellular aggresome, the cells were collected and then stained using PROTEOSTAT® Aggresome detection kit (ENZO, ENZ-51035-K100, New York, NY 10022, USA) according to the manufacturer’s instructions and quantified via flow cytometry using a BD FACSCanto II cytometer (BD Biosciences, San Jose, CA, USA).

Molecular docking

To create the functional peptide segment engaging with the ATP-binding domain of BiP, we performed the molecular docking analysis using Schrodinger 2015 suite (Schrodinger, version 3.1, LLC, New York, NY, 2015). Five peptide candidates surrounding Y713/R714 (PV6, YV5, PE5, PA4, YE4) (Supplementary Fig. S2-1) were selected for molecular docking with BiP’s N-terminal ATP-binding domain (PDB: 3IUC). Since peptide binding sites in the N-terminal ATP-binding domain of BiP have not yet been published, we found five potential peptide binding sites in the N-terminal ATP-binding domain of BiP using the Schrodinger 2015 suite (Supplementary Fig. S2-2), and then used one of them as a ligand binding pocket of five peptides for molecular docking (Supplementary Fig. S2-2, Site 1). The structure of the human BiP protein was retrieved from the Uniprot database (https://www.uniprot.org) and downloaded from the RCSB database (http://www1.rcsb.org) in PDB format. The 2D to 3D structural transformation and conformational optimization of BiP protein were performed using Schrodinger’s LigPrep module, and the Protein Preparation Wizard module was used for protein hydrogenation, hydrogen bonding, removal of water molecules, and energy minimization under OPLS 2005 field. Then the Receptor Grid Generation module is used to generate the docking grid. Finally, the molecular docking process of peptides and proteins is completed with the Ligand Docking panel of the Glide module. The peptide with the highest Glide GScore was selected for further SPR experiments.

Purification of recombinant BiP protein

The human BiP (full-length) DNA was inserted in a pET28a vector (Yeasen, 11905ES03, Shanghai, China) with N-terminal 6 × His-tag and expressed in Escherichia coli BL21 (DE3) cells (Agilent). The strain was cultured in Luria-Bertani (LB) medium at 37 °C for 3 h. When OD_{600 nm} was 0.6, 1 mM gratuitous inducer IPTG (MCE, HY-15921, Nashville, TN 37212, USA) was mixed with medium and cultured continuously at 16 °C for 20 h. The bacterial cells were centrifuged at 9000 r/min for 15 min and the supernatant was lysed by sonication. The proteins were purified in the buffer containing 50 mM HEPES, pH 8.0, and 400 mM NaCl using Ni-NTA agarose (Qiagen, 30230, Shanghai, China). The agarose was washed with 20 mM imidazole buffer to remove the miscellaneous proteins. The BiP protein was eluted in 250 mM imidazole buffer and incubated with PreScission protease overnight at 4 °C to remove 6 × His-tag. To further purify the protein, anion exchange was performed using a HiTrap Q HP column (GE Healthcare) and size exclusion chromatography was performed using a HiLoad 16/60 Superdex 200 column. The protein buffer was exchanged into buffer B (50 mM HEPES, pH 8.0, 50 mM NaCl, 10 mM MgCl₂, 10 mM KCl) for the experiments.

Surface plasmon resonance (SPR) assay

To evaluate the interaction between PE5 and BiP, surface plasmon resonance (SPR) was performed using a BIAcore T200 (Uppsala, Sweden). The recombinant BiP protein (1 nM) as the ligand was immobilized onto the surface of carboxylate glucans in the HTG sensor chip and PBS was used as the running buffer for PE5 or mPE5 and PBS-P with 5% DMSO was used as the running buffer for HA15. Each affix sample was prepared and injected onto the chip surface at seven gradient concentrations ranging from 156 nM to 5000 nM to record the sample binding to the surface. The experimental temperature was 25 °C and the flow rate was 20 μL/min. According to the 1:1 Langmuir binding model, the affinity constants (K_d) were calculated by BIAcore T200 software 3.0.2.

