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Introduction

Gastrointestinal cancers account for a large portion of cancer-related death worldwide, including Korea. Among them, gastric cancer is still one of the most common leading causes of death, and if it is diagnosed in advanced or metastatic stage, the overall prognosis is still poor even though there has been rapid progress for therapeutic modalities of gastric cancer, including surgery or chemotherapy. Thus, there is a need to find novel therapeutic agents effective against malignant gastric disease, especially for advanced or metastatic cancer.

Plumbagin (5-hydroxy-2-methyl-1,4-naphthoquinone) is a quinonoid constituent extracted from the roots of the medicinal plant Plumbago zeylanica L (1). The roots of Plumbago zeylanica have been used for various treatment aims in Oriental medicine fields. One promising effect of plumbagin is that it shows antitumor potential against various types of cancer. For example, an in vivo study demonstrated that plumbagin significantly inhibited the growth of azoxymethane-induced intestinal tumors in rats (2), and several in vitro studies reported that plumbagin showed anti-carcinogenic effects including cell proliferation or invasion or induced cell cycle arrest or apoptosis in breast cancer (3), melanoma (4), non-small lung cancer (5) or prostate cancer cells (6).

Several pivotal studies have clearly demonstrated the molecular mechanisms of antitumor effects of plumbagin in various types of cancer cells. Hafeez et al reported that plumbagin inhibited constitutive expression of epidermal growth factor receptor (EGFR), phosphorylation and DNA binding activity of signal transducer and activator of transcription 3 (STAT3) and nuclear factor-κB (NF-κB) in pancreas cancer cells (7). Manu et al showed that plumbagin has a potential blocking activity of CXC chemokine receptor 4 (CXCR4) and potential for inhibition of invasion and migration in breast and gastric cancer cells (8). A recent in vitro study investigated the underlying mechanism of plumbagin in gastric cancer cells, and demonstrated that plumbagin inhibited NF-κB p65 nuclear translocation and phosphorylation of p65, IκBα and IκBα kinase (IKKα), and downregulated NF-κB-related gene products, such as inhibitor of apoptosis 1 (IAP1), X-linked inhibitor of apoptosis (XIAP), B-cell lymphoma-2 (Bcl-2), Bcl-xL and vascular endothelial growth factor (VEGF) (9). Another well-designed in vitro study showed that plumbagin suppresses STAT3 activation pathway through induction of SH2-containing protein tyrosine phosphatase 1 (SHP1), a non-receptor type protein tyrosine phosphatase (PTPase), in multiple myeloma cells (10). However, impact of plumbagin on STAT3 signaling pathway in gastric carcinogenesis has not been reported yet.

Previously, in vitro, we observed that SHP1 expression was markedly reduced or negative in various gastric cancer cell lines, which was mainly caused by epigenetic silencing mechanism, and exogenous introduction of SHP1 plasmid significantly downregulated Janus kinase 2 (JAK2)/STAT3 pathway and their target genes (unpublished data). From this background, we aimed in this in vitro study to demonstrate the ability of plumbagin to induce SHP1 expression and suppress JAK2/STAT3 signaling pathway in gastric cancer cells.

Materials and methods

Reagents and cell line

Plumbagin (purity >97%) was purchased from Sigma-Aldrich (St. Louis, MO, USA) and, dissolved in dimethyl sulfoxide (DMSO) to a concentration of 100 mmol/l and stored at −20°C, and diluted to indicated concentration immediately before use. Recombinant human interleukin-6 (IL-6) and broad-acting PTPase inhibitor sodium pervanadate was purchased from Sigma-Aldrich. A rabbit polyclonal IgG antibody against human SHP1 (sc-287) and β-actin (sc-47778) was purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Mouse monoclonal IgG antibodies against human STAT3 (no. 9139) and phospho-STAT3 (Tyr 705, no. 4113), and rabbit polyclonal antibodies against human JAK2 (no. 3230) and phospho-JAK2 (Tyr 1007/1008, no. 3771) were purchased from Cell Signaling Technology, Inc. (Beverly, MA, USA).

The human gastric cancer cell line (MKN-28) was obtained from Korean Cell Line Bank (Seoul National University, Seoul, Korea), and cultured in RPMI supplemented with 10% heat-inactivated FBS and penicillin/streptomycin (1.0%) (all from Gibco, Carlsbad, CA, USA). Cells were incubated in a humidified atmosphere of 5% CO2 at 37°C.

Western blot analysis

Total 80–100 μg of cytoplasmic proteins were extracted using CelLytic M (C2978; Sigma-Aldrich) with Complete Mini (pretease inhibitor cocktail; Roche Diagnostics GmbH, Mannheim, Germany). Primary antibodies were diluted at 1:1,000 in the blocking buffer (Tris-buffered saline with Tween-20; Biosesang, Gyeonggi, Korea) containing 5% non-fat skim milk (Difco; Becton-Dickinson and Co., Sparks, MD, USA). Probed membranes were incubated for 12 h at 4°C. The membranes were incubated with goat anti-mouse or anti-rabbit IgG as a secondary antibody for 1 h at room temperature. The protein bands were detected by exposing membrane to enhanced chemiluminescence (Perkin-Elmer, Wathman, MA, USA) for 1 min.

