Stathmin is overexpressed and regulated by mutant p53 in oral squamous cell carcinoma
- Hai-long Ma†1,
- Shu-fang Jin†1,
- Wu-tong Ju†1,
- Yong Fu1,
- Yao-yao Tu1,
- Li-zhen Wang2,
- Zhi-yuan Zhang1Email author and
- Lai-ping Zhong1Email author
© The Author(s). 2017
Received: 20 May 2017
Accepted: 31 July 2017
Published: 14 August 2017
The aim of this study was to investigate the oncogenic function and regulatory mechanism of stathmin in oral squamous cell carcinoma (OSCC).
Two-dimensional electrophoresis and liquid chromatography-tandem mass chromatography were applied to screen differentiated proteins during carcinogenesis in OSCC. Cell Counting Kit-8 (CCK-8) assays, colony formation, migration, flow cytometry, immunofluorescence and a xenograft model were used to detect the function of stathmin. The correlation between stathmin and p53 expression was analyzed using immunohistochemistry. Mutant/wild type p53 plasmids and small interfering RNA were used to examine the regulation of stathmin. Chromatin immunoprecipitation assays and luciferase assays were performed to detect the transcriptional activation of stathmin by p53.
Overexpression of stathmin was screened and confirmed in OSCC patients and cell lines. Silencing expression of stathmin inhibited proliferation, colony formation and migration and promoted apoptosis. Poly ADP ribose polymerase (PARP) and cyclin-dependent kinase 1 (cdc2) were activated after silencing the expression of stathmin. Suppression of tumorigenicity was also confirmed in vivo. Mutant p53 transcriptionally activated the expression of stathmin in HN6 and HN13 cancer cells, but not in HN30 cells harboring wild type p53.
These results suggest that stathmin acts as an oncogene and is transcriptionally regulated by mutant p53, but not by wild-type p53. Stathmin could be a potential anti-tumor therapeutic target in OSCC.
Oral squamous cell carcinoma (OSCC) accounts for more than 300,000 new cases each year and unfortunately has poor clinical outcomes with a 5-year survival rate of only 50–60%, which is even lower in patients with locally advanced disease [1, 2]. Despite substantial progress made in the past few decades to understand the mechanism of carcinogenesis, treatment strategy has developed slowly for OSCC. In terms of treatment, surgery remains the best option for OSCC patients but it is not effective on late-stage metastatic tumors . Betel quid, alcohol, tobacco and human papillomavirus are widely regarded as carcinogens for oral cancer. It is generally accepted that carcinogenesis of OSCC is a” Darwinian process” that involves a series of gene mutations causing the selected growth of mutated cells which replace normal cells in a specific region . TP53, CDKN2A, PIK3CA, PTEN, NOTCH and EGFR are the most frequently mutated genes in head and neck cancer . Mutations in the TP53 gene have been widely studied in OSCC, with about 75% of mutations occurring in the DNA binding domain. Mutations in this region can activate a series of oncogenes to promote tumor progression, while wild-type p53 (wtp53) may inhibit tumor growth . Neomorphic mutant p53 (mutp53) activities are spread over multiple levels, impinging on chromatin structure, transcriptional regulation and microRNA maturation, shaping the proteome and cellular metabolic pathways, and also exerting cytoplasmic functions and displaying cell-extrinsic effects . Patients carrying mutant p53 are always resistant to chemotherapy and radiotherapy with poor prognosis . Therefore, it is crucial to deeply investigate the molecular network involved in gain-of-function (GOF) mutations of p53 in OSCC.
Stathmin 1 (hereafter referred to as stathmin), also known as oncoprotein 18/op18, is a microtubule-destabilizing phosphoprotein that is ubiquitously expressed and involved in regulating the global dynamics of mitotic and interphase microtubules . Stathmin prevents the incorporation of αβ-tubulin dimers in growing microtubules, resulting in eventual microtubule destabilization . Stathmin overexpression has been reported in many types of human cancers and is associated with promotion of cancer cell proliferation, migration, invasion and resistance to paclitaxel [11–18]. In our previous study, we observed an interaction between stathmin and p53 in OSCC . However, the relationship between stathmin and mutant/wild-type p53 remains unclear. Conflicting evidence has been reported regarding the relationship between stathmin and mutant/wild-type p53 expression [20, 21]; thus, the regulatory mechanism between mutp53 and stathmin in OSCC requires further investigation.
In this study, we demonstrate that stathmin was overexpressed in OSCC patients and cell lines, which promoted progression and tumorigenesis. Mechanistically, stathmin was regulated by mutp53 but not by wtp53 in OSCC.