ATPase assay

ATP-to-ADP conversion was measured by the ADP-Glo^TM Kinase Assay Kit (Promega, V6930, Madison, WI 53711-5399, USA). A reaction mixture containing 40 μM recombinant full-length BiP protein and a standard ATPase assay buffer with increased concentrations of PE5, mPE5, or HA15 was prepared in 384 Wells of white OptiPlate (Perkin Elmer, Massachusetts, USA) and then was preincubated for 30 min at 37 °C. Then 1 mM ATP was added to the reaction and incubated for 2 h. Luminescence was read on a Multifunctional microporous plate detector (TECAN, 03030923, Austria).

Annexin V–FITC apoptosis assay

The cells (1 × 10⁵~ 2 × 10⁵/well) were attached in 6-well plates and cultured for 6~12 h at 37 °C in 5% CO₂. PE5 or mPE5 was added to the medium at the indicated concentrations and continuously cultured for 48 h. After washing with cold PBS and resuspending, the cells were stained with Annexin V-FITC and propidium iodide (PI) using Annexin V–FITC Apoptosis Detection Kit (Elabscience, E-CK-A211, Wuhan, China) according to the manufacturer’s protocol. The fluorescence was measured by flow cytometry using a BD FACSCanto II cytometer (BD Biosciences, San Jose, CA, USA).

Xenograft study

1.0 × 10⁶ 4T1 cells were suspended in 100 mL PBS and then were injected subcutaneously into the right-side chest of 8-week-old female BALB/c mice (Taconic Bioscience). When the average tumor size reached 60 mm³, the mice were randomly divided into 5 mice in each group. The control group or the treatment group was injected subcutaneously with PBS or PE5 in the pericancerous epithelium, once every 2 days. The tumor size and volume were measured by calipers once every two days and calculated by the formula: 0.5 × D × d², where D and d were the longest and shortest vertical diameters, respectively. When the tumor size in the control group reached 1000 mm³, the experiment was concluded and the mice were euthanized and tumors were collected and frozen for histology.

Quantification and statistical analysis

All data were determined from at least three independent experiments and presented as mean ± SD. The unpaired two-tailed Student t-test was used to analyze the statistical difference between the two sets of data. One-way ANOVA was used for statistical analysis of the difference between multiple groups. *P-values < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 were considered statistically significant.

Results

Artificially depleting the RHBDF1 gene in breast cancer cells results in ER stress and autophagy

We examined the impact of RHBDF1 gene knockout (R1KO) on ER morphology in MCF-7 breast cancer cells. In contrast to those in mock-transfected cells (MT), ER structures in the R1KO cells showed considerably expanded lumens. Similar changes were observed when we treated MDA-MB-231 breast cancer cells with RHBDF1-specific shRNA (shR1) (Fig. 1a). Transmission electron microscopy analysis of these cells revealed diminished ER integrity and ER cisterna dilation, which are typical of ER stress, in RHBDF1-deficient cells (Fig. 1b). By using ubiquitin and p62 as indicators of protein aggresome formation [39], we found a large amount of ubiquitinated proteins and protein aggresomes in proximity to the ER in RHBDF1-deficient cells (Fig. 1c, d). By using an aggresomal detection test, we found that the level of protein aggregation in RHBDF1-deficient cells was approximately 1.7–2.5 fold greater than that in the control groups (Fig. 1e). Electron microscopy images of the cells revealed the accumulation of autophagic lysosomes in RHBDF1-deficient cells (Fig. 1f), consistent with the occurrence of cell autophagy accompanied by UPR failure. Furthermore, Western blotting analyses revealed the presence of the typical autophagy markers LC3-I, LC3-II and p62 in RHBDF1-deficient cells (Fig. 1g). Furthermore, fluorescence staining of sections of experimental tumors formed by subcutaneously implanted mouse breast cancer 4T1 cells showed a 26.4-fold increase in the level of the autophagy marker LC3B in the R1KO group compared to that in the MT controls (Fig. 1h). These data demonstrate ER protein aggregation and ER stress in breast cancer cells in the absence of RHBDF1 gene expression.