Reverse transcription-polymerase chain reaction (RT-PCR)

Total RNA was extracted from whole cells using TRIzol (Invitrogen Life Technologies, Carlsbad, CA, USA) method, and subsequently complementary DNA (cDNA) was produced by using High Capacity cDNA Reverse Transcription kit (Applied Biosystems, Foster City, CA, USA) and treatment with 1 unit of DNase (Promega Corp., Fitchburg, WI, USA). We performed RT-PCR by modifying a previously described method. In brief, 20 ng of prepared cDNA was used to make 25 μl of PCR product using EconoTaq® Plus Green Master Mix (Lucigen Corp., Middleton, WI, USA). PCR was done under the following conditions: initial denaturation at 94°C (2 min), followed by 30–40 cycles of denaturation at 94°C (15 sec), annealing at 55°C (15 sec), extension at 72°C (15 sec) and final extension at 72°C (10 min). The oligonucleotide sequences are summarized in Table I. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as a housekeeping gene for each sample. PCR products (5 μl) were loaded on a 2% agarose gel, and positive bands were obtained by staining with ethidium bromide (Amresco LLC, Solon, OH, USA).

Table I

Characteristics of primers for RT-PCR.

Table I

Characteristics of primers for RT-PCR.

GenePrimers (5′→3′)GenBank accession no.Size (bp)Reference
SHP1F : ACTGAGCTGCATCTGAG
R : CCACTGACGAGAGC
NM_080549.2240(31)
Cyclin D1F : TTCGATGATTGGAATAGC
R : TGTGAGCTGCTCATTGAG
XM_006718653.1150(44)
VEGF-1F : GGAGTGTGTGCACGAGAGTCAC
R : GGTCGACTGAGAGCT
NM_001287044.1343(45)
SurvivinF : TTCTGCACATCTGAGTCG
R : TGTCGAGAGCTCAGT
NM_001012271.1391(46)
MMP-9F : TTGACAGCGACAGAGTG
R : GCATTCACGTCGTCCTTAT
NM_004994.2179(47)
GAPDHF : GGTCTCTCTGACTCACA
R : AGCATTCGTTGTCATAC
NM_002046116(48)
Water soluble tetrazolium salt-1 (WST-1) cell proliferation assay

To quantify the inhibitory effect of plumbagin on cellular proliferation, we used a commercial WST-1 assay kit (EZ-Cytox; DoGen, Seoul, Korea) as manufacturer’s instructions (11). Briefly, 1×104 MKN-28 cells/well were cultured in a 96-well plate at 37°C for 24 h, and treated with plumbagin at 20 or 40 μM for 3, 6 and 9 h. We also cultured untreated cells for the same time period as a control. After treatment, 10 μl of WST was added in each well for 4 h, and absorbance at 450 nm was measured by an ELISA reader (Epoch; BioTek Instruments, Inc., Seoul, Korea). All the experiments were performed in triplicate.

Wound healing assay

After treated with plumbagin at 20 or 40 μM for 6 h, cells were equally seeded on a 6-well plate chamber, and after attachment, a monolayer wound was made using 200 μl pipette tip. The media were changed to remove floating debris, and the vertical distance between the sides of the wound was measured at 24 and 48 h after wound injury using software (12). All the experiments were performed in triplicate.

Matrigel invasion assay

Following treatment with plumbagin at 20 or 40 μM for 6 h, 4×104 cells/well were placed into the 24-well Matrigel Invasion Chambers (BD Biosciences, Franklin Lakes, NJ, USA) in 2% FBS medium, and in the lower wells 10% FBS was added. After 24 h of incubation, filter membranes were stained with crystal violet, and the number of positive invading cells which penetrated through membrane pore was counted under ×20 magnification in at least five randomly selected separate areas.

Annexin V assay

To compare the different percentage of apoptotic cells by plumbagin treatment, we used an Annexin V-FITC assay kit (Anse Technologies Co., Ltd., Seoul, Korea) according to the manufacturer’s instructions. Briefly, 1×106 MKN-28 cells/well were treated with 20 or 40 μM of plumbagin for 6 h, washed and resuspended in binding buffer, followed by staining with an Annexin V-FITC and propidium iodide (PI) solution for 10 and 30 min, respectively. After staining, samples were analyzed using FACSCalibur Flow Cytometer (Becton-Dickinson and Co., San Jose, CA, USA).

Statistical analysis

The SPSS ver. 19.0 (SPSS, Inc., Chicago, IL, USA) was used for all analyses. Continuous data are presented as mean ± standard deviation. A Student’s t-test was performed for continuous data, and p<0.05 was considered as statistically significant.

Results

Plumbagin inhibits phosphorylation of constitutive STAT3 in MKN-28 cells

First, we investigated the effect of plumbagin to modulate the activity of JAK2/STAT3 signaling in the gastric cancer cells. MKN-28 cells were treated with different concentration (10, 20 and 40 μM) of plumbagin for 6 h, and western blot analysis was performed to measure phosphorylation level of JAK2 at tyrosine 1007/1008 and STAT3 at tyrosine 705. Plumbagin significantly inhibited phosphorylation of JAK2 and STAT3 from 20 μM, and in contrast, total JAK2 and STAT3 showed similar level regardless of plumbagin treatment, which supports that plumbagin negatively modulates JAK2/STAT3 activity mainly via dephosphorylation rather than protein degradation. Because SHP1 is one of non-transmembrane PTPase which negatively modulate JAK2/STAT3 signaling in epithelial cells (13), we also observed induction of SHP1 expression by plumbagin treatment starting at 20 μM (Fig. 1A). Our data suggest that plumbagin might dephosphorylate and downregulate JAK2/STAT3 activity by induction of SHP1 expression in MKN-28 cells. We also observed the time-course to inhibit phosphorylation of JAK2/STAT3 and to induce SHP1 by treatment with 40 μM of plumbagin for indicated time points. Phosphorylation of JAK2/STAT3 was ameliorated at 6 h, whereas SHP1 expression appeared at 3 h, which suggests that a time-lag might exist between induction of SHP1 and downregulation of JAK2/STAT3 activity in MKN-28 cells (Fig. 1B).

