γ-Actin plays a key role in endothelial cell motility and neovessel maintenance
© Pasquier et al.; licensee Biomed Central. 2015
Received: 17 September 2014
Accepted: 29 December 2014
Published: 6 February 2015
Angiogenesis plays a crucial role in development, wound healing as well as tumour growth and metastasis. Although the general implication of the cytoskeleton in angiogenesis has been partially unravelled, little is known about the specific role of actin isoforms in this process. Herein, we aimed at deciphering the function of γ-actin in angiogenesis.
Localization of β- and γ-actin in vascular endothelial cells was investigated by co-immunofluorescence staining using monoclonal antibodies, followed by the functional analysis of γ-actin using siRNA. The impact of γ-actin knockdown on the random motility and morphological differentiation of endothelial cells into vascular networks was investigated by timelapse videomicroscopy while the effect on chemotaxis was assessed using modified Boyden chambers. The implication of VE-cadherin, VEGFR-2 and ROCK signalling was then examined by Western blotting and using pharmacological inhibitors.
The two main cytoplasmic isoforms of actin strongly co-localized in vascular endothelial cells, albeit with some degree of spatial preference. While β-actin knockdown was not achievable without major cytotoxicity, γ-actin knockdown did not alter the viability of endothelial cells. Timelapse videomicroscopy experiments revealed that γ-actin knockdown cells were able to initiate morphological differentiation into capillary-like tubes but were unable to maintain these structures, which rapidly regressed. This vascular regression was associated with altered regulation of VE-cadherin expression. Interestingly, knocking down γ-actin expression had no effect on endothelial cell adhesion to various substrates but significantly decreased their motility and migration. This anti-migratory effect was associated with an accumulation of thick actin stress fibres, large focal adhesions and increased phosphorylation of myosin regulatory light chain, suggesting activation of the ROCK signalling pathway. Incubation with ROCK inhibitors, H-1152 and Y-27632, completely rescued the motility phenotype induced by γ-actin knockdown but only partially restored the angiogenic potential of endothelial cells.
Our study thus demonstrates for the first time that β-actin is essential for endothelial cell survival and γ-actin plays a crucial role in angiogenesis, through both ROCK-dependent and -independent mechanisms. This provides new insights into the role of the actin cytoskeleton in angiogenesis and may open new therapeutic avenues for the treatment of angiogenesis-related disorders.
KeywordsCytoskeleton Actin Angiogenesis Vascular endothelial cells ROCK signalling
Angiogenesis is defined as the formation of new blood vessels from pre-existing ones. It is crucial for organ growth during development but also throughout adult life to repair wounded tissues. Furthermore, an imbalance in this process directly contributes to numerous pathologies such as cancer, diabetes, age-related macular degeneration, ischemic disorders and rheumatoid arthritis [1,2]. The multi-step and complex process leading to the formation of a new vascular network relies on the activation of endothelial cells followed by their proliferation, migration and morphological differentiation into capillary tubes. The cytoskeleton which directly regulates and controls an impressive array of cell functions, including cell shape maintenance, cell division, vesicle and organelle transport, cell motility and differentiation, plays a major role in angiogenesis. Studies focusing on the anti-angiogenic properties of microtubule-targeting drugs – reviewed by Pasquier et al.  – have provided major insights into the role of microtubules in this process. However, very little is known about the specific role of actin isoforms in angiogenesis.
In vertebrates, there are 6 functional actin genes and the expression of the six actin isoforms is regulated both spatially and temporally in a tissue-specific manner. Four of these isoforms (i.e. α-cardiac muscle actin, α-skeletal muscle actin, α-smooth muscle actin and γ-smooth muscle actin) are mainly expressed in muscle cells, while the cytoplasmic isoforms β- and γ-actin are ubiquitous . Interestingly, the β- and γ-actin isoforms are almost identical proteins, differing only by 4 amino acid residues at the N-terminal end (positions 1, 2, 3 and 9). Distinct localization of β- and γ-actin mRNAs in several cell types, such as neurons, myoblasts and osteoblasts, has suggested for almost 20 years a spatial segregation of the two isoforms [5,6]. However, the spatial and functional segregation of β- and γ-actin was confirmed only recently in fibroblasts and epithelial cells by Chaponnier and colleagues, using newly developed monoclonal antibodies . In particular, β-actin appears to play a role in cell attachment and contraction by preferentially localizing to stress fibres whereas γ-actin is mainly organised as a meshwork in cortical and lamellipodial structures and thus plays a crucial role in cell motility . In accordance with this finding, we recently demonstrated that γ-actin specifically regulates cell motility by modulating the Rho-associated kinase (ROCK) signalling pathway and therefore influencing the phosphorylation of focal adhesion protein paxillin and myosin regulatory light chain 2 in neuroblastoma cells . Elsewhere, key functional differences between β- and γ-actin were also recently revealed by mouse knock-out experiments. Indeed, β-actin knock-out mice are not viable, in part due to severe growth and migration defects of β-actin null embryonic cells, which are not observed in γ-actin null embryonic cells . In contrast, γ-actin knock-out mice are viable, despite suffering increased mortality at birth and progressive hearing loss, which suggests that γ-actin is required for cytoskeleton maintenance but not for development . Spatial segregation and functional differences led us to hypothesize that β- and γ-actin may play distinct roles in endothelial cells and differentially contribute to angiogenesis. We therefore investigated the localization of β- and γ-actin in vascular endothelial cells and undertook the functional analysis of γ-actin by RNAi to decipher its specific function in endothelial cell adhesion, motility and morphological differentiation into vascular networks, thus revealing a key role in angiogenesis.