Fig. 1. Loss of RHBDF1 causes ER protein aggregation in breast cancer cells.

a Immunofluorescence staining of the ER membrane in breast cancer cells. MCF-7 cells (MT or R1KO) and MDA-MB-231 cells (shCtrl or shR1) were imaged by ER probe DIOC6(3) (5 μM) for 5 min. Shown were representative images from three experiments, >50 cells per experiment. b Ultrastructural analysis by transmission electron microscopy (TEM) in breast cancer cells. N: nucleus, ER: endoplasmic reticulum, CT: cisternae. c, d Immunofluorescence staining of ubiquitin/p62 and ER probe DIOC6(3) in RHBDF1 knockout MCF-7 cells or control cells. e Protein aggregates analysis using PROTEOSTAT Aggresome Detection Kit and FACS. f Ultrastructural images of autolysosomes in breast cancer cells. N: nucleus, AL: autolysosome, ER: endoplasmic reticulum. g Western blot analysis of autophagy markers. h Immunofluorescence staining of mouse 4T1 breast tumor sections 26 days post cancer cell implantation for the autophagy marker LC3B. Shown were representative images (left) and data summary (right) from 5 mice. Data were presented as the mean ± SEM, n = 5, **P < 0.01.

RHBDF1 is pivotal to the maintenance of BiP protein stability in breast cancer cells

Since the BiP protein is recognized as a major executor of the UPR in the ER lumen [14, 40], we measured the impact of RHBDF1 depletion on BiP protein stability. By using Western blotting, we discovered that the levels of the BiP protein decreased by 52% and 30%, respectively, in R1KO and shR1-treated cells compared to those in control cells (Fig. 2a, Supplementary Fig. S1-1). Immunofluorescence staining of BiP confirmed a reduction in the amount of BiP by 60%–72% in RHBDF1-deficient cells (Fig. 2b, Supplementary Fig. S1-2). When the cells were further treated with thapsigargin (TG), an ER stress inducer, the BiP protein level progressively increased in the control cells; however, this increase was significantly slower in R1KO cells (Fig. 2c). We noticed that even though RHBDF1 gene knockout exerted no effect on BiP gene transcription, which we discerned by using RT-qPCR (Supplementary Fig. S1-3), the stability of the BiP protein was markedly changed in R1KO cells, with the half-life of the BiP protein decreased by as much as 61% in R1KO cells compared to control cells when the cells were cultured in the presence of the protein synthesis inhibitor cycloheximide (CHX) (Fig. 2d). To further investigate the mechanisms underlying the decreased BiP level in R1KO cells, we treated the cells either with the lysosome inhibitor leupeptin or with the proteasome inhibitor MG132 for 2 h and found that the BiP level increased by approximately 2-fold in R1KO cells in the presence of MG132 compared to that in vehicle-treated cells; however, there was no change in the presence of leupeptin (Fig. 2e). This is in agreement with previous findings showing that the proteasome pathway is crucial to BiP degradation [41, 42]. Moreover, the BiP ubiquitination rate was markedly increased in the R1KO cells in the presence of MG132 (Fig. 2f), indicating that RHBDF1 depletion results in proteasomal degradation of BiP. Furthermore, we artificially overexpressed RHBDF1 tagged with C-terminal HA (R1-HA) in R1KO cells and then measured BiP protein levels and the extent of BiP ubiquitination. We found that reintroduction of RHBDF1 into R1KO cells led to re-established BiP protein levels (Fig. 2g) and a decreased BiP ubiquitination rate (Fig. 2h). These findings suggest that RHBDF1 function is pivotal to the maintenance of BiP protein stability in breast cancer cells.