Plumbagin inhibits IL-6-induced STAT3 phosphorylation in MKN-28 cells

Previous in vitro studies demonstrated that stimulation with human recombinant IL-6 upregulates phosphorylated STAT3 level to promote invasive activity in gastric cancer cells (14,15). Thus, we investigated whether plumbagin could modulate IL-6-induced phosphorylation of STAT3 in gastric cancer cells. MKN-28 cells were stimulated with 50 ng/ml of IL-6 at indicated time points (30 and 60 min), and we observed that phosphorylated STAT3 was significantly upregulated from 30 min, whereas SHP1 expression continued to be weakly positive. However, treatment with 20 and 40 μM of plumbagin for 6 h, phosphorylated STAT3 ameliorated even in 60 min stimulation with 50 ng/ml of IL-6, and SHP1 expression was restored during stimulation (Fig. 2). These findings suggest that plumbagin can also suppress IL-6-induced STAT3 phosphorylation as well as constitutive STAT3 phosphorylation, and SHP1 might play a critical role in this process.

Inhibition of SHP1 restores phosphorylation of JAK2 and STAT3 in MKN-28 cells

Because we observed that SHP1 was implicated in the dephosphorylation and inactivation of JAK2/STAT3 signaling, for the next step, we validated this mechanism by using PTPase inhibitor sodium pervanadate (10,16,17). Treatment with indicated concentration of pervanadate and 40 μM of plumbagin for 6 h, and western blot analysis showed that plumbagin downregulated phosphorylation of JAK2 and STAT3, this effect was restored by adding 25 and 50 μM of pervanadate, whereas SHP1 expression showed the opposite pattern, it was restored by treatment with plumbagin but ameliorated by combination with pervanadate (Fig. 3). Taken together, these findings also support our suggestion that SHP1 might be closely related to the mechanism of plumbagin-induced inhibition of JAK2/STAT3 activity in MKN-28 cells.

Plumbagin downregulates STAT3-associated target genes in MKN-28 cells

In various human malignancies including gastric cancer, STAT3 is commonly activated and acts as a pivotal transcription factor to upregulate multiple target genes involving proliferation, invasion/metastasis and anti-apoptosis, such as VEGF-1, matrix metalloproteinase-9 (MMP-9), Bcl-xL, survivin or cyclin D1 (18). To investigate the effect of plumbagin in regulating target gene expression related to STAT3 pathway in gastric cancer cells, treatment with 20 or 40 μM of plumbagin was performed for 6 h with or without stimulation of IL-6, and by RT-PCR. Plumbagin restored SHP1 expression in both constitutive and IL-6-stimulated conditions, and reduced gene expression of VEGF-1, MMP-9, Bcl-xL, survivin and cyclin D1, which were maximally reduced by 40 μM concentration (Fig. 4). These findings suggest that plumbagin modulates mRNA expression of STAT3-related target genes via restoration of SHP1 expression in MKN-28 cells.

Plumbagin inhibits cell proliferation, migration and invasion, and induces apoptosis in MKN-28 cells

To determine functional effects of plumbagin for STAT3-related cellular proliferation, migration, invasion and apoptosis in gastric cancer cells, MKN-28 cells were treated with 20 or 40 μM of plumbagin. By performing WST-1 cell proliferation assay, we observed that plumbagin significantly inhibited cell proliferation in a time- and dose-dependent manner, with maximal inhibitory effect at 40 μM in 6 h treatment (Fig. 5). After treatment with plumbagin for 6 h, we made wound injury in a 6-well plate using a pipette tip, and we observed that plumbagin significantly inhibited wound closure at 24 and 48 h after injury and this inhibitory effect was more prominent at 40 than 20 μM of plumbagin (Fig. 6). We treated MKN-28 cells with 20 or 40 μM of plumbagin for 6 h, and cultured for 24 h in Transwell plate with a pore membrane, and then fixed and stained the cells by crystal violet. We also observed that plumbagin significantly reduced the relative number of invading cells, and 40 μM of plumbagin was more effective than 20 μM (Fig. 7). Finally, we investigated pro-apoptotic effect of plumbagin by Annexin V assay, and relative number of apoptotic cells was significantly increased by 40 μM of plumbagin, rather than 20 μM (Fig. 8). Taken together, these findings suggest that plumbagin significantly inhibits cell proliferation, migration and invasion, and induces cellular apoptosis in MKN-28 cells, and these functional effects are dose-dependent.

Discussion

In this in vitro study, we showed that plumbagin restored SHP1 expression to downregulated JAK2/STAT3 activity and their target genes, and consequentially, led to anti-proliferative, migratory, invasive and pro-apoptotic effects in gastric carcinoma cells. To our knowledge, our study firstly demonstrates that plumbagin inhibits STAT3 pathway through induction of SHP1 activity in stomach cancer cells. Little has been reported about anti-cancer role of plumbagin in gastric cancer, and only few studies focused on its inhibitory effects on NF-κB or CXCR4 pathway (8,9), or cytoxic effect through generation of reactive oxygen species (ROS) (19). Also, recent studies reported several candidate molecules to downregulate JAK2/STAT3 activity and exhibit antitumor effect in gastric cancer cells (20–23). However, none of them showed molecular link between SHP1 and JAK2/STAT3 pathway in gastric cancer cells.