Two-dimensional electrophoresis (2-DE) and liquid chromatography-tandem mass chromatography (LC-MS/MS)
2-DE and LC-MS/MS in human immortalized oral epithelial cells (HIOECs) and in HB96 cells were described thoroughly in our previous study . Briefly, HIOECs and HB96 cells were lysed, sonicated and protein was quantified. First-dimensional IEF was completed with an IPGphor IEF System (Amersham Biosciences, Uppsala, Sweden) and second-dimensional SDS–PAGE was performed with a Hoefer SE 600 Ruby System (Amersham). Differentially expressed protein spots were excised and digested for mass spectroscopy. The peptide mixtures were isolated and identified by a Finnigan LTQ mass spectrometer coupled with the Surveyor HPLC system (Thermo, Sunnyvale, CA). Differentially expressed protein identification in MS/MS raw data was determined using the SEQUEST program in the BioWorks 3.1 software suite (University of Washington, licensed to Thermo Finnigan) based on the International Protein Index human database version 3.15.1.
For immunohistochemistry, tissue samples from 60 patients and 16 healthy subjects were prepared according to the following protocol for immunohistochemical staining. Briefly, the sections were heated by water bath at 100 °C with citrate buffer solution (pH 6.0) for 20 min to retrieve antigen. The primary antibodies were rabbit monoclonal anti-stathmin (Abcam, ab52630, dilution 1:500; Cambridge, MA) and mouse monoclonal anti-human p53 (Abcam, ab28, 1:500). Microscopic examination of stathmin immunohistochemical staining was conducted by two blinded pathologists. The intensity of the stathmin immunoreaction was scored as following: 0 = negative, absence of stained cells; 1 = weak; 2 = moderate; 3 = strong. The immunohistochemical staining score was calculated by multiplying the percentage of positive cells and the staining intensity as described in the literature . This study was approved by the Ethics Committee of Ninth People’s Hospital, Shanghai Jiao Tong University School of Medicine. Informed consent was obtained from all patients for use of their tissues.
For immunofluorescence, Alexa Fluor 488-Affini goat anti-rabbit IgG (Abcam, ab150077, 1:250) was used as the secondary antibody. DAPI was used to stain cell nuclei. The morphology of microtubules was observed using rat monoclonal anti-tubulin antibody (Abcam, ab6160, 1:500) under a laser scanning confocal microscope (LSM-710, Carl Zeiss, Gottingen, Germany).
The cell lines used in the present study were HIOEC, HB96 (derived cancerous cell line from HIOEC), HN4, HN6, HN12, HN13, CAL27, HN30 and SCC25. HIOECs and HB96 cells were obtained from our previously established in vitro cellular carcinogenesis model of OSCC . HN4, HN6, HN12 and HN13 lines were obtained from tongue squamous cell carcinoma patients, while HN30 cells were derived from a pharyngeal carcinoma. All of the HN lines were provided by the University of Maryland Dental School, Baltimore, MD. CAL27 and SCC25 lines were purchased from ATCC (Manassas, VA). The mutant phenotype of TP53 in cells was detected as described by Song et al. . All OSCC cell lines used in this study were HPV negative. HIOECs were cultured in defined keratinocyte-SFM (Gibco, Carlsbad, CA) and SCC25 were cultured in DMEM/F12 (Gibco). The other cell lines were cultured in DMEM (Gibco). Both DMEM and DMEM/F12 were supplemented with 10% fetal bovine serum, 1% glutamine, and 1% penicillin-streptomycin (Gibco). All cells were cultured in a humidified atmosphere of 5% CO2 at 37 °C.
Western blot assay
Protein extraction and western blots were performed as previously described . Membranes were probed with primary rabbit antibodies against stathmin, p53, beta-actin, cyclin D1, bcl-2 and GAPDH, purchased from Abcam; or with antibodies against cleaved-PARP, PARP, cyclin B1, cdc2 and p-cdc2 (Tyr15), purchased from Cell Signaling Technology, Danvers, MA; and finally with fluorescent-based anti-rabbit IgG secondary antibody (Fermentas, Vilnius, Lithuania). Immunoreactive bands were scanned and analyzed using Image J software (NIH, Bethesda, Maryland) and the Odyssey Infrared Imaging System (LI-COR Biosciences, Lincoln, NE).
