Background
Human glioblastoma multiforme (GBM) is the most common and malignant type of brain tumors. Current treatment options such as surgical intervention, radiation therapy or cytotoxic chemotherapy do not significantly improve the median survival beyond approximately 12 to 18 months for patients with GBM [
1,
2]. Therefore, the identification and the development of novel and more efficient therapeutic approaches remain a crucial task for this disease.
Since GBM is characterized by particularly high levels of neovascularization, a therapeutic strategy based on angiogenic blockade appears to be promising. Actually, a number of strategies targeting new blood vessel formation have shown some success in preclinical models of GBM [
3,
4] and several clinical trials with anti-angiogenic agents are ongoing [
5]. An important feature of angiogenesis is the interaction of endothelial cells (EC) with surrounding extracellular matrix (ECM). Integrin binding mediates cell adhesion of ECs to surrounding ECM and regulates their survival, growth and mobility [
6]. Integrins and αVβ5 are predominantly expressed in proangiogenic ECs [
7,
8] and especially integrin αVβ3 has been found to be upregulated in ECs of GBM tumors [
9,
10]. Cilengitide, a cyclic pentapeptide mimicking the Arg-Gly-Asp (RGD) binding site of integrin ligands, was identified as a potent and selective integrin antagonist that interfered with binding of ECM components to αVβ3 and αVβ5 integrins [
11]. In preclinical models cilengitide had synergistic therapeutic effects with radioimmunotherapy in breast cancer and orthotopic brain tumor models [
12,
13]. However, expression of αVβ3 and αVβ5 integrins is not restricted to activated ECs. Both integrins are also in brain tumor cells [
14‐
16]. In fact, we have recently shown that cilengitide inhibits integrin-dependent signaling and induces apoptosis not only in endothelial but also in glioma cells thereby explaining the profound activity of integrin inhibitors in this disease [
17]. These data suggest that anti-angiogenic molecules directed towards integrins may have a multi-targeting effect on both endothelial and glioma cells.
An additional aspect to be considered for the design of novel therapeutic strategies against GBM is the ability of these tumors to escape anti-angiogenic monotherapy [
18,
19]. Therefore, it might be necessary to target multiple pro-angiogenic pathways in order to achieve significant anti-tumorigenic effects.
Here, we studied two angiogenic inhibitors targeting different angiogenic pathways, endostatin (ES) and tumstatin (Tum), and evaluated the anti-tumorigenic activity of the individual factors and a combination of both factors in an in vivo model of GBM. ES has been reported to interfere with integrin α5β1 and VEGFR-2 in ECs, while Tum binds αvβ3 and αVβ5 integrins and induces apoptosis in ECs [
20,
21]. In addition, microarray analysis of tumor tissue was performed to identify activation of alternative pro-tumorigenic signalling pathways in tumor cells.
Discussion
Angiogenesis plays a central role in tumor growth and metastasis. Since GBM tumors are highly vascularised, therapeutic strategies based on angiogenic blockade are particularly attractive for this entity. However, it has been observed that initial responses to anti-angiogenic therapy are frequently followed by tumor progression resulting in only limited survival advantage [
29‐
31]. This evasive resistance implies adaptation of tumors to angiogenic inhibitors via activation of alternative pathways that sustain tumor growth.
Accordingly, our approach was designed to simultaneously target different angiogenic signaling pathways and to investigate the activation of possible resistance mechanisms in a GBM model.
Our results show for the first time that the combined application of the integrin inhibitors ES and Tum significantly augment the inhibitory effect over each of the individual substances and that the ES + Tum combination exerts its antitumorigenic effects by both antiangiogenic and direct antitumorigenic activities. Finally, we found an up-regulation of the prolactin receptor in tumor cells treated with the ES + Tum combination and demonstrate a role of this receptor in the control of glioma cell proliferation in vitro.
In the present study, the antiangiogenic substances were delivered to a subcutaneous graft of G55 glioma cells using
ex vivo modified PAE cells, which were encapsulated in alginate microbeads. The microencapsulation technology ensures a continuous release of proteins, and has been successfully used by us and others in different animal models [
32,
33].