SHP1 is non-receptor-type PTPase, which is encoded by PTPN6 gene located on human chromosome 12p13 (24), and it has been reported as a negative regulator of JAK2/STAT3 activity by dephosphorylation of JAK2 and STAT3 to act as a PTPase (25,26). Previous studies demonstrated that SHP1 is inactivated by aberrant methylation of CpG island promoter in various hematopoietic malignancies, and their functional roles have been extensively investigated in hematopoietic cancer cells (27–30). However, only few studies reported CpG island promoter hypermethylation in epithelial cells such as colon cancer cells (13,31), and little is known about reduced gene expression or promoter hypermethylation of SHP1 in gastric cancer cells except that several studies briefly reported the methylation rate of CpG island promoter of SHP1 in gastric carcinoma tissues (32,33). In colon cancer cells, SHP1 expression was mainly regulated by DNA methylation and upregulated by DNA methyltransferase inhibitors such as 5-aza-2′-deoxycytidine (5-aza-dC). Furthermore, increased SHP1 expression by transfection with SHP1 plasmid vector or treatment with demethylating agent such as 5-aza-dC substantially decreased p-JAK2/p-STAT3 level (13). We observed previously that SHP1 expression was epigenetically regulated and closely related with STAT3 activity in gastric cancer cells (unpublished data). Thus, as the next step, we searched for a candidate molecule which can induce SHP1 expression in gastric cancer cells, and demonstrated that plumbagin might be a potential inducer of SHP1 to inhibit JAK2/STAT3 pathway.

Previous pivotal studies extensively investigated the crucial role of STAT3 for initiation and progression of gastric cancer. Persistent infection of CagA-positive Helicobacter pylori (H. pylori) strain activates constitutive STAT3 via chronic JAK2 activity, which in turn promotes target gene transcription associated with proliferation, invasion, metastasis and angiogenesis. In terms of gastric epithelial cells, IL-6 family ligands such as IL-6 and IL-11 is associated with chronic inflammation by CagA-positive H. pylori and development of gastric cancer (18,34). The suppressors of cytokine signaling (SOCS) family proteins such as SOCS-1 or -3 have been reported as important regulators of JAK2/STAT3 pathway by negative feedback mechanism (35,36). Numerous PTPases have been presented as promising targets to inhibit STAT3 signaling in various kinds of cancer, including SHP1 (37), SHP2 (38), PTP-1D (39) and PTEN (40). However, little is known about the expression level and roles of PTPase in gastric tumorigenesis, and their inhibitory action on STAT3 signaling is still controversial. Several previous studies have focused on the effect of SHP2, and an in vitro study demonstrated that phosphorylated CagA preferentially activates SHP2/extracellular signal-regulated kinase (ERK) pathway and induces cell growth inhibition (41), whereas immunohistochemical studies using human stomach tissues showed that SHP2 expression was significantly enhanced in H. pylori-infected gastric cancer (42,43). Previously, we observed that exogenous expression of SHP1 in gastric cancer cells significantly inhibited cellular proliferation, migration and invasion (unpublished data), however, this phenomenon should be further validated in immunohistochemical studies using human gastric carcinoma tissues. This study might further support the hypothesis that SHP1 negatively regulates STAT3 activity because induction of SHP1 by plumbagin inhibited JAK2/STAT3 signaling and inhibition of SHP1 reversed the plumbagin effects on JAK2/STAT3 pathway.

In conclusion, our study suggests that plumbagin might have promising anti-cancer potential via upregulation of SHP1 expression and inhibition of JAK2/STAT3 pathway in gastric cancer cells, and SHP1 might be an alternative target to regulate STAT3 activity in gastric carcinogenesis. Further preclinical studies concerning the effects of plumbagin in STAT3 overexpressing gastric cancer should be performed, and also other promising agents which can upregulate SHP1 expression in gastric cancer cells need to be investigated.

Acknowledgements

This research was financially supported by grants from Korea University (Seoul, Korea) (grant no. R1211041) and Pacific Pharma Corp. (Seoul, Korea) (grant no. Q1307141).

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Introduction

Breast cancer is the second leading cause of cancer related death among women worldwide. It was estimated that there will be 230,480 new cases of invasive breast cancer and 39,520 new deaths among US women in 2011 (1). Among the molecular targets for treatment purpose, epidermal growth factor receptor (EGFR) and human epidermal growth factor receptor 2 (HER2) were most widely investigated as potential targets for anticancer drug development since the first report of the overexpressions in human breast cancers indicates a poor prognosis (2,3). EGFR overexpression has been found in 15% of unselected, but half of triple-receptor-negative (TRN) breast cancers (4), and HER2 is amplified in 20 to 30% of breast cancers. All these changes lead to enhanced malignant phenotypes and highly aggressive disease (5,6). HER2-overexpressing breast cancers also exhibited the capability of resistance to many first-line chemotherapy and tamoxifen (7–9). Moreover, many fundamental studies revealed that EGFR and HER2 have a close relationship with the aberrant activation of Ras proteins, which is implicated in facilitating virtually all aspects of the malignant phenotype, including cellular proliferation, transformation, invasion and metastasis (10).

Growth factor receptor bound protein 2 (Grb2) is one of the most important proteins participating in EGFR and HER2 signal transduction. It consists of one Src-homology 2 (SH2) domain and surrounded by two Src-homology 3 (SH3) domains. The SH2 domain interacts with phosphotyrosines (pTyr) on target proteins, while the SH3 domains interact with proline-rich sequences (11). Once the membrance receptor is activated, Grb2 serves as an adaptor protein in many signal transductions including the mitogen-activated protein kinase (MAPK) cascade for promoting cell division and/or differentiation (12). Silencing the Grb2 expression reduced cell growth in vitro indicating the GRB2 protein could be a good target for cancer therapy (13). The SH2 domain is one of the most prevalent protein-binding modules for protein-protein interaction which mediate the formation of multiprotein complexes during signaling. SH2 domains specifically function in protein tyrosine kinase (PTK) pathways due to the dependence of their binding on tyrosine phosphorylation (14). In cells, the specific association of SH2 domains and tyrosine phosphopeptides is to mediate the reversible relocalization of proteins, which is important for efficient propagation of PTK signals (15). The specificity of SH2 domain has allowed these domains to function as probes to detect tyrosine phosphorylation in signaling proteins (16,17). We hypothesized that Grb2-SH2 domain may serve as a negative inhibitor if it binds to an activated receptor in a living cell, but lacks SH3 domains to bind those downstream modules.