Real-time PCR assay
Total RNA from cultured cells was isolated at 80% confluence with TRIzol reagent (Takara, Dalian, China) according to the manufacturer’s instructions. Total RNA (1 μg) was reverse transcribed into first strand cDNA with PrimeScript reverse transcriptase (Takara, Dalian, China), random primers and oligo dT primer in a 20 μl reaction mixture according to the manufacturer’s protocol. Real-time PCR was performed with the StepOnePlus™ Real Time System (Applied Biosystems, Foster City, CA). The primer sequences were as follows. Stathmin: 5′-CTCGGACTGAGCAGGACTTTC-3′ (forward) and 5′-ATTCTTTTGACCGA GGGCTG-3′ (reverse); p53: 5′-CCAGAAAACCTACCAGGGCA-3′ (forward) and 5′-GAATGCAAGAAGCCCAGACG-3′ (reverse); and GAPDH: 5′-CCTCTGACTTCAACAGCGAC-3′ (forward) and 5′-TCCTCTTGTGCTCTTGCTGG-3′ (reverse). Relative quantification of mRNA levels compared to the internal control gene GAPDH was calculated according to the 2-ΔΔct method. Melting and amplification curves were performed to determine the specificity of stathmin amplification. All samples were assayed in triplicate.
Small interfering RNA (siRNA) and plasmid transfection
The siRNAs against stathmin, p53 and scrambled control sequence were designed as follows. siRNA-stathmin: 5′-CAGCCCUCGGUCAAAAGAAtt-3′ (si#1); 5′-GCUGCCAAACUGGAACGUUtt-3′ (si#2); siRNA-p53: 5′-GTCCAGATGAAGCTCCCAGAAdTdT-3′; siRNA–negative control (NC): 5′-UUCUCCGAACGUGUCACGUdTdT-3′. siRNAs were chemically synthesized by Shanghai GenePharma Co., Shanghai, China. In addition, si#2 sequence was packaged into the lentiviral pGLV-h1-GFP-puro vector for in vivo experiments. Cells were transfected using Lipofectamine™ 3000 Transfection Reagent (Invitrogen, Carlsbad, CA) according to the manufacturer’s instruction. Transfection efficiency for the siRNAs in HN4 and HN13 cells was 50% ~ 70%. Transfected cells (1000–5000 per well) were seeded in 96-well plates and cultured for 5 days. Cell Counting Kit-8 (CCK-8; Dojindo, Kumamoto, Japan) solution (10 μl) was added to each well followed by incubation at 37 °C for an additional 3 h. The absorbance at 450 nm was measured to assess cell viability. The experiment was performed in triplicate.
Colony formation assay
After transient transfection for 24 h with siRNA, 1000 cells were plated in 6-well plates for 3 weeks until macroscopic colonies were visible. The colonies were fixed and stained using crystal violet. Colonies of more than 50 cells were counted under a dissecting microscope. The data were expressed as means ± SD from three independent experiments.
Briefly, 3–5 × 104 transfected cells were seeded in the upper chambers of filter inserts containing serum-free medium. The lower chambers held 500 μl medium per well containing 10% FBS in 24-well plates. After 24–36 h, the cells were stained using 5% crystal violet. The number of cells migrating to the lower chamber was counted. The experiment was performed in triplicate.
In vitro wound healing assay
Transfected cells were incubated overnight yielding confluent monolayers suitable for wounding. Wounds were scratched using a 10 μl pipette tip. At 0 h and 24 h, photographs were taken with a Nikon DMCI microscope (Nikon, Tokyo, Japan). The distance migrated by invasive cells to heal the wound area during this period was measured.
Cells transiently transfected with siRNA were harvested 48 h after incubation. These cells were then quantified by flow cytometry using the FITC-Annexin V Apoptosis Detection Kit (BD Biosciences, Franklin Lakes, NJ) according to the manufacturer’s instructions. The experiments were performed in triplicate.
Tumorigenicity assay in vivo
To evaluate the antitumor effect of stathmin in vivo, an OSCC xenograft model was generated in BALB/C nude mice (4–6 weeks old, 18–22 g) by subcutaneously injecting 2 × 106 HN13 cells after lentivirus transfection. The transfected cells were selected in medium containing 4 μg/μl puromycin to harvest stable stathmin-knockdown cells and scrambled control cells. Tumors were measured once per week for 5 weeks, and tumor weights were measured after sacrifice.
To construct the stathmin expression vector, the code domain sequence (CDS; NM_203401.1) was cloned into the eukaryotic expression vector pcDNA3.1 (Invitrogen). Wild-type p53 CDS (NM_ 000546) and a mutant sequence template (Shanghai GenePharma; 524A > G, 733 U > G, 844 U > C) were cloned into pcDNA3.1.