Fig. 2. RHBDF1 is critical for maintaining BiP protein stability in breast cancer cells.

a Western blot of BiP in MCF-7 cells (MT or R1KO). Shown were representative images (up) and data summaries (down) from three experiments. b Immunofluorescence staining for BiP and ER marker PDI in MCF-7 cells. Shown were representative images (left) and data summary (right) from three experiments, >50 cells per experiment. c, d BiP levels in MCF-7 cells (MT or R1KO) treated with Thapsigargin (TG, 10 μM) (c) or CHX (10 μg/mL) (d) according to the indicated times. Shown were representative images (up) and data summaries (down) from three experiments. e BiP levels in MCF-7 cells (MT or R1KO) treated with MG132 (20 μM) and Leupeptin (400 nM) for 2 h. Shown were representative images (up) and data summaries (down) from three experiments. f Ubiquitinated BiP in MCF-7 cells (MT or R1KO) pretreated with MG132 (10 μM) for 12 h before collection. g Western blot of BiP in MCF-7/R1KO cells transfected with RHBDF1-HA. h Ubiquitinated BiP in R1KO MCF-7 cells overexpressed with RHBDF1-HA. Shown were representative images from three experiments. All data were presented as the mean ± SEM, n = 3, *P < 0.05, **P < 0.01, ***P < 0.001.

The multitransmembrane domain (MTD) of RHBDF1 interacts with BiP

To determine whether RHBDF1 directly interacts with BiP, we overexpressed HA-tagged RHBDF1 (R1-HA) in MCF-7 cells, carried out immunoprecipitation (IP) assay and identified putative binding proteins via mass spectrometry. We found that BiP was a major component of RHBDF1-binding proteins under the experimental conditions (Fig. 3a). The results from additional IP experiments indicated that RHBDF1 directly interacts with either endogenous BiP or artificially overexpressed BiP (MYC tagged) (Fig. 3b-d). Immunofluorescence analysis with anti-RHBDF1 and anti-BiP antibodies also indicated the colocalization of RHBDF1 and BiP (Fig. 3e). Since the tertiary structure of the RHBDF1 protein remains undetermined [26, 43–45], we dissected the primary sequence of RHBDF1 into three fragments, namely, the N-terminal domain (NTD, 1–411 aa), the first loop domain (FLD, 411–655 aa), and the remaining multitransmembrane and C-terminal domain (MTCTD, 655–855 aa) (Fig. 3f). Based on a topological study of human RHBDF1 [38], we expressed these fragments in AD293 cells and then carried out co-IP experiments with BiP. We found that only the MTCTD fragment coimmunoprecipitated with BiP (Fig. 3g, h). We then divided MTCTD into two fragments, MTD (655–828 aa) and CTD (800–855 aa) (Fig. 3f), repeated the IP experiments, and found that MTD, but not CTD, coimmunoprecipitated with BiP (Fig. 3i). It is thus plausible that RHBDF1 interacts with the BiP protein through the MTD segment.

Fig. 3. RHBDF1 directly interacts with the N-terminal domain of BiP at its multitransmembrane domain.

a Pull-down analysis with anti-HA magnetic beads in MCF-7 cells transfected with pLVX-RHBDF1-HA or control pLVX-HA. b Endogenous immunoprecipitation analysis with anti-RHBDF1 antibodies in MCF-7 cells. c, d Immunoprecipitation analysis with anti-HA (c) or anti-MYC magnetic beads (d) in AD293 cells co-transfected with RHBDF1-HA and BiP-MYC. e Immunofluorescence staining of RHBDF1 (red) and BiP (green) in MCF-7 cells. Colocalized RHBDF1 and BiP proteins show up in yellow. f Schematic diagram of different lengths of RHBDF1 constructed for immunoprecipitation. g, h The immunoprecipitation with anti-HA magnetic beads (g) and anti-MYC magnetic beads (h) in AD293 cells co-transfected with NTD (1-411)-HA, FLD (411–655)-HA, or MTCTD (655–855)-HA and BiP-MYC. i The immunoprecipitation with anti-MYC magnetic beads in AD293 cells co-transfected with MTD (655–828)-HA or CTD (800–855)-HA and BiP-MYC. j Schematic diagrams of RHBDF1 mutant sites constructed to determine the binding site with BiP. k, l The immunoprecipitation with anti-HA magnetic beads (k) and anti-MYC magnetic beads (l) in AD293 cells co-transfected with L3M-HA or L5M-HA and BiP-MYC. m Schematic diagram of two truncations of BiP constructed for immunoprecipitation. n The immunoprecipitation with anti-MYC magnetic beads in AD293 cells co-transfected with RHBDF1 mutants and two truncations of BiP. Shown were representative images from three experiments.