In order to investigate the possibility of our hypothesis, we utilized PTD as a tool to bring SH2 domains into living cells. The most popular PTD is HIV-1 TAT48-60 (GRKKRRQRRRPPQ), which has 12 amino acid arginine-rich peptides with the ability to rapidly translocate outside proteins into cells both in vivo and in vitro. Fusion of this PTD with proteins and peptides was proved to facilitate effective transduction of the fused cargos into cultured cells and animal tissues while preserving their biological activity (18,19). In this study, we constructed, expressed, and purified a fusion protein containing an SH2 domain derived from Grb2 and a PTD domain, and investigated its potential functions in breast cancer cells.

Materials and methods

All chemicals were of analytical grade and purchased from commercial suppliers.

Bacterial strains, plasmids and cell lines

E. coli [DH5α and BL21 (DE3)] (Invitrogen, Carlsbad, CA, USA) were cultured in Luria-Bertani broth medium (LB) and stored at −70°C. pET-16b was a commercial product (Novagen, Billerica, MA, USA) and stored at −20°C. The reconstructed plasmid pET-16b-ptd was a gift from Professor Hua Han, Department of Medical Genetics and Developmental Biology, Fourth Military Medical University (20). The breast cancer cell lines HER2-negative MDA-MB-231 and HER2-positive SK-BR-3 were preserved in the Department of Molecular Biology of the Fourth Military Medical University and was routinely cultured in Dulbecco’s modified Eagle’s medium (DMEM) or RPMI-1640 medium supplemented with 10% fetal bovine serum (Gibco by Invitrogen, Carlsbad, CA, USA), 100 U/ml penicillin and 100 μg/ml streptomycin. Cell cultures maintained at 37°C in a humidified 5% CO2 atmosphere.

Expression vector construction

The Grb2-SH2 coding DNA sequence was the native sequence of Grb2-SH2 as reported in Genbank (CCDS11721.1) without any modification. Grb2-SH2 cDNA was generated by PCR using the cDNAs of human lymphocytes constructed by the Department of Medical Genetics and Developmental Biology of Fourth Military Medical University using the following primers: SH2 sense primer, 5′-GGA TCC CTA CCG CAG GAA TAT C-3′ and SH2 anti-sense primer, 5′-CTC GAG AAA CCA CAT CCG TGG-3′. PCR products were purified using QIAquick Gel Extraction Kit (Qiagen, Hilden, Germany). Resulting PCR products were digested with BamHI and XhoI into the PGEM-T-Easy reporting plasmid and subsequently sub-cloned into digested pET-16b-ptd containing a PTD coding sequence (5′-TAT GTA TGG TAG GAA GAA ACG TCG ACA GCG TCG TCG-3′) derived from HIV-1 TAT48–60 (GRKKRRQRRRPPQ) and a ten-Histidine-tag sequence for easy purification to construct the expression vector pET-16b-ptd/grb2-sh2. Meanwhile, we also designed a mutant contrast for Grb2-SH2 sequence with a loss of 60 bases in sequence using a forward primer (5′-GAA GTT CAA TTC TTT GCG GTA GGG ATC-3′) and a reverse primer (5′-CCG GGG ATC CCT ACC GCA AAG AAT TG-3′) and inserted into digested PGEM-T-grb2-sh2 and pET-16b-ptd/grb2-sh2. The subsequent DNA sequencing and BLAST program (http://www.ncbi.nlm.nih.gov/BLAST/) confirmed all insertions to be correct.

Protein expression and purification

The reconstructed plasmids were transformed into E. coli BL21 (DE3) and a single colony was picked and grown overnight in 5 ml LB supplemented with 100 μg/ml ampicillin at 37°C then diluted 1:100. Protein expression induced by 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) (Sigma-Aldrich, St. Louis, MO, USA). After induction for 4 h at 33°C, approximately 10 g of wet weight cells from 1-liter culture were harvested by centrifugation at 6,000 x g for 30 min at 4°C followed by re-suspension in 60 ml of Tris-HCl (50 mM, pH 8.0) containing 1 mM phenylmethylsulfonyl fluoride (PMSF) for enzyme stability. The E. coli cells were then pulse-sonicated for 10×1 m (10 sec working and 50 sec resting on ice, 300 W). The lysate was centrifuged at 15,000 x g for 30 min at 4°C. The recombinant His-tagged PTD-Grb2-SH2 proteins were purified from the cell lysate using 1 ml of HisTrap™ Ni++ charged columns (Amersham Pharmacia Biotech, Uppsala, Sweden). The lysate was dialyzed against binding buffer (20 mM Tris-HCl, pH 7.4, 500 mM NaCl, 5 mM imidazole) overnight at 4°C. Subsequently, the dialyzed lysate was injected into the Ni-NTA resin column and binding for 10 min. The unbound proteins were washed away by using 10 volumes of binding buffer. The His-tagged proteins were eluted with elution buffer (20 mM Tris-HCl, pH 7.4, 500 mM NaCl, 20 mM imidazole). The eluate was analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and western blot analysis. The fractions containing the purified fusion protein were pooled and dialyzed against 100 volumes of 20 mM Tris-HCl (pH 7.4) overnight at 4°C. The protein concentration was determined using a DC protein assay kit (Bio-Rad, Hercules, CA, USA) with a bovine serum albumin (BSA) standard. The purified protein was stored at 4°C for use within 1 week or at −70°C for long-term storage.