Material and methods
HMEC-1 endothelial cells were originally isolated from dermal microvessels and immortalized by transfection with SV40 large T antigen . They were obtained from the Cell Culture Laboratory in the Hôpital de la Conception (Assistance Publique Hôpitaux de Marseille, Marseille, France) and grown in MCDB-131 medium (Invitrogen, Mount Waverley, Australia) containing 10% heat-inactivated Fetal Calf Serum (FCS), 2 mM L-glutamine, 1% penicillin and streptomycin, 1 μg/mL hydrocortisone and 10 ng/mL epithelial growth factor (BioScientific, Gymea, Australia). BMH29L cells are bone marrow derived endothelial cells that were immortalized by ectopic expression of human telomerase reverse transcriptase . They were kindly provided by Dr Karen MacKenzie (Children’s Cancer Institute Australia) and grown in Medium 199 (Invitrogen) containing 10% heat-inactivated FCS, 5% male human serum AB only (Sigma-Aldrich, Castle Hill, Australia), 1% penicillin and streptomycin, 1% heparin, 5 ng/mL recombinant human FGFβ (fibroblast growth factor β; Sigma-Aldrich) and 20 μg/mL Endothelial Cell Growth Factor (ECGF; Roche, Dee Why, Australia). Both cell lines were routinely maintained in culture on 0.1% gelatin-coated flasks at 37°C and 5% CO2. Cell lines were regularly screened and are free from mycoplasma contamination.
γ-actin gene expression was silenced in endothelial cells using the siRNA sequence previously described (5′-AAGAGATCGCCGCGCTGGTCA-3′; Qiagen, Doncaster, Australia) . An alternative siRNA sequence (5′-CAGCAACACGTCATTGTGTAA-3′; Qiagen) was also used in confirmation experiments . β-actin gene expression was targeted using the siRNA sequence previously described (5′-AATGAAGATCAAGATCATTGC-3′; Qiagen) . The optimum amount of siRNA was determined to be 200 and 500 pmol for HMEC-1 and BMH29L cells, respectively and was used in all subsequent experiments. A non-silencing control siRNA, which has no sequence homology to any known human gene sequence, was used as a negative control in all experiments (Qiagen). Cells were transfected using the Nucleofector® II device (Lonza, Mount Waverley, Australia) as previously described . Briefly, HMEC-1 and BMH29L cells were resuspended in nucleofector® solution R and V, respectively, and transfected with siRNA using specifically optmized nucleofector® programs (T-016 and S-003 for HMEC-1 and BMH29L, respectively). All subsequent experiments were performed 72 h after siRNA transfection, when the level of γ-actin protein expression was the lowest.
The expression of γ-actin mRNA was examined using quantitative RT-PCR. Total RNA was extracted and DNase treated using the Qiagen RNeasy Plus kit according to the manufacturer’s instructions (Qiagen) and cDNA synthesis was performed using High capacity cDNA reverse transcription kit with RNAse inhibitor (Applied Biosystems, Mulgrave, Australia). Real-time PCR was performed on 7900HT Fast Real-time PCR system using the TaqMan® gene expression Master Mix (Applied Biosystems). γ-actin mRNA primer and probe sequences used were as follows: forward, 5′-CAGCTCTCGCACTCTGTTCTTC-3′; reverse, 5′-ACATGCCGGAGCCATTGT-3′; probe, 5′-CGCGCTGGTCATT-3′. All data were normalized to the housekeeping gene Ppia (peptidilprolyl isomerase A, TaqMan® Endogenous Control, Applied Biosystems). Gene expression levels were determined using the ΔΔC t method, normalized to the housekeeper gene and expressed relative to a calibrator .