The efficacy of each angiogenic inhibitor was demonstrated on EC proliferation and wound assays in vitro and the combination of ES + Tum showed even additive inhibitory effects on endothelial cell proliferation. Local release of single inhibitors ES and Tum by encapsulated PAE cells resulted in inhibition of tumor growth in subcutaneously implanted GBM by about 58% and 50%, respectively, when compared to the control group, respectively. Strikingly, the combined application of ES and Tum inhibited tumor growth by about 83% tumor growth inhibition.
While these observations correlated with a pronounced decrease of vascular density in ES- treated tumors, treatment with Tum resulted in only minimal reduction of blood vessel density, suggesting that
in vivo tumor growth reduction mediated by Tum is mainly caused by a direct antitumorigenic activities and less through antiangiogenic mechanisms. A direct αVβ3 -dependent growth-inhibitory effect of Tum on glioma cells
in vitro and
in vivo has been previously describe by Kawaguchi et al. [
34]. On the other hand, the extent of tumor growth inhibition caused by the Es + Tum combination was higher than expected compared with the reduction level of vessel density. This fact prompted us to hypothesize that the ES + Tum combination exerts direct anti-neoplastic effects on glioma cells
in vivo, in addition to its antiangiogenic effect. This hypothesis was confirmed in our i
n vitro experiments, which showed reduced proliferation rates of glioma cells after treatment with the ES + Tum combination, but not after treatment with the single inhibitors. Moreover, the ES + Tum combination caused morphological changes and induced apoptosis in glioma cells. Since previous studies have demonstrated that integrin antagonists affect cell cycle progression and viability of glioma cell lines, even inhibiting signaling pathways similar to ECs [
34], we suggest that ES and Tum act through their respective integrin receptors on glioma cells, ultimately leading to inhibition of proliferation and induction of apoptosis. Nevertheless, further studies are necessary to clarify the effects of ES + Tum on glioma cells at the molecular level.
In order to gain further insights into possible mechanisms that enable tumor cells to escape anti-angiogenic therapies, we performed cDNA arrays using mRNA from tumor tissue treated with encapsulated PAE-WT cells or PAE cells releasing ES or Tum, either individually or in combination. Surprisingly, we identified only a few genes with a significant increase or decrease in expression level (mean signal log ratio ‡1.0) in the ES-, Tum- or ES + Tum-treated groups when compared with the control group. We focused our interest on the hormone prolactin (PRL) and its cognate receptor PRLR, which were up-regulated after treatment with Tum and ES + Tum, respectively. Validation of PRLR up-regulation in ES + Tum tissue sections by immunohistochemistry revealed a heterogeneous staining pattern with an intensive PRLR staining localized in well-defined tumor regions. Double immunostaining with apoptotic marker M30 and PRLR further showed that those areas with high levels of PRLR contained none or few apoptotic cells, whereas apoptotic regions presented low or no expression of PRLR. Similar results were obtained
in vitro after immunofluorescence staining for cleaved caspase-3 and PRLR in glioma cells treated with ES + Tum (data not shown). Based on these results, we assume that a subpopulation within the G55 cells does not undergo apoptosis after ES + Tum-treatment but rather proliferates via activation of the PRLR/PRL signaling axis. In Glioblastoma “Cancer Stem Cells” (CSC), a small subpopulation of self-renewing “stem-like” cancer cells, have been demonstrated to show resistance to commonly used anticancer therapies such as radiation [
35] and chemotherapy [
36,
37]. Clark et al. [
38] have observed a compensatory activation of multiple ERBB family receptors in GBM CSCs deprived of EGFR signal, suggesting an intrinsic GBM resistance mechanism for EGFR-targeted therapy. To what extent the PRLR-positive subpopulation found in ES + Tum-treated tumor consist of CSC needs to be further investigated in future studies.
Several studies have documented the involvement of the ligand PRL in the growth control of different tumors such as breast [
39], liver [
40] and prostate [
41] and further, PRL antagonists such as hPRL-G129R has been demonstrated to inhibit breast cancer growth
in vitro and in vivo [
42]. However, only little is known about the role of the PRLR/PRL-signaling axis in glioma cells. PRLR expression has been found in rat and human glioma cells [
28,
43] but also in benign intracranial tumors [
44]. Ducret et al. [
45] have shown that PRL induces a dose-dependent increase in proliferation and survival of U87-MG glioma cells. In line with these results we have detected PRLR mRNA expression in two additional glioma cell lines (G28 and G55) and could demonstrate that PRL stimulates cell proliferation in a dose-dependent manner, indicating that these cells express a functional PRLR.