Given that BiP is present in the ER lumen [13] and that the RHBDF1 MTD fragment is linked by two loops (third and fifth loops) on the lumen side of the ER membrane, we carried out site-specific mutations of RHBDF1 in the two lumen side loops, designated L3M (Y713A/R714A) and L5M (D₇₆₈NFAH₇₇₂ to AAADA) (Fig. 3j). We overexpressed the mutant RHBDF1 individually with BiP-MYC in AD293 cells using wild-type RHBDF1 as the control. We performed IP assays and found that L3M-HA, but not L5M-HA, showed less binding affinity for BiP (Fig. 3k, l). Additionally, since the ATP-binding domain (NBD) and substrate-binding domain (SBD) of BiP are located in the N- and C-terminus, respectively [10], we constructed two BiP fragments, namely, BiP-NBD and BiP-SBD (Fig. 3m). We found via co-IP assays that BiP-NBD exhibited significantly higher affinity for RHBDF1 (Fig. 3n). These data are consistent with the view that the third ER luminal loop of RHBDF1, which contains the amino acid residues Y713-R714, and the ATP-binding domain (NBD) of BiP are involved in the RHBDF1–BiP protein interaction.

PE5, a pentapeptide mimicking the BiP binding site of RHBDF1, inhibits the ATPase activity of BiP

Considering the primary structural motifs of RHBDF1 surrounding Y₇₁₃-R₇₁₄, we carried out molecular docking analysis with the N-terminal ATP-binding domain of BiP (PDB: 3IUC) as the target (Supplementary Table S1, Supplementary Fig. S2). We found that the pentapeptide PYRAE (PE5) fit in the ATP-binding domain of BiP (Fig. 4a, b). We then synthesized PE5 and a mutant pentapeptide in which the Tyr and Arg residues were replaced with Ala (PAAAE; mPE5) as the control (Supplementary Fig. S3, Supplementary Table S2). We produced His-tagged BiP recombinant protein (Supplementary Fig. S4-1, S4-2) and carried out a surface plasmon resonance (SPR) assay to determine the interaction between BiP and PE5 or mPE5. We found that PE5, but not mPE5, showed binding affinity (K_d = 57.7 μM) for BiP (Fig. 4c). This binding affinity was similar to that of the known BiP inhibitor HA15 [22, 46] (K_d = 43.3 μM) (Fig. 4c). We then synthesized biotin-PE5 (Supplementary Fig. S4-3, Supplementary Table S2) and biotin-mPE5 (Supplementary Fig. S4-4, Supplementary Table S2) and then carried out biotin pull-down experiments with MCF-7 cell lysates. We found that biotin-PE5, but not biotin-mPE5 or biotin alone, interacted with BiP (Fig. 4d). Additionally, we carried out colocalization experiments with PE5 and BiP in MCF-7 cells by using an anti-BiP antibody and the immunofluorescent peptide Mca-PE5 (Supplementary Fig. S3-4, Supplementary Table S2) and found that Mca-PE5 and BiP colocalized to the ER (Fig. 4e). Moreover, we found by using an immunoprecipitation assay that PE5 disrupted the RHBDF1-BiP interaction (Fig. 4f). Furthermore, by using an ADP kinase assay, we found that PE5 inhibited the ATPase activity of BiP, with an IC₅₀ value of 92.33 μM, which was on the same order as the inhibitory effect of HA15 (60.08 μM) (Fig. 4g). These data indicate that PE5, which mimics the BiP binding site of RHBDF1, can not only interact with BiP but can also inhibit the ATPase activity of the latter.