Western blotting analysis
For western blotting analysis, cells were lysed in RIPA buffer containing a cocktail of protease and phosphatase inhibitors (Sigma-Aldrich). Equal amounts of protein (10–15 μg) were resolved on 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis or 4-15% pre-cast Criterion acrylamide gels (Bio-Rad Laboratories, Gladesville, Australia) before electrotransfer onto nitrocellulose membrane. Immunoblotting was performed using antibodies directed against β-actin (clone AC-74, Sigma-Aldrich), γ-actin – courtesy of Pr Peter Gunning , GAPDH (Abcam, Cambridge, UK), phospho-myosin light chain 2 (Cell Signaling Technology, Beverly, MA, USA), VE-cadherin (Cell signaling technology) and VEGFR-2 (Cell signaling technology). The membranes were then incubated with horseradish peroxidase-conjugated IgG secondary antibodies and protein detected with ECL Plus (GE Healthcare Life Sciences, Uppsala, Sweden). The blots were scanned and densitometric analysis performed as previously described .
Vascular endothelial cells were seeded on gelatin-coated 8-well Permanox Lab-Tek chamber slides (Applied Biosystems) after siRNA transfection. β-actin and γ-actin were stained as previously described  with slight modifications. Specifically, cells were fixed with 3.7% formaldehyde for 20 min at RT and permeabilized with 100% methanol for 20 min at −20°C. Cells were incubated with the following primary mAbs: anti-β-actin (mAb 4C2, IgG1 – courtesy of Pr Christine Chaponnier ) and anti-γ-actin (mAb 2A3, IgG2b – courtesy of Pr Christine Chaponnier ). The following secondary Abs were used: FITC-conjugated goat anti-mouse IgG1 (Southern Biotechnology, Birmingham, AL) and TRITC-conjugated goat anti-mouse IgG2b (Southern Biotechnology). For tubulin staining, cells were fixed and permeabilized in 100% methanol at 20°C for 15 min and blocked with 10% FCS for 30 min. Microtubules were then stained with anti-βI-tubulin primary antibody (Abcam), followed by Alexa Fluor 488 anti-mouse secondary antibody (Invitrogen). For paxillin and phalloidin dual staining, cells were fixed with 3.7% formaldehyde/PBS for 10 min and permeabilized with 0.1% Triton X-100/PBS for 5 min. Focal adhesions were then stained with anti-paxillin primary antibody (BD Biosciences), followed by Alexa Fluor 488 anti-mouse secondary antibody (Invitrogen) and Alexa 568-conjugated phalloidin (Invitrogen). All slides were mounted on coverslips with ProLong Gold anti-fade reagent containing DAPI (Invitrogen) and imaged using the 63X oil-immersion objective of an Axiovert 200 M fluorescent microscope coupled to an AxioCamMR3 camera driven by the AxioVision 4.8 software (Carl Zeiss, North Ryde, Australia). The thickness of actin stress fibres was determined performing a line scan perpendicular to the fibres using Image J, while the size of paxillin-containing adhesion sites was measured using the AxioVision 4.8 software.
Colocalization between β-actin and γ-actin channels was assessed by measuring the Pearson’s Correlation Coefficient and visually inspecting two-dimensional histograms (fluorograms). The Pearson’s coefficient measures the linear relationship between the pixel intensities of two channels. In the case of positive correlation this value can drop either due to decreasing colocalization, or due to differences in stoichiometry in structures. The fluorogram can distinguish these two scenarios, low colocalization shows by dispersion of points whilst varying stochiometries show multiple tight linear clusters. Measurements were made using the Coloc 2 plugin in ImageJ (imagej.nih.gov/ij). Image background was carefully subtracted from each channel and regions of interest were drawn around each cell to exclude extracellular pixels from the measurement. Measurements were performed on single cells to ensure variations in expression and staining did not contribute to multiple stoichiometries.
For the adhesion assay, cells were pre-labeled in situ with 10 μM Cell Tracker Green CMFDA (Invitrogen) in serum-free medium for 30 min and 50,000 cells were then seeded onto 24-well plates, pre-coated for 2 hours at 37°C with various extra-cellular matrix (ECM) proteins: fibronectin (2 μg/mL), laminin (10 μg/mL) or type I collagen (10 μg/mL). After 1 hour incubation, cells were washed twice with PBS and the number of adhered cells was assessed with a Victor 3 plate reader (Perkin-Elmer, Glen Waverley, Australia) at 492/517 (Abs/Em). All readings were then normalized to the negative control (no ECM).
