Electric field stimulation

The experimental setup and EF exposure protocols were similar to those reported previously (Zhao et al., 1996). In brief, HUVEC cells were seeded in a specially made trough formed by two parallel (1 cm apart) strips of glass coverslip (No. 1, length of 22 mm) fixed to the base of the dish with silicone grease (Dow Corning, MS4). Scratch lines were made perpendicular to the long axis of chambers with a fine sterile needle and used as reference marks for directed cell migration. The cells were incubated for 24–48 hours (37°C, 5% CO₂), allowing them to settle and adhere to the base of the dish, before a roof of No. 1 coverslip was applied and sealed with silicone grease. The final dimensions of the chamber, through which current was passed, were 22×10×0.2 mm. Agar-salt bridges not less than 15 cm long were used to connect silver/silver-chloride electrodes in beakers of Steinberg’s solution (58 mM NaCl, 0.67 mM KCl, 0.44 mM Ca(NO₃)₂, 1.3 mM MgSO₄, 4.6 mM Trizma base, pH 7.8–8.0), to pools of excess culture medium at either side of the chamber. This prevents diffusion of electrode products into the cultures. Field strengths were measured directly at the beginning of, the end of and during each experiment. No fluctuations in field strength were observed. For drug inhibition experiments, the cells were incubated with the VEGFR inhibitor 4-[(4′-chloro-2′-fluoro)phenylamino]-6,7-dimethoxyquinazoline (50 μM), the PI3K inhibitor LY294002 (50 μM), an Akt inhibitor 1-L-6-hydroxymethyl-chiro-inositol 2-[(R)-2-O-methyl-3-O-octadecylcarbonate] (50 μM), the Rho kinase inhibitor Y27632 (50 μM), both Akt and Rho kinase inhibitors (10 μM each) or latrunculin (50 nM) for 1 hour before EF stimulation. The same concentration of drug was present during EF exposure in a CO₂ incubator.

Reorientation of endothelial cells in EFs requires VEGFR activation

VEGF activation is one of the pivotal elements in angiogenic responses and enhanced angiogenesis by electric stimulation in vivo is mediated through VEGFR activation (Kanno et al., 1999; Patterson et al., 1999; Linderman et al., 2000). To test whether EF-induced endothelial cell orientation might involve VEGF signaling, we quantified levels of VEGF. EF exposure (200 mV mm⁻¹, the same as that measured at skin wounds) significantly enhanced levels of VEGF released into the culture medium. Marked elevation of VEGF in the culture medium was observed as early as 5 minutes after onset of the EF; this was reduced at 1 hour and 2 hours, rose again at 4 hours, and reached a high level by 24 hours (Fig. 4).

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Fig. 4

Exposure to a physiological EF increased VEGF release from endothelial cells. HUVEC cells were cultured in serum-free DMEM and exposed to an EF of 200 mV mm⁻¹. VEGF in the medium was quantified by ELISA. Values are means±s.e.m.

Inhibition of VEGFR activation by inhibiting both VEGFR-1 and VEGFR-2 with the drug 4-[(4′-chloro-2′-fluoro)phenylamino]-6,7-dimethoxyquinazoline completely abolished the reorientation of cells in an EF (Fig. 3B). This drug is a potent VEGFR inhibitor that inhibits the receptor tyrosine kinase activity (50% inhibitory concentrations of 2.0 μM and 100 nM for VEGFR-1 and VEGFR-2, respectively). It is very selective for VEGFR-1 and VEGFR-2 tyrosine kinase activity compared with that associated with the epidermal growth factor (EGF) receptor (50-fold and 3800-fold, respectively) (Hennequin et al., 1999). The morphology of the cells treated with VEGFR inhibitor was very similar to control cells. Cells still elongated, although their long axis was slightly reduced, but they were oriented randomly (Fig. 1C). Inhibition of VEGFRs could conceivably have had detrimental effects on the long-term viability of cells and this could have influenced their orientation responses. To test for this, we compared the orientation response after a short period of inhibitor and EF application. The orientation response was completely abolished at 4 hours and 8 hours in an EF after VEGFR inhibition. The Oi values of the cells treated with VEGFR inhibitor were −0.16±0.05 and −0.05±0.05 in EF for 4 hours and 8 hours, respectively, which is significantly different from the non-inhibitor-treated values of 0.36±0.05 and 0.53±0.05 (P<0.01).