Interestingly, we observed a strong up-regulation of PRLR in glioma cells treated with ES + Tum
in vitro. PRLR expression in contrast was not influenced by oxygen deprivation as observed after incubation of G55 cells under hypoxic and normoxic conditions for 24 hours, 48 hours and 5 days (data not shown). These observations suggest that up-regulation of PRLR in GBM tumors after ES + Tum treatment was not a secondary response to the anti-angiogenic treatment, but rather mediated through direct action of both integrin targeting factors on tumor cells. Although little is known about the effects of ES and Tum on glioma cells at the molecular level, an integrin-mediated auto-regulation of cell proliferation and apoptosis in glioma cells have been recently described by our group and others [
17,
34]. In addition, an integrin-PRLR cross-talk has recently been described in breast cancer cells [
46]. The fact that cilengitide, an integrin αvβ3/αvβ5 inhibitor, partially blocked ES + Tum-mediated effect on PRLR expression point to an integrin dependent mechanism. It is therefore tempting to speculate that the combined application of ES and Tum triggers up-regulation of PRLR in glioma, resulting in augmented PRL signalling and ultimately in increased tumor growth and/or stimulation of angiogenesis [
47]. Our
in vitro data confirm to some extent this hypothesis as they show for the first time that PRLR overexpression significantly increases glioma cell growth. The PRLR-mediated increase of cell growth was abrogated by inhibition of Jak2, a tyrosine kinase that has been described as major downstream regulator of PRLR-signalling [
28]. Moreover, we found a 4fold up-regulation of PRL expression in PRLR-overexpressing cells when compared to mock-transfected cells, suggesting a PRL-autocrine loop that stimulates glioma cell growth. Beside the already mentioned pro-proliferative activity of PRLR in diverse tumor entities, several groups have reported about a PRLR/PRL-mediated inhibition of apoptosis especially in response to chemotherapy. In breast cancer cells PRL confers resistance against cisplatin by activating a detoxification enzyme [
48,
49] and in ovarian carcinoma cells PRL and its receptor inhibit apoptosis induced by serum starvation or cisplatin treatment [
49]. These observations might explain the fact that ES + Tum-mediated cell growth inhibition
in vitro was significantly less pronounced in PRLR overexpressing cells than in control cells.
Material and methods
Expression vectors and transfection procedure
CMV (human cytomegalovirus) promoter driven plamids were used to generate expression vectors for angiogenic inhibitors. Murine ES was introduced into pcDNA3.1 plasmid (Invitrogen Life Technologies, Carlsbad, CA) as described previously [
32]. The cDNA coding for Tum was obtained by RT-PCR from total RNA extracted from HDMECs (human dermal microvascular endothelial cells) using following primer-pair: forward-primer 5′ccgagctcggatccaggtttgaaaggaaaa3′ and reverse-primer 5′cgctcgagggtgtcttttcatgcacacct3′, and was cloned into pSecTag2/Hygro (Invitrogen Life Technologies). The cDNA encoding full length PRLR together with a C-terminal HA-tag was cloned in pcDNA3.1(−)/Hygro plasmid (Invitrogen Life Technologies). The PRLR cDNA was obtained by RT-PCR from total RNA extracted from the cell line MCF-7 using following primer-pair: forward-primer 5′aacactcgagaaggcagccaacatgaaggaaaat3′ and reverse-primer: 5′tgggtaccttaagcgtaatctggaacatcgtatgggtagtgaaaggagtgtgt3′. Porcine aortic endothelial (PAE) cells were transfected with 2 μg plasmid DNA (ES, Tum), and the glioma cell line G55 with 1 μg plasmid DNA (PRLR) using Lipofectamine Plus® (Invitrogen Life Technologies) according to manufacturer instructions. Positive cells were selected by application of the appropriate antibiotics, and again expanded.