The chemotaxis assay was performed as previously described . Briefly, the underside of 8 μm transparent polyethylene terephthalate membrane inserts (BD Falcon) was pre-coated with 0.1% gelatin for 1 h. The cells were pre-labeled in situ with 10 μM Cell Tracker Green CMFDA (Invitrogen) in serum-free medium for 30 min and 100,000 cells were then seeded onto the insert in assay medium (0.5% BSA in serum-free medium). Assay medium supplemented with 5% FCS, 0.1 ng/mL VEGF-A, 5 ng/mL FGFβ or 20 μg/mL ECGF was then added to the bottom of the insert and used as chemoattractant. A negative control was included in each experiment by adding serum-free medium to the bottom of the insert. The plates were incubated for 6 h at 37°C and 5% CO2. Excess cells on the upper side of the insert were then gently swabbed off with a cotton tip and migrated cells at the underside of the insert were measured with the same plate reader used for the adhesion assay. All readings were then normalized to the negative control (serum-free medium).
Random motility assay
Wound healing assay
An optimized wound healing assay was used as previously described , with slight modifications. Endothelial cells were grown to confluence in specific culture inserts (Ibidi, Martinsried, Germany). After 24 h, the culture inserts were removed, leaving a definite cell-free gap of approximately 400 μm, and the cells were washed with PBS before their incubation in culture medium. The colonization of the cell-free gap was analysed by time-lapse videomicroscopy using the 5X objective of the same microscope device used for immunofluorescence experiments. Photographs were taken every 10 minutes for 20 h and plates were kept at 37°C and 5% CO2 throughout the duration of the experiment. The migration rate was calculated digitally by quantification of the cell-free area at the different time points using the AxioVision 4.7 software.
In vitro Matrigel™ assay
Matrigel™ (BD Biosciences, North Ryde, Australia) assay was used to determine the effect of γ-actin knockdown on endothelial cell morphogenesis into capillary tubes, as previously described . Briefly, 24-well plates were coated at 4°C with 270 μL of a Matrigel™ solution (1:1 dilution in culture medium), which was then allowed to solidify for 1 h at 37°C before cell seeding. Cells were allowed to undergo morphogenesis and form capillary-like structures and photographs were taken after 8 h using the 5X objective of the same microscope device used for immunofluorescence experiments. Angiogenesis was then quantitatively evaluated by measuring the total surface area of capillary tubes formed in at least 10 view fields per well using the AxioVision 4.7 software.
A non-enzymatic methodology was also established to analyse the potential changes in protein expression that occur during the morphological differentiation of endothelial cells into vascular networks. Briefly, 3.2 × 105 cells were seeded onto 6-well plates previously coated with Matrigel™ and harvested at different time points of the morphological differentiation process (i.e. 15 min, 1, 2, 4 and 8 h). Cells were first incubated with a Cell Recovery Solution (BD Biosciences) for 1 h at 4°C under agitation to allow complete dissolution of the Matrigel™, then pelleted, washed with cold PBS and finally lysed as described in the western blotting section.
Rho-associated kinase (ROCK) signalling inhibition
ROCK signalling was interrupted as previously described , through the use of two specific ROCK inhibitors, H-1152 (Merck Millipore, Kilsyth, Australia) and Y-27632 (Sigma-Aldrich). Stock solutions of both inhibitors were prepared in water and stored at 4°C. Inhibitors (1–10 μM) were added to siRNA-transfected cells at 48 h post-transfection, and remained in culture medium for a further 24 h and during wound-healing and angiogenesis assays.
All experiments were performed at least in triplicate. Statistical significance was determined using two-sided student’s t test in the GraphPad Prism 4 software (GraphPad Software, Inc).
Spatial distribution of β- and γ-actin in vascular endothelial cells
Knockdown of cytoplasmic γ-actin expression by RNAi
Cytoplasmic γ-actin expression is essential for the morphological differentiation of endothelial cells into vascular networks
Vascular regression induced by γ-actin knockdown is associated with impaired VE-cadherin up-regulation
Cytoplasmic γ-actin plays a key role in endothelial cell motility and chemotaxis
Effects of γ-actin knockdown on random motility parameters
Average cell velocity (μm/min)
0.53 ± 0.03
0.37 ± 0.03
−30 ± 5 %**
Persistence time (min)
6.3 ± 0.3
8.6 ± 0.6
+36 ± 7 %*
Random motility coefficient (μm2/min)
0.90 ± 0.05
0.55 ± 0.07
−38 ± 8 %**
In order to further investigate the effects of γ-actin knockdown on endothelial cell migration, wound healing experiments were performed using time-lapse videomicroscopy. Figure 6D shows representative photographs taken from control (top panel) and γ-actin siRNA-treated cells (bottom panel) after 12 h incubation. At this time point, recovery from the wound was 93.0 ± 3.4% and 67.2 ± 3.5% in control and γ-actin siRNA-treated cells, respectively (p < 0.001). As shown in Figure 6E, γ-actin knockdown resulted in a significant decrease in wound recovery at all time points until 15 h. Linear regression analysis showed that control siRNA-treated BMH29L cells reached 50% wound recovery after 6.7 ± 0.4 h, whereas it took 9.4 ± 0.6 h (p < 0.01) to γ-actin siRNA-treated BMH29L cells (data not shown). Similar results were obtained with HMEC-1 cells, which took 7.8 ± 0.5 h and 9.8 ± 0.5 h to reach 50% wound recovery (p < 0.01), when they were transfected with control and γ-actin siRNA, respectively (data not shown).