SDS-PAGE and western blot analysis

SDS-PAGE was followed the procedure described by Laemmli (21). For western blot analysis, the proteins were separated by SDS-PAGE on a 15% gel and transferred onto a polyvinylidene difluoride (PVDF) membrane (Bio-Rad) in transfer buffer [25 mM Tris, 192 mM glycine, 15% (vol/vol) methanol] at 4°C at 100 V for 30 min. After blocking with 5% milk in phosphate-buffered saline (PBS, 50 mM phosphate and 0.9% NaCl; pH 7.2) at room temperature for 1 h or overnight at 4°C, the membrane was incubated for 1 h with horseradish peroxidase (HRP)-labeled antibody against the His-tag (Qiagen; 1:2,000 with 2.5% milk in PBS). The membrane was then washed five times with PBS-T (PBS plus 0.05% Tween-20) and two times with PBS. The target proteins were visualized with the enhanced chemiluminescence detection system (ECL, GE Healthcare, Piscataway, NJ, USA).

Immunofluorescence assay

Cells on cover slips were incubated with the target proteins for 4 h. The samples were fixed in 4% paraformaldehyde, permeabilized in PBS with 0.1% Triton X-100, blocked in 2% normal rabbit serum in PBS, and then incubated overnight at 4°C with mouse anti-His polyclonal antibody (Santa Cruz Biotechnology, Santa Cruz, CA, USA) at a 1:400 dilution. Cells were extensively washed with PBS and incubated with the fluorescein-labeled rabbit anti-mouse secondary antibody (Dako, Glostrup, Denmark) at a 1:100 dilution at room temperature for 2 h followed by further washing. The results were assessed under a reflected light fluorescence microscope (BH2-RFC, Olympus, Tokyo, Japan).

Cell viability assay

The MTT [3-(4, 5-dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium bromide] assay was used to investigate the cytotoxicity of the fusion protein. Breast cancer cells were plated in 96-well plates (5×104 cells per well) in septuplicate. The fusion proteins (including target protein and mutant contrast) were added into the culture medium (RPMI-1640 medium supplemented with 10% fetal bovine serum) in different concentrations (0.05 mg/ml, 0.1 mg/ml or 0.2 mg/ml). At the indicated time, 20 μl aliquots of 5 mg/ml MTT (Sigma) in PBS were added to each well and incubated for 4 h followed by the addition of 150 μl of Me2SO. The A490 values were assayed in a Sunrise microplate reader (Tecan, Groedig, Austria). Proliferation in vitro was also determined by 5-ethynyl-2-deoxyuridine (EdU, Ribobio, Guangzhou, China). EdU incorporation was determined using a Cell-Light™ EdU Cell Proliferation Kit (Ribobio), according to the supplier’s instructions. The electronic microscope was used to observe the ultrastructure of treated breast cancer cells.

Cell apoptosis analysis

Annexin V-FITC/PI staining was performed using the Elite ESP flow cytometry (FACSCalibur, Becton-Dickinson Immunocytometry Systems, San Jose, CA, USA) at 488 nm according to the manufacturer’s guidelines. Briefly, cells were incubated with PI and Annexin V-fluorescein isothiocynate in the darkness at room temperature. Flow cytometric analysis was immediately performed for apoptosis analysis and the data were analyzed using the Cell Quest Pro software (BD Biosciences, San Jose, CA, USA). Transmission electron microscope observation was carried out with the assistance of the Laboratory Center of Electron Microscope from Fourth Military Medical University.

Statistical analysis

Statistical analysis was performed using the SPSS 13.0 software package for Windows.

Results

Sequence synthesis, cloning, expression, purification and identification

Using the recombinant DNA technology, the PCR amplification fragments consistent with that expected for Grb2-SH2 (310 bp) and Grb2-SH2-Mutant (250 bp) were detected on an agarose gel and the DNA sequences were confirmed by automatic sequencing. The successful construction of the expressed recombinant pET-16b-ptd/grb2-sh2 and pET-16b-ptd/grb2-sh2-Mutant was confirmed by restriction mapping (Fig. 1A). In the pET system, target genes are positioned downstream of the bacteriophage T7 late promoter. Typically, products contain a prophage (λDE3) encoding the highly processive T7 RNA polymerase under control of the IPTG-inducible lacUV5 promoter that ensures tight control of recombinant gene basal expression. SDS-PAGE showed the expression of soluble proteins with a molecular mass value consistent with that expected for PTD-Grb2-SH2 (17.6 kDa) and PTD-Grb2-SH2-Mutant (16 kDa). Recombinants PTD-Grb2-SH2 and PTD-Grb2-SH2-Mutant were expressed in E. coli after induction of IPTG. In order to obtain maximum soluble protein expression, we adjusted conditions, including temperature and time. Finally, we determined that a temperature of 33°C and induction time of 4 h was optimal for obtaining maximum amount of soluble protein (Fig. 1B). HisTrap™ Ni++ charged columns (Amersham Pharmacia, Piscataway, NJ, USA) were used to purify the His-tagged proteins. Purification products were analyzed by 15% SDS-PAGE and identified by western blot analysis (Fig. 1C).