Reorientation of endothelial cells involved the PI3K-Akt pathway

VEGFR activation leads to endothelial cell migration, cell survival and proliferation, which require the activation of Akt, a downstream effectors of PI3K (Shiojima and Walsh, 2002). Both the PI3K inhibitor LY294002 (50 μM) and the Akt inhibitor (50 μM) significantly decreased the orientation response (Fig. 3B). The concentration of either drug alone would be expected to inhibit PI3K and Akt activation completely but neither drug inhibited perpendicular reorientation completely, and significant Oi values remained (Fig. 3B), indicating that other signaling mechanisms must be involved.

Role of Rho and integrin in EF-induced reorientation of endothelial cells

The Rho family of GTPases regulates VEGF-stimulated endothelial cell motility and reorganization of the actin cytoskeleton, which are important in endothelial cell retraction and in the formation of intercellular gaps (Wojciak-Stothard et al., 1998). The Rho kinase inhibitor, Y27632, decreased the orientation response significantly, with Oi values of 0.55±0.05, 0.45±0.05 and 0.24±0.05 at 10 μM, 20 μM and 50 μM, respectively. Significant Oi values nonetheless remained even at 50 μM, indicating that multiple signaling mechanisms must be involved. Mitogen-activated-protein kinase inhibition with U0126 (50 μM), like Y27632 (0.33±0.03), decreased the orientation to a similar extent.

Because both Akt and Rho kinase inhibitors individually showed partial inhibition, perhaps the two enzymes function in different pathways to induce cell reorientation. To test this, a combination of the two inhibitors was used. The orientation response was abolished completely by using Akt and Rho kinase inhibitors together (both at 10 μM) (Oi=−0.10±0.06; compared to control=0.80±0.09, P<0.0001) (Fig. 3B).

Integrins, especially αvβ3, are important in endothelial cell movement and alignment to shear stress and mechanical stimulation (Davies, 1991; Davies, 1995; Tzima et al., 2001). HUVEC cells were incubated with a blocking antibody against αvβ3 (LM609) (20 μg ml⁻¹) for 1 hour and then exposed to an EF (200 mV mm⁻¹) with the antibody present. Blocking αvβ3 had no effect on orientation to the EF, cells reoriented normally (Oi=0.72±0.03, n=110, compared with the control=0.80±0.09, n=124, P>0.05).

Small EFs elongated endothelial cells

HUVEC cells elongated dramatically in an EF (Fig. 1B, Fig. 2). By contrast, cells cultured with no EF retained a more-cobblestone-like appearance (Fig. 1A). Striking cell elongation was induced by a voltage drop of about 0.7–4.0 mV across a cell of ~15 μm in diameter. We quantified the elongation of the cells using a long:short axis ratio. A perfectly round cell has a long:short axis ratio of 1 and, as cells elongate, the ratio increases. Control cells (no EF) showed no increase in long:short axis over 24 hours in culture (Fig. 5A). Elongation responses were both time and voltage dependent. The long:short axis ratio of EF exposed cells indicated gradual cell elongation throughout the 24 hour experimental period. The voltage dependency of the elongation response was more obvious at later times, with a greater long:short axis ratio for cells cultured at higher EFs (Fig. 2, Fig. 5A). The threshold for EF-induced endothelial cell elongation was between 50–75 mV mm⁻¹ (Fig. 5A), again 0.5–0.75 mV across a cell 10 μm in diameter. The elongation response of endothelial cells was more marked than that seen previously at the same EF strengths, in corneal and lens epithelial cells (Zhao et al., 1996).

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Fig. 5

Time and voltage dependency of elongation of endothelial cells in a small physiological EF. (A) Cell elongation as a function of time and voltage (n=65–598 from at least two independent experiments). (B) Effects of various drugs on EF-induced cell elongation. Inhibition of PI3K (LY), Akt (Akt-i), Rho (Y27632) and actin polymerization (Latrunc) significantly decreased the elongation response, as did inhibition of VEGR (VEGFR-i), and the combined inhibition of both Akt and Rho (Akt-i + Y). VEGFR-i, VEGFR inhibitor (50 μM); LY, PI3K inhibitor LY294002 (50 μM); Akt-i, Akt inhibitor (50 μM); Y27632, Rho inhibitor (50 μM); Akt-i + Y27632, Akt and Rho inhibitors (10 μM each); Latrunc, Latrunculin (50 μM). Cells were subjected to EFs of 200 mV mm⁻¹ for 24 hours. n=47–343 from more than two independent experiments. *, P<0.002 compared with cells exposed to 200 mV mm⁻¹ without drug treatment; **, P<0.0001 compared with cells exposed to 200 mV mm⁻¹ without drug treatment; #, P<0.001 compared with same drug treatment but not exposed to EFs; ##, P<0.0001 compared with same drug treatment but not exposed to EFs.