Cytoplasmic γ-actin regulates endothelial cell motility through ROCK signalling pathway
Cytoplasmic γ-actin regulates angiogenesis through ROCK-dependent and ROCK-independent mechanisms
Actin-binding and regulatory proteins have been proposed to represent attractive targets for the development of innovative anti-angiogenic therapies . However, the role of the actin cytoskeleton in angiogenesis has not been completely elucidated. Basic cell biology studies are crucially needed to map the specific role played by each component of the actin cytoskeleton in order to find new ways to target angiogenesis in pathologies such as cancer and rheumatoid arthritis. Here, we investigated the cellular distribution and localization of the two cytoplasmic isoforms of actin, β and γ, in vascular endothelial cells and performed the first functional analysis of γ-actin in these cells, revealing a key role for this protein in angiogenesis.
Co-immunofluorescence staining using newly developed antibodies directed against β- and γ-actin revealed a strong colocalization of the two cytoplasmic actin isoforms in vascular endothelial cells, albeit some level of spatial preference. Our findings are in accordance with the preferential localization of β-actin in radial stress fibres and membrane ruffling and γ-actin in the cortical microfilament meshwork previously reported in epithelial cells, fibroblasts  and more recently in neuroblastoma cells . The subcellular localization of β- and γ-actin expression appears to be mediated by localization of their respective mRNA . Furthermore, studies have shown that β- and γ-actin differentially interact with accessory proteins such as tropomyosins  and non-sarcomeric myosins . These differential interactions may govern the specific functions of β- and γ-actin, and therefore contribute to the spatial and temporal regulation of cytoskeletal dynamics.
Significant knockdown of β-actin could not be achieved in vascular endothelial cells without inducing major cytotoxicity, suggesting that these cells are particularly sensitive to changes in β-actin expression. In contrast, γ-actin expression could be successfully knocked down by 50-60% in endothelial cells, indicating an isoform-specific function for actin in endothelial cells. This difference in response to β- and γ-actin knockdown is consistent with recent studies showing that β-actin knockout mice are not viable  unlike γ-actin knockout mice, which despite suffering increased mortality and showing progressive hearing loss during adulthood, are viable .
Interestingly, γ-actin knockdown almost completely suppressed the capacity of endothelial cells to maintain capillary-like structures on Matrigel™. Timelapse videomicroscopy analysis revealed that γ-actin was dispensable in the early steps of the morphological differentiation process (i.e. vascular sprouting) but required to sustain newly formed vascular networks. Previous studies have shown that the major endothelial cell adhesion molecule, VE-cadherin, plays a crucial role in angiogenesis – reviewed in [25-27]. The development of a VE-cadherin knockout (Cdh5−/−) mouse model thus revealed that this type II cadherin is dispensable in de novo blood vessel formation but essential to prevent the regression and ensure the maturation of nascent blood vessels [28,29]. We therefore hypothesized that VE-cadherin may be involved in the vascular regression induced by γ-actin knockdown. Our results showed that the expression of VE-cadherin is rapidly up-regulated during the morphological differentiation of endothelial cells, which is consistent with the study by Prandini et al. demonstrating that VE-cadherin promoter activity is enhanced during angiogenesis . Furthermore, we found that the up-regulation of VE-cadherin was impaired in γ-actin siRNA-treated cells, which may contribute to vascular regression in these cells. This finding is in agreement with a recent study showing that VE-cadherin is involved in the cessation of endothelial cell sprouting . Elsewhere, recent studies have shown that the major receptor involved in angiogenesis, VEGFR-2, may be in part regulated by the actin cytoskeleton. For instance, its localization to the plasma membrane has been shown to be mediated by myosin 1C, an unconventional motor protein trafficking along actin filaments . In addition, inhibition of Rho GTPase Rac1 was found to down-regulate the expression VEGFR-2 . However, we demonstrated herein that VEGFR-2 expression was up-regulated in endothelial cells during the angiogenic process independently of γ-actin expression.