Cell culture and microencapsulation
Commercially available HUVECs and HDMECs were cultured in EGM-2 medium (Lonza, Basel, Switzerland) containing 2% fetal calf serum (FCS). PAE cells were maintained in F-12/HAM medium supplemented with 10% FCS. The human glioblastoma cell line G55 [
50] was kindly provided by Prof. Katrin Lamszus from the Department of Neurosurgery, University Hospital Hamburg-Eppendorf, and cultured in Modified Eagle’s Medium supplemented with 10% FCS. All cells were maintained in 5% CO
2/95% air atmosphere in a humidified incubator at 37°C. Wild-type or stably transfected PAE cells were encapsulated in Alginat microbeads as described previously [
22,
23]. Cells were resuspended in a 2% sodium alginate–saline solution (Pronova Ultra-Pure MVG; FMC BioPolymer AS d/b/a NovaMatrix, Sandvika, Norway) to a final concentration of 2 × 10
6 cells/ml. For
in vitro experiments conditioned medium (CM) was collected after a culture period of 48 hours. For long stimulation experiments CM was collected after a culture period of 4 days and subsequently diluted 1:3 with serum-reduced medium.
Cell viability and proliferation assay
HUVEC and G55 cells (5 × 104 per well) were seeded on 48-well tissue culture plates and incubated in basal medium or in CM or mixtures of CM from PAE-WT, PAE-Tum and PAE-ES cells additionally containing 4% FCS. Each stimulation experiment was performed in triplicate. After 24 and 48 hours of incubation at 37°C, cells were trypsinized and counted using the Vi-Cell XR (Beckman Coulter, Germany).
Cellular viability and proliferation was assessed using the WST-1 assay (Roche Applied Science, Mannheim, Germany) following the manufacturer´s instructions. Stably PRLR-transfected or mock-transfected G55-WT or G55 cells (8 × 103 cells per well) were cultured under serum deprivation in presence of AG490 (0.1, 1 and 10 μM) and/or 2 nM prolactin (R&D Systems, Minneapolis, MN). To quantify cell viability, cells were incubated with WST-1 reagent for 1 h and the absorbance was measured using a plate reader at 450 nm (reference 650 nm). Each stimulation experiment was performed in quintuplicate. Cell viability of experimental cells was related to cell viability of control (untreated) cells, which was set to 100%.
Apoptosis assay
G55 cells were seeded at subconfluent density into multiwell tissue culture plates. After culture overnight, cells were washed twice with PBS, and medium was replaced with CM from PAE-WT, PAE-Tum or PAE-ES cells, or a mixture of CM from PAE-Tum and PAE-ES cells. Incubations of cultures were continued for 24 hours before cells were analysed for apoptosis. For analysis, adherent cells were detached and pooled with floating cells. Apoptosis was assessed by flow cytometric analysis of cells stained with FITC-conjugated annexin-V and PI (BD Biosciences Pharmigen, San Diego, CA). Values represent the mean of three independent experiments.
Western blotting
Supernatants of transfected PAE cells were tested for transgene expression using Western blot analysis. CM from PAE-ES cells was incubated overnight at 4°C with heparin agarose (Sigma-Aldrich, St. Louis, MO) for protein concentration. Supernatant of PAE-Tum cells were concentrated overnight at 4°C by Nickel Cam™ HC resin (Sigma-Aldrich) according to manufacturer instructions. ES, Tum and PRLR were detected by murine ES polyclonal antibody (R&D Systems) [
24], His-probe polyclonal antibody (Santa Cruz Biothechnology, Santa Cruz, CA) [
24] and HA antibody (H6908; Sigma-Aldrich), respectively. The signal was visualized by Lumigen™ PS-3 detection reagent (GE Healthcare, Buckinghamshire, UK)) and exposed to an Amersham Hyperfilm™ECL (GE Healthcare).
In vitro wound healing assay
HDMEC cells (7× 105) were cultured in 24-well tissue culture plates in endothelial growth medium with supplements. After reaching confluence each well was scratched with a standardized pipette tip, resulting in an EC-free wound. Medium was replaced with CM of WT or transfected PAE cells additionally supplemented with 4% FCS. Photographs of each well were taken direct after scratching and after 20 hours incubation. The width of the gap was determined using AxioVision40 V4.8 software (Carl Zeiss Imaging Solutions, Jena, Germany) and values representing the closing wound were compared between experimental groups. Values represent the mean of three independent experiments.
In vivo tumor model
Animal experiments were conducted according to the UKCCR guidelines for the welfare of animals in experimental neoplasia [
51]. G55 cells (1× 106) were subcutaneously injected into SCID mice. Microbeads containing 1 × 106 WT or transfected PAE cells were implanted at the same site 7 days later. In the combination groups 1 × 106 PAE cells producing each inhibitor were injected. Each experimental group consisted of 5 animals. After 10 days, animals were sacrificed and tumors were excised and weighed. One half of each tumor was fixed in 10% formaldehyde and embedded in paraffin for immunohistochemistry. The other half was frozen in liquid nitrogen and used for RNA isolation.