Figure 1

Construction, expression and purification of PTD-Grb2-SH2 and PTD-Grb2-SH2-Mutant. (A) The Grb2-SH2 coding frame is presented with 10 histidines and the HIV-1 TAT domain. (B) The expression of the recombinants. Lane M, protein molecular weight marker; lane 1, E. coli BL21(DE3) lysates after IPTG; lane 2, pET-16b plasmid after IPTG; lane 3, pET-16b-ptd/grb2-sh2 before IPTG; lane 4, pET-16b-ptd/grb2-sh2 recombinant after IPTG; lane 5, the crude lysate supernatant of induced pET-16b-ptd/grb2-sh2 after IPTG; lane 6, pET-16b-ptd/grb2-sh2-Mutant before IPTG; lane 7, pET-16b-ptd/grb2-sh2-Mutant after IPTG; lane 8, the crude lysate supernatant of induced pET-16b-ptd/grb2-sh2-Mutant after IPTG. (C) The purification and identification of the recombinants. Lane M, protein molecular weight marker; lane 1, the purified PTD-Grb2-SH2-Mutant fusion protein; lane 2, the purified PTD-Grb2-SH2 fusion protein; lane 3, the purified PTD-Grb2-SH2-Mutant fusion protein was identified by western blot analysis; lane 4, the purified PTD-Grb2-SH2 fusion protein was identified by western blot analysis.

Transduction of the recombinants into breast cancer cell line MDA-MB-231

An immunofluorescence assay with an anti-His-tag antibody was used to investigate whether the PTD-Grb2-SH2 and PTD-Grb2-SH2-Mutant could be transduced into living cells as designed. Purified PTD-Grb2-SH2 and PTD-Grb2-SH2-Mutant were added to cultured MDA-MB-231 cells for 2 h of incubation. Then, both recombinants were observed in the cells under a fluorescent microscope (Fig. 2). As shown in Fig. 2B and D, the recombinants including PTD-Grb2-SH2 and PTD-Grb2-SH2-Mutant were dispersed throughout the cytoplasm and mainly located in the nucleus, indicating HIV-1 TAT48-60 (GRKKRRQRRRPPQ) helped the target peptides to pass through both the cellular and nuclear membranes in living cells as reported (22).

Growth inhibition of PTD-Grb2-SH2 in the breast cancer cell lines MDA-MB-231 and SK-BR-3

The fresh recombinants of PTD-Grb2-SH2 and PTD-Grb2-SH2-Mutant were incubated with the breast cancer cell lines MDA-MB-231 and SK-BR-3 to investigate whether the new proteins could affect the proliferation of breast cancer cells in vitro. Fig. 3A and B show that the proliferation of MDA-MB-231 was inhibited by PTD-Grb2-SH2 in a time-dependent manner, but not by the mutant protein. Due to the similar inhibition effects of PTD-Grb2-SH2 in two different breast cancer cell lines, we chose the SK-BR-3 cells to look for the optimal concentration. Fig. 3C reveals that the recombinant PTD-Grb2-SH2 exhibited significant toxicity to breast cancer cells in a dose- and time-dependent manner in vitro. EdU identified the proliferation rates of MDA-MB-231 and SK-BR-3 cells. After incubation with PTD-Grb2-SH2 (0.1 mg/ml) for 12 h, a decreased rate of cell proliferation was detected in MDA-MB-231 cells, compared to the untreated group (24.6±1.4 vs. 39.6±1.3%, P<0.01). The proliferation rate of SK-BR-3 cells was also found to be decreased in the treated group, compared to the control group (13.4±1.1 vs. 37.6±2.2%, P<0.01). However, there is no difference between the cells treated with PTD-Grb2-SH2-Mutant (0.1 mg/ml) and control in either breast cancer cell line (Fig. 4).

PTD-Grb2-SH2 induces apoptosis in MDA-MB-231 and SK-BR-3 cells

By light microscopy, after 12 h incubation with the fusion protein (0.1 mg/ml), many breast cancer cells began to shrink and lose their normal fibroblast-like shape compared with the untreated cells. We gathered the treated cells to be observed under an electron microscope. Characteristic forms associated with cellular apoptosis could be observed, including the shrinkage of cellular and nuclear membranes and the appearance of many high-density structures and vesicles in MDA-MB-231 and SK-BR-3 cells were observed. Many vesicles appeared in the cytoplasm and the chromosome condensed into armillary shapes and concentrated beneath karyotheca, resulting in crescent or ring shapes (Fig. 5). We identified these changes as apoptotic phenomena, suggesting that PTD-Grb2-SH2 may induce apoptosis in breast cancer cells. To further determine the cytotoxicity of PTD-Grb2-SH2 on cell proliferation, MDA-MB-231 and SK-BR-3 cells were exposed to PTD-Grb2-SH2 (0.1 mg/ml) for 12 h. In order to differentiate this from necrosis and to confirm it as apoptosis, we performed fluorescein-conjugated Annexin V (Annexin V-FITC) flow cytometry. We quantitated the number of cells undergoing apoptosis. Our results showed that PTD-Grb2-SH2 induced apoptosis in 14.2% of MDA-MB-231 cells and 19.5% of SK-BR-3 cells, compared to the controls (6.3 and 11.3%, respectively) (Fig. 6).

Discussion

Although the discovery and characterization of HER2 and herceptin have resulted in great progress in breast cancer treatment, many patients still eventually relapse. Consequently, there is an urgent need for additional therapeutic strategy other than HER2 signaling pathway. Grb2 is an important adapter protein and the first trigger for many cellular signal pathways involved in the processes of cell proliferation and mitogenesis (23). Blocking the interaction between pTyr-containing activated receptors and the SH2 domain of Grb2 is considered to be an effective and non-cytotoxic strategy in the development of new anti-proliferate agents due to its potential to shut down the Ras activation pathway (24). This makes the Grb2-SH2 domain an ideal target for breast cancer treatment.