To better understand how γ-actin knockdown impaired the angiogenic process, we investigated its impact on endothelial cell adhesion, chemotaxis and motility. While partially depleting γ-actin expression did not alter cell adhesion, it significantly reduced the capacity of endothelial cells to respond to chemotactic stimuli, randomly explore their microenvironment and colonize a new area. This result is in line with other studies demonstrating the crucial role played by the actin cytoskeleton in cell locomotion in non-endothelial cells – reviewed by Le Clainche and Carlier . It is also consistent with the results recently reported for γ-actin depleted epithelial cells , arguing in favour of a crucial role of γ-actin in the control of directional motility in diverse cell types.
We previously reported that γ-actin regulates neuroblastoma cell migration by modulating ROCK signalling pathway . Here, we extended this finding to vascular endothelial cells. Indeed, γ-actin knockdown was associated with accumulation of thicker actin stress fibres, larger focal adhesions and increased phosphorylation of myosin light chain 2 in endothelial cells, thus strongly suggesting activation of the ROCK signalling pathway. Furthermore, incubation with pharmacological inhibitors of ROCK signalling completely restored the motility of γ-actin siRNA-treated cells, demonstrating that the γ-actin knockdown motility phenotype is dependent upon ROCK activation. In contrast, ROCK inhibitors only partially rescued the angiogenic potential of γ-actin knockdown cells, suggesting that γ-actin regulates angiogenesis through both ROCK-dependent and ROCK-independent mechanisms.
Interestingly, VE-cadherin engagement has been shown to activate numerous biochemical pathways that can affect cell shape and adhesion, including the ROCK signalling pathway [35-37]. Furthermore, a direct link was recently reported between actin stress fibres and the protein complex forming adherens junctions, VE-cadherin/β-catenin/α-catenin [38,39]. Here we provide evidence of the reverse relationship since decreasing the expression of γ-actin partially blocked the required up-regulation of VE-cadherin expression during morphological differentiation of endothelial cells. Additional experiments are currently underway to fully decipher the mechanisms involved in the crosstalk between γ-actin, ROCK signalling and VE-cadherin, which appears to play a critical role in the maintenance of newly formed vascular networks.
Collectively our results demonstrate for the first time that γ-actin plays a crucial role in angiogenesis through the regulation of endothelial cell motility and neovessel maintenance, via ROCK-dependent and -independent mechanisms. This finding opens potential therapeutic avenues for the treatment of angiogenesis-related disorders by targeting key factors involved in the γ-actin/VE-cadherin/ROCK signalling network.
Fetal calf serum
- FGFβ :
Fibroblast growth factor β
Human microvascular endothelial cell line 1
Myosin light chain 2
Small interfering RNA
Rho-associated protein kinase
Vascular endothelial cadherin
Vascular endothelial growth factor receptor 2
The authors would like to thank Dr Christine Chaponnier (University of Geneva, Switzerland) for generously providing the β- and γ-actin monoclonal antibodies and Dr Karen MacKenzie (Children’s Cancer Institute Australia) for providing the BMH29L endothelial cell line. This work was supported by the Children’s Cancer Institute Australia for Medical Research, which is affiliated with the University of New South Wales and Sydney Children’s Hospital, and by grants from the Cancer Institute New South Wales (EP) and Cancer Council New South Wales (MK). EP was supported by a Cancer Institute New South Wales “Early Career Development” Fellowship. MK is supported by a NHMRC Senior Research Fellowship (APP1058299) and an Australian Research Council Centre of Excellence in Convergent Bio-Nano Science and Technology.