Chromatin immunoprecipitation assay (ChIP)
To explore the regulatory mechanism of stathmin in OSCC, ChIP was conducted in OSCC cell lines. ChIP assays were performed according to the manufacturer’s instructions using the SimpleChIP Enzymatic Chromatin IP Kit (magnetic beads; CST). Briefly, p53 and protein complexes were cross-linked inside HN6, HN13 and HN30 cells by the addition of formaldehyde (1% final concentration) to the cells in culture. Chromatin was digested with micrococcal nuclease and sonicated into 150–900 bp DNA/protein fragments. An aliquot of the cross-linked protein complexes was immunoprecipitated by incubation with either p53-specific antibody (CST) or IgG antibody overnight at 4 °C with rotation. Chromatin-antibody complexes were isolated from solution by incubation with ChIP-Grade Protein G Magnetic Beads for 1 h at 4 °C with rotation. The bead-bound immune complexes were then washed and eluted from the beads with elution buffer. Eluates were heated at 65 °C overnight to reverse the formaldehyde cross-linking and DNA was extracted. DNA samples from chromatin immunoprecipitation preparations were analyzed by PCR using primers spanning the stathmin gene (NM_203401.1) in the promoter region (forward, 5′-AATGGGGAGCTGGTTCGGA-3′; reverse, 5′-GTGTAGTCCTGTCCCGGAGG-3′). The transcription factor binding sites of p53 were predicted using the website of JASPAR (http://jaspar.genereg.net/)  and PROMO [28, 29]. The two predicted binding sites of CCAGGCCCACACCTG and GCAACCCCCGGCATT were amplified using the above promoter primers. Primers used in the real-time PCR mentioned above amplified the CDS region. The nonpromoter regions were used as control.
Luciferase assays were used to confirm the transcriptional activation leading to p53 GOF. 293 T and SCC25 cells were transiently transfected using transfection reagent Lipofectamine 3000 (Invitrogen). Cells were seeded in 12-well plates (1 × 105 cells/well) and grown to 70–80% confluence. Each stathmin promoter-luciferase construct was co-transfected into cells with pRL-TK (TK promoter Renilla luciferase construct as internal control). Luciferase activity was determined 24–48 h after transfection using a dual luciferase reporter assay system (Beyotime, Shanghai, China). Cell lysates (200 μl/well) were used for measurement of relative luciferase units in a luminometer by first mixing 20 μl of cell lysates with 100 μl of luciferase assay reagent to measure firefly luciferase activity, then subsequently adding 100 μl of Renilla luciferase reagent to measure Renilla luciferase activity. Data were normalized to Renilla luciferase activity (internal control) and expressed as arbitrary units. siRNA against p53 was co-transfected with p53 plasmids. NSC319726 is a mutp53 (R175H) reactivator, exhibiting growth inhibition in cells expressing mutp53 and showing no inhibition of wtp53 cells . NSC319726 was added after transfection with the mutp53 plasmids. All experiments were performed in triplicate.
All data were analyzed using the statistical software SPSS 13.0 for Windows (SPSS Inc., Chicago, IL). Student’s t-test and one-way analysis of variance were used to analyze immunohistochemical results. All hypothesis-generating tests were two-tail at a significance level of 0.05 (*P < 0.05, **P < 0.01). Data were presented as means ± standard error of the mean.
Identification of stathmin overexpression in OSCC
The correlation between stathmin expression and clinical characteristics in OSCC
parametric test value
t = 1.189
< 60 years
t = 0.218
≥ 60 years
F = 0.039
t = −0.470
t = −0.775
F = 14.757
< 0.001 *
t = 0.181
t = −0.702
Stathmin overexpression promotes progress of OSCC in vitro and in vivo
Positive correlation between stathmin and p53 expression in OSCC tissues
Stathmin expression is regulated by mutp53, not wtp53 in OSCC
In this study, we demonstrated that overexpression of stathmin promoted tumor progression and was regulated by mutp53 in OSCC. Overexpression of stathmin has been detected in several malignancies. Stathmin overexpression in hepatoma promoted local invasion, polyploidy formation, early recurrence and poor prognosis, suggesting that stathmin can be an effective therapeutic target . Proteins from pooled microdissected nasopharyngeal carcinoma and adjacent non-tumorigenic nasopharyngeal epithelial tissues were separated by 2-DE to find significant overexpression of stathmin in the tumors . Stathmin knockdown by siRNA in melanoma cells drastically repressed cell proliferation and migration, whereas ectopic expression of stathmin increased cell proliferation and migration , consistent with our results. From the reports in various carcinomas, overexpression of stathmin was very consistent in several tumors and acted as a potential oncogene.