Immunohistochemistry and immunofluorescence
Paraffin-embedded tissue samples were serially sectioned at a thickness of 5 μm, and every 20th section was used for analysis. Tissue sections were consecutively stained with hematoxylin and eosin. Blood vessels were visualized using murine polyclonal CD31 antibody [
52] (Dianova, Hamburg, Germany). Monoclonal Ki67 [
52], polyclonal prolactin receptor and M30 CytoDEATH antibodies were purchased from Dako (Dako Denmarck A/S, Glostrup, Denmark), abcam (Cambridge, UK) and Roche Applied Science respectively. Immunhistochemical staining was performed as previously described [
52]. For double immunhistochemical analyses, M30 and PRLR antibodies were visualized with Diaminobenzidine (DAB) and 3-Amino-9-ethylcarbazole (AEC), respectively. A blocking step in between using the Avidin-Biotin Blocking Kit (Vector Laboratories, Inc., Burlingame, CA) was performed. For immunofluorescence detection of PRLR in G55 cells, 3× 10
5 cells were seeded on chamber slides and treated with CM or mixtures of CM from PAE-WT, PAE-Tum and PAE-ES cells for 3 days. After fixation with cold ice methanol, staining was performed as previously described [
52]. Microvessel density was quantified by counting CD31-positive vessels in 10 arbitrarily chosen visual fields (10 × magnification) per tumor from totally 4 to 5 tumors from each experimental animal group using AxioVision40 V4.8 software (Carl Zeiss Imaging Solutions, Jena, Germany).
TUNEL assay of apoptotic cells
For the in situ detection of fragmented DNA, tissue sections were subjected to terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) using the in situ cell death detection kit, POD (Roche Diagnostic GmbH) according to the manufacturer´s instructions. Nuclei were counterstained with haematoxylin. TUNEL-negative nuclei were stained blue, while TUNEL-positive nuclei were stained brown.
RNA isolation and microarray analysis
Frozen tumor samples were homogenized with a micro tissue disintegrator. Tissue homogenates were first treated with Tri®Reagent (Sigma-Aldrich) for RNA Isolation succeeded by purification with the RNeasy Mini Kit (Qiagen, Hilden, Germany) following manufactures protocols. Quality and concentration of isolated RNA was determined using the Agilent RNA 6000 Nano Kit (Agilent Technologies Inc., Waldbronn, Germany) and NanoDrop6000 Photometer (Peqlab Biothechnologie GmbH, Erlangen, Germany). From each experimental animal group, three RNA samples were selected for further microarray analyses. Sense strand cDNA was generated from 100 ng total RNA using the Ambion WT Expression Kit (Ambion Inc., Applied Biosystems, Austin, TX). Procedures for labelling, fragmentation and hybridization were performed using the Terminal Labeling Kit and Hybridization, Wash and Stain Kit following Affymetrix protocols. All experiments were performed using Affymetrix Human Gene 1.0 ST Array containing 28.869 genes (Affymetrix Inc., Santa Clara, CA). Microarrays were scanned with the GeneChip Scanner 3000 7G (Affymetrix Inc.) using the GeneChip Command Console 3.0 (Affymetrix Inc.). The signals were processed using Genelevel RMA Sketch algorithm with following software: Affymetrix® Expression Console™ 1.1 software. Comparison analyses were carried out with a T-Test (between subjects) (TIGR, The Institute for Genomic Research, MeV 4).
Statistical analysis
Statistical analyses were performed using the Statistical Package for Social Sciences (SPSS) program (SPSS Inc. Chicago, IL, USA), version 15 with a Mann–Whitney-U-Test for tumor growth, microvessel density data and wound assays and with the unpaired Student t test with Welch correction for proliferation experiments. Probability value (p-value) ≤ 0.05 was considered statistically significant.
Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
LOF, JW, GS and WF designed this research work. LOF, JH and MK performed experiments and analysed data in this manuscript. LOF, JW, UB, EMMP, WF, CB and GS interpreted the data and wrote the manuscript. All authors read and approved the final manuscript.