- Carmeliet P. Angiogenesis in life, disease and medicine. Nature. 2005;438:932–6.View ArticlePubMedGoogle Scholar
- Ferrara N, Kerbel RS. Angiogenesis as a therapeutic target. Nature. 2005;438:967–74.View ArticlePubMedGoogle Scholar
- Pasquier E, Andre N, Braguer D. Targeting microtubules to inhibit angiogenesis and disrupt tumour vasculature: implications for cancer treatment. Curr Cancer Drug Targets. 2007;7:566–81.View ArticlePubMedGoogle Scholar
- Tondeleir D, Vandamme D, Vandekerckhove J, Ampe C, Lambrechts A. Actin isoform expression patterns during mammalian development and in pathology: insights from mouse models. Cell Motil Cytoskeleton. 2009;66:798–815.View ArticlePubMedGoogle Scholar
- Hill MA, Gunning P. Beta and gamma actin mRNAs are differentially located within myoblasts. J Cell Biol. 1993;122:825–32.View ArticlePubMedGoogle Scholar
- Bassell GJ, Zhang H, Byrd AL, Femino AM, Singer RH, Taneja KL, et al. Sorting of beta-actin mRNA and protein to neurites and growth cones in culture. J Neurosci. 1998;18:251–65.PubMedGoogle Scholar
- Dugina V, Zwaenepoel I, Gabbiani G, Clement S, Chaponnier C. Beta and gamma-cytoplasmic actins display distinct distribution and functional diversity. J Cell Sci. 2009;122:2980–8.View ArticlePubMedGoogle Scholar
- Shum MS, Pasquier E, Po’uha ST, O’Neill GM, Chaponnier C, Gunning PW, et al. gamma-Actin regulates cell migration and modulates the ROCK signaling pathway. FASEB J. 2011;25:4423–233.View ArticlePubMedGoogle Scholar
- Bunnell TM, Burbach BJ, Shimizu Y, JM E. β-Actin specifically controls cell growth, migration, and the G-actin pool. Mol Biol Cell. 2011;22:4047–58.View ArticlePubMed CentralPubMedGoogle Scholar
- Belyantseva IA, Perrin BJ, Sonnemann KJ, Zhu M, Stepanyan R, McGee J, et al. Gamma-actin is required for cytoskeletal maintenance but not development. Proc Natl Acad Sci U S A. 2009;106:9703–8.View ArticlePubMed CentralPubMedGoogle Scholar
- Ades EW, Candal FJ, Swerlick RA, George VG, Summers S, Bosse DC, et al. HMEC-1: establishment of an immortalized human microvascular endothelial cell line. J Invest Dermatol. 1992;99:683–90.View ArticlePubMedGoogle Scholar
- MacKenzie KL, Franco S, Naiyer AJ, May C, Sadelain M, Rafii S, et al. Multiple stages of malignant transformation of human endothelial cells modelled by co-expression of telomerase reverse transcriptase, SV40 T antigen and oncogenic N-ras. Oncogene. 2002;21:4200–11.View ArticlePubMedGoogle Scholar
- Verrills NM, Po’uha ST, Liu ML, Liaw TY, Larsen MR, Ivery MT, et al. Alterations in gamma-actin and tubulin-targeted drug resistance in childhood leukemia. J Natl Cancer Inst. 2006;98:1363–74.View ArticlePubMedGoogle Scholar
- Harborth J, Elbashir SM, Bechert K, Tuschl T, Weber K. Identification of essential genes in cultured mammalian cells using small interfering RNAs. J Cell Sci. 2001;114:4557–65.PubMedGoogle Scholar
- Pasquier E, Tuset MP, Street J, Sinnappan S, MacKenzie KL, Braguer D, et al. Concentration- and schedule-dependent effects of chemotherapy on the angiogenic potential and drug sensitivity of vascular endothelial cells. Angiogenesis. 2012;16:373–86.View ArticlePubMed CentralPubMedGoogle Scholar
- Winer J, Jung CK, Shackel I, Williams PM. Development and validation of real-time quantitative reverse transcriptase-polymerase chain reaction for monitoring gene expression in cardiac myocytes in vitro. Anal Biochem. 1999;270:41–9.View ArticlePubMedGoogle Scholar
- Schevzov G, Vrhovski B, Bryce NS, Elmir S, Qiu MR, O’Neill GM, et al. Tissue-specific tropomyosin isoform composition. J Histochem Cytochem. 2005;53:557–70.View ArticlePubMedGoogle Scholar
- Pasquier E, Sinnappan S, Munoz MA, Kavallaris M. ENMD-1198, a new analogue of 2-methoxyestradiol, displays both antiangiogenic and vascular-disrupting properties. Mol Cancer Ther. 2010;9:1408–18.View ArticlePubMedGoogle Scholar
- Kouvroukoglou S, Dee KC, Bizios R, McIntire LV, Zygourakis K. Endothelial cell migration on surfaces modified with immobilized adhesive peptides. Biomaterials. 2000;21:1725–33.View ArticlePubMedGoogle Scholar
- Valentiner U, Haane C, Nehmann N, Schumacher U. Effects of bortezomib on human neuroblastoma cells in vitro and in a metastatic xenograft model. Anticancer Res. 2009;29:1219–25.PubMedGoogle Scholar
- Adler J, Parmryd I. Quantifying colocalization by correlation: the Pearson correlation coefficient is superior to the Mander’s overlap coefficient. Cytometry A. 2010;77:733–42.View ArticlePubMedGoogle Scholar