In this study, we designed, constructed, expressed and purified a fusion protein that contained a PTD domain and a Grb2-SH2 domain. The PTD domain is considered to deliver proteins of more than 120 kDa into living cells and tissues (25,26). Fusion proteins containing HIV-1 TAT have been reported to be successfully transduced into tumor cells and applied for anticancer therapy (27–29). We chose TAT48–60 (GRKKRRQRRRPPQ) as the PTD domain for transduction because this motif is the smallest one carrying PTD function without interfering with the target protein’s function (22). Our data confirmed that both the fusion protein and mutant protein successfully passed through the cellular and nuclear membranes of living breast cancer cells with the help of PTD domain. TAT48-60 delivery is a convenient research tool to study the function of small target peptides or small proteins in living cells. In previous studies we used this PTD domain to successfully deliver SH3 domains into the leukemia cell line K562 and the hepatocarcinoma cell line HepG-2 (30,31). The Grb2-SH2 domain is a highly preserved domain in both prokaryote and eukaryote, and the fusion proteins we designed are small proteins with molecular weight less than 20 kDa. Although the prokaryotic expression system we chose is not a theoretically proper expression system for a eukaryotic protein, we confirmed the right construction by automatic sequencing and identified the target proteins by western blot analysis to prevent miss-construction and expression.

In function assay, we directly incubated the recombinants with HER2-negative MDA-MB-231 and HER2-positive SK-BR-3 to test the effects on the proliferation of breast cancer cells. Our data exhibited that PTD-Grb2-SH2 inhibited the proliferation of both breast cancer cell lines, but the PTD-Grb2-SH2-Mutant did not exert any inhibition to these cell lines, indicating the loss in basic sequence of Grb2-SH2 will result in dysfunction of the target protein. Our results also revealed that PTD-Grb2-SH2 exhibited significant toxicity to breast cancer cells in a dose- and time-dependent manner in vitro, which is an appropriate characteristic for anticancer drug developing. To date, many strategies have been used to inhibit the function of Grb2 in order to block crucial intracellular signals. Some studies have focused on designing and synthesizing phosphotyrosines or their mimics based upon space structures of Grb2 (32). Grb2 is made up of one SH2 domain surrounded by two SH3 domains. While both SH2 and SH3 domains possess a strong ability to recognize and specifically bind to their ligands, SH2 domains are the major known binding modules for tyrosine-phosphorylated proteins and are the prototype for protein-protein interactions that mediate the formation of multi-proteins complexes during signaling (33). Integrating these basic results with our data, we think that the strength and specificity of Grb2-SH2 for its ligands give this domain the ability of inhibiting Grb2 related signaling in living breast cancer cells.

We chose HER2-negative MDA-MB-231 and HER2-positive SK-BR-3 in the function assay as Grb2 intermediates the mitogen-activated protein kinase (MAPK) pathway and also regulates receptor trafficking (34–36). It works with various RTKs, with EGFR being its major binding partner (37,38), which regulates many cell functions. Our data showed PTD-Grb2-SH2 inhibited HER2-negative and -positive breast cancer cells, indicating this fusion protein possesses non-specificity for single pathway. This is also the reason why we did not test the downstream molecules or pathways for this protein. There are too many molecules and pathways associated with Grb2 that remain to be elucidated in further experiments. Nevertheless, we still used electron microscopy and flow cytometry assay to identify the apoptotic phenomena induced by PTD-Grb2-SH2.

In conclusion, our study successfully constructed a target fusion protein expressing both PTD and Grb2-SH2 domains and showed that expression of the fusion protein resulted in growth inhibition and cell death in breast cancer cells regardless of HER2-phenotype. We proved that the TAT48–60 is a useful tool and could be capable of delivering outside proteins through the plasma membrane of living cells, and even delivering a protein directly to the cell nucleus. This technique will help us to demonstrate protein function in living cells. We also demonstrated that the SH2 domain is a highly conserved protein functional domain and can maintain its biological activity even when expressed in bacteria. The extent of inhibition depend on the concentration of the protein and the length of the time, moreover, it suggested that the protein might induce apoptosis in breast cancer cells. These results indicate that the PTD-Grb2-SH2 protein has the potential to be developed for treatment of breast cancer.

Acknowledgements

This study was supported by grant no. 30901457 from National Natural Science Foundation of China.

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Dierck K, Machida K, Mayer BJ and Nollau P: Profiling the tyrosine phosphorylation state using SH2 domains. Methods Mol Biol. 527:131–155. 2009. View Article : Google Scholar : PubMed/NCBI

34

Jiang X, Huang F, Marusyk A and Sorkin A: Grb2 regulates internalization of EGF receptors through clathrin-coated pits. Mol Biol Cell. 14:858–870. 2003. View Article : Google Scholar : PubMed/NCBI

35

Yamazaki T, Zaal K, Hailey D, Presley J, Lippincott-Schwartz J and Samelson LE: Role of Grb2 in EGF-stimulated EGFR internalization. J Cell Sci. 115:1791–1802. 2002.PubMed/NCBI

36

Burke P, Schooler K and Wiley HS: Regulation of epidermal growth factor receptor signaling by endocytosis and intracellular trafficking. Mol Biol Cell. 12:1897–1910. 2001. View Article : Google Scholar : PubMed/NCBI

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Schulze WX, Deng L and Mann M: Phosphotyrosine inter-actome of the ErbB-receptor kinase family. Mol Syst Biol. 1:2005.0008. 2005. View Article : Google Scholar : PubMed/NCBI

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Seiden-Long I, Navab R, Shih W, et al: Gab1 but not Grb2 mediates tumor progression in Met overexpressing colorectal cancer cells. Carcinogenesis. 29:647–655. 2008.PubMed/NCBI

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