VEGFR, PI3K-Akt and Rho signaling are involved in the elongation response

The signaling elements required for reorientation are also involved in elongation, but there are subtle differences. The VEGFR inhibitor (50 μM) had no effect on the long:short axis ratio of control cells but significantly decreased the long:short axis ratio in EF-treated cells (P<0.002, Fig. 5A). Both the PI3K inhibitor LY294002 and the Akt inhibitor also significantly decreased the long:short axis ratio (both P<0.0001 versus control; Fig. 5A). Cells treated with these drugs elongated less, with LY294002 the more effective in suppressing EF-induced elongation. The Rho kinase inhibitor, Y27632 also significantly decreased the long:short axis ratio (P<0.0001, Fig. 5B), whereas the αvβ3-blocking antibody significantly inhibited the elongation response (3.12±0.008 compared with the control 3.65±0.15, P=0.007).

Cytoskeleton alignment and the consequence of actin filament disruption

To control changes in cell shape, reorientation and migration, extracellular stimuli initiate intracellular signaling that modifies cytoskeletal organization. Both actin filaments and microtubules were aligned in the direction of cell elongation (Fig. 1D-I). Latrunculin A, a toxin inhibiting actin polymerization, completely abolished the EF-induced elongation response (Fig. 5B) and suppressed the orientation response significantly (P<0.001) but not fully (Fig. 3B).

DC EFs have direct effects on endothelial cells

Electrical stimulation is a promising approach for therapeutic angiogenesis (Kanno et al., 1999; Patterson et al., 1999). Weak pulsatile electric stimulation caused significant increases in blood flow and capillary density in the ischemic limb of rat because it stimulated VEGF secretion from muscle cells in vitro and in vivo. Muscle contraction was not needed. Our results suggest that, in addition, there could be direct effects of this type of electrical stimulation on endothelial cells. We have shown that a small physiological EF markedly stimulated release of VEGF from cultured endothelial cells and that the ensuing activation of VEGFRs and associated signaling elements promoted cell elongation, cell reorientation and directed cell migration, all of which are central to angiogenesis. It is crucially important that angiogenesis is highly controlled and organized spatially. Currently, chemical gradients are the major players thought to guide new blood vessel growth. Our data therefore introduce the DC EF as a novel, extracellular, perhaps physiological cue orchestrating endothelial cell behaviors in culture. This single cue has a remarkable series of direct effects, which collectively are precursors of new vessel formation in vivo.

DC electric fields stimulate VEGF production in endothelial cells

Pulsed DC electrical stimulation (10 Hz, 300 microseconds) induces VEGF expression by muscle cells through increasing VEGF-encoding mRNA at a transcriptional level, which started to rise 12 hours after EF stimulation and peaked at 24–48 hours. This response might play a role in sensing tissue hypoxia following excessive muscle activation (Hang et al., 1995). We showed that a constant DC EF induced significant VEGF production by endothelial cells within 5 minutes after the onset of an EF and showed a first peak after 30 minutes. This is too early to result from VEGF transcription de novo and indicates that the EF must increase VEGF secretion. By contrast, the later elevation of VEGF at 24 hours in a DC EF (Fig. 4) could be due to new VEGF synthesis. This difference in VEGF production might be due to the differences in stimulation modalities (pulsed DC or constant DC) and cell types (muscle cells or endothelial cells). The early rise of VEGF within the immediate vicinity of endothelial cells is necessary and causes the initiation of the behavioral responses of endothelial cells.

VEGFR, PI3K-Akt and Rho kinase signaling in EF-induced responses

The proximal element that transduces the EF and induced the pre-angiogenic responses is the VEGFR with downstream signaling elements involving PI3K-Akt, Rho-ROCK and the F-actin cytoskeleton. Our finding that Rho-p160ROCK is involved in EF-induced cell alignment is in keeping with a similar role in cell alignment in shear stress (Li et al., 1999). Apart from its involvement in Ca²⁺ regulation, p160ROCK or Rho kinase can enhance the phosphorylation of myosin light chain (Kimura et al., 1996). A small DC EF might activate the Rho-p160ROCK pathway, which in turn modulates myosin light chain phosphorylation, or intracellular Ca²⁺ to regulate the cell alignment.