Fig. 4. PE5 targets BiP to inhibit its ATPase activity.

a PE5-binding site of BiP (PDB: 3IUC) in 3D docking interactions of PE5 and BiP by Schrodinger 2015 suite. b 2D docking interactions plot of PE5 with ATP-binding domain of BiP. c SPR assay for determination of interaction between PE5 and BiP. d The immunoprecipitation with streptavidin beads in MCF-7 cells lysates with biotin, biotin-PE5, or biotin-mPE5 treatment. e The immunofluorescence analysis for Mca-PE5 and BiP in MCF-7 cells. Shown were representative images from three experiments, >50 cells per experiment. f The immunoprecipitation with anti-HA beads in MCF-7 cells lysates in the presence of PE5. The cells were co-transfected with R1-HA and BiP-MYC. g The ATPase activity of BiP with PE5 or mPE5 treatment. IC₅₀ is presented as the mean ± SD of three independent experiments performed in duplicate.

PE5 induces ER protein aggregation and ER stress-mediated apoptosis in breast cancer cells

Since the disruption of BiP activity may result in diminished UPR while enhancing ER protein aggregation and proteotoxic stress, we treated MCF-7 cells with PE5 and analyzed its effect on these changes. By using transmission electron microscopy, we found pulverized ER structures with dilated cisternae, which are characteristics of ER stress, in PE5-treated MCF-7 cells (Fig. 5a). Additionally, we found a PE5 dose-dependent increase in protein aggregates in PE5-treated cells but not in PBS- or mPE5-treated cells (Fig. 5b). Moreover, we measured the levels of ER stress-associated proteins such as ATF4, CHOP, cleaved Caspase-3, and cl-PARP in PE5-treated MCF-7 cells and found that the levels of each of these proteins were noticeably higher in PE5-treated cells than in mPE5-treated controls under otherwise identical experimental conditions (Fig. 5c, d, Supplementary Fig. S5). The experiments were repeated with another breast cancer cell line, the MDA-MB-231 cell line, and similar results were obtained (Fig. 5c, d). The level of the autophagy marker LC3B also increased by 1.6-fold under these experimental conditions (Fig. 5c). Apoptosis assays using Annexin V-FITC demonstrated a dose-dependent increase in the apoptosis rate by 1.9- and 2.7-fold with 100 μM and 200 μM PE5, respectively, compared with the effect of PBS or mPE5 treatment (Fig. 5e). These findings indicate that PE5, which can inhibit the ATPase activity of BiP, caused ER stress and cancer cell apoptosis with considerable potency. Moreover, considering our results showing that RHBDF1 stabilizes the BiP protein through protein-protein interactions and that PE5 disrupts the RHBDF1-BiP interaction (Fig. 4f), we suspected that treatment with PE5 may affect the protein stability of BiP by disrupting the RHBDF1-BiP interaction, resulting in destabilized and degraded BiP proteins; however, the BiP protein level in PE5-treated cells or tumor tissues did not decline significantly (Supplementary Fig. S5 and Fig. 6e). We are leaning toward the conclusion that PE5 treatment of the cells leads to severe ER stress, which might facilitate the production of new BiP proteins.

Fig. 5. PE5 induces ER protein aggregation and ER stress-mediated apoptosis in breast cancer cells.

a Ultrastructural analysis by transmission electron microscopy (TEM) in MCF-7 cells treated with PE5 or mPE5 for 48 h. ER: endoplasmic reticulum, CT: cisternae. b Protein aggregates analysis in MCF-7 cells treated with PE5 or mPE5 for 48 h using PROTEOSTAT Aggresome Detection Kit and FACS. Data from three experiments were presented as the mean ± SEM, n = 3, *P < 0.05, **P < 0.01. c, d Relative protein levels of ER stress markers in MCF-7 or MDA-MB-231 cells treated with PE5 or mPE5 at 50-200 μM (c) or at 4-24 h (d). Shown were representative images (up) and summaries (down) from three experiments. Data from three experiments were presented as the mean ± SEM, n = 3, *P < 0.05, **P < 0.01. e The apoptosis assay in MCF-7 or MDA-MB-231 cells treated with PE5 or mPE5 using Annexin V-FITC apoptosis detection kit. The cells in the viable, early stage, and late stage of apoptosis/necrosis are in the bottom left quadrant, the bottom right quadrant, and the top right quadrant of each panel (Annexin V–negative, PI-negative) (left). The percentage of apoptotic cells is shown in a histogram (right). Shown were the representative images from three experiments.