Recently, possible mechanisms that may underlie the multiple functions of stathmin in cancer have attracted much attention. In our study, cleaved PARP (the 89-kDa cleaved product) was increased after silencing stathmin, which indicated the pro-apoptotic effect of stathmin knockdown in OSCC. Furthermore, bcl-2 and cyclin D1 increased after ectopic expression of stathmin. In agreement with these results, silencing of stathmin resulted in decreased expression of bcl-2 and survivin proteins and activation of caspase-3 and caspase-9 [38, 39].
Cdc2, also known as cyclin-dependent kinase 1, mainly regulates the progression of the G2/M phase. A conserved tyrosine (Tyr15 in humans) leads to inhibition of cdc2; its phosphorylation is thought to alter ATP orientation, preventing efficient kinase activity. Activation of cdc2 (try15) blocked cells at the G2/M phase which then induced apoptosis. In contrast, cyclin B1, the partner of cdc2, was unaltered in our study. It has also been reported that expression of p53 and p21 increased after silencing stathmin in gallbladder carcinoma . In our study, we confirmed that upregulation of p-cdc2 (try15) after stathmin silencing arrested the cell cycle and induced apoptosis in OSCC. The results suggest that the downstream signaling molecules of stathmin differ in various cancers.
Overexpression of stathmin was transcriptionally activated by mutp53 but not by wtp53 in OSCC in our study. In over 70% of cases, the TP53 mutations are missense, most frequently within the region of DNA binding. Although the spectrum of the TP53 missense mutations is vast – counting about 1800 different amino-acid changes– several hotspots in p53 mutants, particularly affecting residues R273, R248, R175 and G245, have been reported to be present with a higher frequency in head and neck cancer . Cells expressing mutp53 exhibited aggressive cancer phenotypes, such as enhanced cell survival, proliferation, invasion and adhesion, altered mammary tissue architecture and invasive cell structures . This was confirmed in SCC25 cells after transduction with mutp53 plasmids. Stathmin depletion caused a large percentage of the apoptosis occurring in both normal and cancer cell lines lacking p53 , while significant inhibition of proliferation was also observed in OSCC cell lines with mutp53. This suggests that stathmin depletion could be used therapeutically to induce apoptosis in tumors with or without p53 expression. In our study, three p53 mutants (R175H, G245C and R282W) were constructed into one plasmid to mimic the complicated mutations of p53 in OSCC. These three mutants contributed to the mutp53 gain of oncogenic function to promote invasive growth of head and neck cancer cells via inhibition of AMPK activation . Upregulation of stathmin was also shown to be mediated by GOF mutations in p53 in human hepatoma , as well as in breast cancer cell lines harboring mutp53 . The results of our study are in accord with the above reports. In type II high grade epithelial ovarian carcinomas, stathmin favored the binding and the phosphorylation of mutp53 by DNA-PKCS, eventually modulating mutp53 stability and transcriptional activity . It was speculated that a positive feedback loop between mutp53 and stathmin synergistically promoted the progression of malignant tumors. However, wtp53 has also been reported to downregulate stathmin expression [20, 45]. This discrepancy may be related to separate molecular driving mechanisms in different tumors, but our results nonetheless provide new insight into the interaction between mutp53 and stathmin overexpression in OSCC, which extends the network driving mutp53. The transcriptional regulation of STMN1 by p53 varies in different carcinomas, which deepens our understanding of GOF p53-driven tumors. Our results confirm that stathmin is a novel target of mutp53 and jointly promotes tumorigenesis and tumor progression.
In summary, we demonstrated that stathmin was overexpressed in OSCC, which promoted progression and tumorigenesis. Silencing stathmin activated cdc2 at try15 and cleaved PARP to induce cell cycle arrest and apoptosis in OSCC. Importantly, overexpression of stathmin was transcriptionally activated by GOF mutations of p53, not by wtp53. Thus, our findings indicate that stathmin may be a potential therapeutic target for the treatment of OSCC.
This study was supported by research grants 81672660 and 81472519 from the National Natural Science Foundation of China, by research grant 17SG18 from Shuguang Program of Shanghai Municipal Education Commission, by research grants JYLJ002, BXJ201728, and 15ZH2008 from Shanghai Jiao Tong University School of Medicine.
Availability of data and materials
Data sharing is not applicable to this article as no datasets were generated or analyzed during the current study.
HM, SJ, and WJ analyzed and interpreted both the clinical and experimental data; YF and YT analyzed the experimental data; ZZ and LZ analyzed the clinical data; LW and JL performed the histological examination; HM and LZ were the major contributors in writing the manuscript. All authors read and approved the final manuscript.
Ethics approval and consent to participate
Human Research Ethics Committee of Ninth People’s Hospital Shanghai Jiao Tong University School of Medicine (approved number: 2008–12).
Consent for publication
The authors declare that they have no potential competing interest.
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