Thus, PI3K-Akt and Rho-Rock are two major pathways that act separately and downstream of EF-VEGFR activation to induce cell elongation and orientation. The integrin αvβ3 interacts with VEGFR signaling during mechanotransduction in endothelial cells responding to shear stress (Wang and Gotlieb, 1999) but this is not involved in EF-induced orientation. This is interesting, given the common downstream signaling elements shared by cells orienting to shear stress and to a small EF (Chen et al., 1999). Thus, EFs use one of the most important signaling strategies to modulate endothelial cell behaviors. There are parallels here with directional migration of epithelial cells, which is stimulated by a chemical gradient of EGF. EF-induced epithelial cell migration is initiated by EGF receptor signaling (Zhao et al., 2002b; Zhao et al., 1996; Fang et al., 1998).

Cytoskeleton involvement

Cytoskeleton reorganization underlies the cell elongation, orientation and migration induced by various stimulations (Wang and Gotlieb, 1999; Girard and Nerem, 1993; Thoumine et al., 1995; Schenk et al., 1999). EF-induced endothelial cell orientation and alignment also may involve actin filament (F-actin) and microtubule rearrangements, because both reoriented and came to align with the long axis of the cells after exposure to an EF (Fig. 1G-I). Latrunculin-A treatment completely abolished the elongation response but cell reorientation in an EF remained at around half the control value (P<0.001). Interestingly, this suggests that actin filament reorganization might play a different role in the control of cell reorientation than in cell elongation, and provides an experimental model with which to investigate the interaction of EF-induced cell shape change and cell reorientation.

Membrane potential perturbation

It has been suggested that cells align perpendicular to an EF in order to minimize the voltage drop across themselves (Robinson, 1985). HUVEC cells have a membrane potential (mean±s.d.) of −16±12.7 mV as single cell, and −23.6±5.5mV as confluent cells in culture (Vargas et al., 1994). The threshold inducing perpendicular alignment, elongation and directional migration was 50–100 mV mm⁻¹, equivalent to 1–2 mV across a cell 20 μm in diameter. This represents a fluctuation within the normal variation of membrane potentials observed in those cells. In 70% of those cells, Vargas et al. (Vargas et al., 1994) observed that a small outward current was activated with a threshold of 0 mV (depolarization of membrane potential), whereas inward rectifying currents were activated with a threshold of 100 mV (hyperpolarization). In other words, small currents at membrane potentials between −100 mV and −20 mV indicate that channels in HUVEC cells were mostly closed in that range (Vargas et al., 1994). This suggests that the applied electric fields might be too small to signal via the modified membrane potential to induce perpendicular alignment. Not all cell types align perpendicularly in an EF. Human small cell lung cancer cells and differentiated Dictyostelium ameba for instance, elongate parallel to the EF or showed no preferential orientation (Zhao et al., 2002a).

It is interesting to notice the similarity between the appearance of the cells exposed to EFs (Fig. 1B) and that of cells exposed to flow shear stress (Tzima et al., 2001; Li et al., 1999). DC EFs do cause electro-osmotic water movement (McLaughlin and Poo, 1981). However, it is unlikely that electro-osmotic water movement mediates the effects of a DC EF. First, the electro-osmotic water movement is parallel to the EF vector; if water movement was the cause of the cell alignment, the cells wound have aligned with long axis parallel to the EF vector. The cells aligned perpendicular to the field direction (Fig. 1B, Fig. 2). Second, the water movement is not detectable by volume measurement before and after EF exposure in our experimental setup. Flow shear stress of 10 dyn cm⁻² does not induce cell alignment in sparse culture, similar to cell density in Fig. 2 (Masuda and Fujiwara, 1993). In a chamber similar to our construction, the shear stress =6 μQ/bh² (dyn cm⁻²), where Q is the flow rate (ml second⁻¹), μ is the fluid viscosity (dyn seconds cm⁻². 0.011 for culture medium at 37°C), h is the channel height (~200 μm) and b is the channel width (1 cm for our chamber) (Albuquerque et al., 2000; Levesque and Nerem, 1985). A flow rate about 240 ml hour⁻¹ would be needed to generate a shear stress of 10 dyn cm⁻². Such flow rates do not happen in our system, although marked cell alignment was observed (Fig. 1B, Fig. 2).

M.Z. is a Wellcome Trust University Award Senior Lecturer. The work was supported by The Wellcome Trust (058551 to M.Z.) and British Heart Foundation (FS/2000056, and PG/99191 to M.Z., C.D.M., J.V.F.). We thank Y. Yin, K. A. Mullard, P. Morgan and J. A. Barker for some of the initial experiments.