- Thoenes L, Gunther M. Novel approaches in anti-angiogenic treatment targeting endothelial F-actin: a new anti-angiogenic strategy? Curr Opin Mol Ther. 2008;10:579–90.PubMedGoogle Scholar
- Schevzov G, Lloyd C, Hailstones D, Gunning P. Differential regulation of tropomyosin isoform organization and gene expression in response to altered actin gene expression. J Cell Biol. 1993;121:811–21.View ArticlePubMedGoogle Scholar
- Müller M, Diensthuber RP, Chizhov I, Claus P, Heissler SM, Preller M, et al. Distinct functional interactions between actin isoforms and nonsarcomeric myosins. PLoS One. 2013;8:e70636.View ArticlePubMed CentralPubMedGoogle Scholar
- Cavallaro U, Liebner S, Dejana E. Endothelial cadherins and tumor angiogenesis. Exp Cell Res. 2006;312:659–67.View ArticlePubMedGoogle Scholar
- Lampugnani MG, Dejana E. Adherens junctions in endothelial cells regulate vessel maintenance and angiogenesis. Thromb Res. 2007;120:S1–6.View ArticlePubMedGoogle Scholar
- Vestweber D. VE-cadherin: the major endothelial adhesion molecule controlling cellular junctions and blood vessel formation. Arterioscler Thromb Vasc Biol. 2008;28:223–32.View ArticlePubMedGoogle Scholar
- Carmeliet P, Lampugnani MG, Moons L, Breviario F, Compernolle V, Bono F, et al. Targeted deficiency or cytosolic truncation of the VE-cadherin gene in mice impairs VEGF-mediated endothelial survival and angiogenesis. Cell. 1999;98:147–57.View ArticlePubMedGoogle Scholar
- Crosby CV, Fleming PA, Argraves WS, Corada M, Zanetta L, Dejana E, et al. VE-cadherin is not required for the formation of nascent blood vessels but acts to prevent their disassembly. Blood. 2005;105:2771–6.View ArticlePubMedGoogle Scholar
- Prandini MH, Dreher I, Bouillot S, Benkerri S, Moll T, Huber P. The human VE-cadherin promoter is subjected to organ-specific regulation and is activated in tumour angiogenesis. Oncogene. 2005;24:2992–3001.View ArticlePubMed CentralPubMedGoogle Scholar
- Abraham S, Yeo M, Montero-Balaguer M, Paterson H, Dejana E, Marshall CJ, et al. VE-cadherin-mediated cell-cell interaction suppresses sprouting via signalling to MLC2 phosphorylation. Curr Biol. 2009;19:668–74.View ArticlePubMedGoogle Scholar
- Tiwari A, Jung JJ, Inamdar SM, Nihalani D, Choudhury A. The myosin motor Myo1c is required for VEGFR2 delivery to the cell surface and for angiogenic signalling. Am J Physiol Heart Circ Physiol. 2013;304:H687–96.View ArticlePubMed CentralPubMedGoogle Scholar
- Meissner M, Michailidou D, Stein M, Hrgovic I, Kaufmann R, Gille J. Inhibition of Rac1 GTPase downregulates vascular endothelial growth factor receptor-2 expression by suppressing Sp1-dependent DNA binding in human endothelial cells. Exp Dermatol. 2009;18:863–9.View ArticlePubMedGoogle Scholar
- Le Clainche C, Carlier MF. Regulation of actin assembly associated with protrusion and adhesion in cell migration. Physiol Rev. 2008;88:489–513.View ArticlePubMedGoogle Scholar
- Lampugnani MG, Zanetti A, Breviario F, Balconi G, Orsenigo F, Corada M, et al. VE-cadherin regulates endothelial actin activating Rac and increasing membrane association of Tiam. Mol Biol Cell. 2002;13:1175–89.View ArticlePubMed CentralPubMedGoogle Scholar
- Kouklis P, Konstantoulaki M, Malik AB. VE-cadherin-induced Cdc42 signaling regulates formation of membrane protrusions in endothelial cells. J Biol Chem. 2003;278:16230–6.View ArticlePubMedGoogle Scholar
- Nelson CM, Pirone DM, Tan JL, Chen CS. Vascular endothelial-cadherin regulates cytoskeletal tension, cell spreading, and focal adhesions by stimulating RhoA. Mol Biol Cell. 2004;15:2943–53.View ArticlePubMed CentralPubMedGoogle Scholar
- Millan J, Cain RJ, Reglero-Real N, Bigarella C, Marcos-Ramiro B, Fernández-Martín L, et al. Adherens junctions connect stress fibres between adjacent endothelial cells. BMC Biol. 2010;8:11.View ArticlePubMed CentralPubMedGoogle Scholar
- Noda K, Zhang J, Fukuhara S, Kunimoto S, Yoshimura M, Mochizuki N. Vascular endothelial-cadherin stabilizes at cell-cell junctions by anchoring to circumferential actin bundles through alpha- and beta-catenins in cyclic AMP-Epac-Rap1 signal-activated endothelial cells. Mol Biol Cell. 2010;21:584–96.View ArticlePubMed CentralPubMedGoogle Scholar
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly credited. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.