Fig. 6. PE5 inhibits breast cancer development in mice.

a Schematic diagram of animal experiments. b The tumor volume analysis of mice treated with PBS or PE5. c The tumor weight of mice treated with PBS or PE5. d The body weight of mice treated with PBS or PE5. e Western blotting analysis of ER stress and apoptosis-related proteins in PE5-treated or PBS-treated tumors. f Immunofluorescence analysis for ER stress markers in tumor sections from mice treated with PBS or PE5. Shown were representative images (up) and summaries (down) from 5 mice. Data were presented as the mean ± SEM, n = 5, **P < 0.01, ***P < 0.001.

PE5 inhibits tumor development in mouse models of breast cancer

We determined whether PE5 showed anticancer activity by evaluating its effect on a tumor model of mouse 4T1 breast cancer cells (Fig. 6a); mouse cells instead of human breast cancer cells were used because both the third lumenal loop junction of the murine RHBDF1 protein and the ATP-binding domain of the murine BiP protein are identical to their counterparts in the human proteins (UniProt, Q96CC6 RHDF1_HUMAN and Q6PIX5 RHDF1_MOUSE, P11021 BIP_HUMAN and P20029 BIP_MOUSE). When the tumor-bearing animals were treated with PE5 (10 mg/kg per subcutaneous injection every two days), the growth rates of the tumors were approximately 2 fold lower than those of the vehicle-treated tumors (Fig. 6b), with the average tumor weight of the experimental group being approximately 40% of that of the control group at the end of the 26-day treatment period (Fig. 6c) without inducing apparent toxicity (Fig. 6d). Additionally, by performing Western blotting analysis, we found that ER stress and levels of apoptosis-related proteins, such as ATF4, CHOP, cleaved Caspase-3, and cleaved PARP, were markedly increased in the PE5-treated tumors compared to the PBS-treated tumors (Fig. 6e). Consistent with the aforementioned ER stress-inducing mechanism, the protein levels of ATF4, CHOP, cl-PARP, and LC3B in PE5-treated tumors all increased by approximately 2-fold compared to those in the vehicle-treated control group, as assessed by immunostaining of the tumor sections (Fig. 6f). These findings suggest that PE5 inhibits tumor growth by triggering ER stress and ER stress-mediated apoptosis in breast cancer cells.

Discussion

The structure and functionality of BiP, an important factor in preserving the homeostasis of ER proteins, have been extensively researched [10, 13, 47]. It remains unclear, however, how the protein stability of BiP is maintained. We show in this study that the ER membrane protein RHBDF1 directly interacts with BiP and stabilizes the latter. We and others have shown that RHBDF1 interacts with a variety of proteins and modifies their activities [29, 36–38]. It is thus plausible that RHBDF1 acts as a chaperone on the ER membrane to facilitate correct folding and structural stabilization of BiP. We further demonstrate that a pentapeptide, PE5, which mimics a structural motif of the potential BiP-binding site on the RHBDF1 protein, disrupted the RHBDF1-BiP interaction, inhibited the ATPase activity of BiP, and induced ER stress-mediated apoptosis in breast cancer cells. These findings suggest that RHBDF1 and BiP may exhibit paired functions in the maintenance of ER functions to mediate proper protein folding in breast cancer cells. Our findings indicate that RHBDF1 may function as a molecular chaperone with several active domains that may be utilized in interactions with a multitude of proteins and thus may play a significant spatiotemporal role in the modulation of a variety of biological processes, as shown in the mediation of GPCR ligand-initiated transactivation of EGFR [31], the stabilization of HIF-1α under hypoxic conditions [29], and the disruption of the apicobasal polarity of epithelial cells [37].

Interestingly, no significant difference was found in the degree of protein aggregation between RHBDF1-overexpressing MCF-7 cells and control cells regardless of the TG treatment or in the ATPase activity of proteins pulled down with MYC affinity beads between RHBDF1-HA- and BiP-MYC-overexpressing cells and cells with only BiP-MYC overexpressed (Supplementary Fig. S6). This finding suggests that RHBDF1 binds BiP but does not inhibit its ATPase activity, whereas PE5 binds BiP and inhibits its ATPase activity. We think that this outcome is because RHBDF1 transiently binds BiP, facilitating its folding, which releases it from the complex; however, PE5 binds BiP to consolidate, disrupting the binding of ATP to BiP and resulting in the inhibition of BiP ATPase activity. Moreover, we suspect that PE5 may also affect the protein stability of BiP, as the results showed that PE5 disrupted the RHBDF1-BiP interaction (Fig. 4f) and that the level of BiP was sometimes reduced in the PE5-treated cells compared to control cells, but not significantly (Supplementary Fig. S5 and Fig. 6e). Under ER stress, the level of BiP proteins increased quickly in cells to restore ER homeostasis. HA15, which functions as a BiP ATPase inhibitor but has no effect on its protein stability, targets BiP to trigger ER stress conditions, and the levels of BiP increased markedly in HA15-treated cells [17, 22]. We found that BiP, ATF4, and CHOP levels were increased significantly in HA15-treated cells compared to DMSO-treated cells, whereas ATF4 and CHOP but not BiP levels were increased in PE5-treated cells (Supplementary Fig. S6-4), suggesting that in PE5-treated cells, ER stress was induced, but the total level of BiP was not increased. These findings are consistent with the view that PE5 not only triggers ER stress by directly targeting BiP to inhibit its ATPase activity but also destabilizes BiP by disrupting the RHBDF1-BiP interaction, which is one of the many mechanisms by which BiP stability is regulated.

Our finding showing that the amino acid residues Y₇₁₃-R₇₁₄ of RHBDF1 are involved in the interaction with the ATP-binding domain of BiP providing a potentially important target for the development of anticancer therapeutic agents aimed at disrupting the ability of cancer cells to overcome ER stress, realized by maintaining a properly functional UPR. The pentapeptide PE5 can induce ER stress-mediated apoptosis by specifically targeting the ATPase activity of BiP and disrupting the initial UPR, demonstrating the feasibility of this approach. Signaling proteins such as ATF4, CHOP, cl-Caspase-3, and cl-PARP, which are known to be closely associated with ER stress-initiated cell apoptosis, as well as the autophagy marker LC3B, were highly upregulated in cancer cells treated with PE5, further supporting the validity of targeting the RHBDF1-BiP interaction for cancer drug development.

In summary (Fig. 7), our data are consistent with the view that the stability of BiP is critically mediated by RHBDF1, whereas the disruption of the interaction of the two proteins, such as that achieved by the use of the pentapeptide PE5, leads to BiP protein degradation and the accumulation of unfolded proteins and ER stress. Additionally, it is interesting that the pentapeptide PE5 can not only disrupt the RHBDF1-BiP interaction but also can directly target BiP and inhibit the ATPase activity of BiP, suggesting that PE5 can bind to BiP in such a manner that it causes conformational changes that result in the loss of the ATPase activity of BiP. PE5 may thus provide a scaffold for the development of a new BiP inhibitor. As the role of BiP has been studied in a broad variety of cancer cell types, we anticipate that the mechanism described in this study may be applicable to other malignancies even though our investigation was restricted to breast cancer.

Fig. 7. Schematic representation depicting RHBDF1-BiP interaction in the ER.

RHBDF1 may serve as a molecular chaperone to assist BiP protein folding. Based on the structural motifs of RHBDF1 involved in the interaction with BiP, a pentapeptide PE5 is designed and found to be able to target directly BiP and inhibit the ATPase activity of BiP, giving rise to ER stress-mediated apoptosis in breast cancer cells.

Author contributions

ZSZ and LYL conceived the study and wrote the manuscript with input from other authors. SJR performed most of the biochemical and cellular experiments. HL, XLZ and YYS provided technical assistance. YW, YJZ, and XJQ contribute to data analysis manuscript preparation.

Data availability

All data supporting the findings of this study are available from the corresponding author on reasonable request.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41401-023-01163-x.