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PHYS THER
Vol. 86, No. 1, January 2006, pp. 50-59
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Research Reports

Periodic Direct Current Does Not Promote Wound Closure in an In Vitro Dynamic Model of Cell Migration

Charles Godbout and Jérôme Frenette

C Godbout, PT, is a PhD student, Department of Rehabilitation, Laval University, Quebec City, Quebec, Canada
J Frenette, PT, PhD, is Associate Professor, Department of Rehabilitation, Laval University, Quebec City, Quebec, Canada

(jerome.frenette{at}crchul.ulaval.ca) Address all correspondence to Dr Frenette at CHUL Research Center, T-R-93, 2705 Blvd Laurier, Sainte-Foy, Quebec, Canada G1V 4G2


Submitted February 8, 2005; Accepted June 20, 2005


   Abstract
 
Background and Purpose. A prevailing paradigm is that electrical fields can promote cell migration and tissue healing. To further validate this paradigm, we tested the hypothesis that periodic direct current (DC) can enhance wound closure using an in vitro dynamic model of cell migration.

Methods and Results. Layers of primary fibroblasts were wounded and treated with DC under various voltages. Repair area, cell velocity, and directionality as well as lamellipodium area were evaluated at different times. Direct current had no beneficial effect on cell migration. Moreover, prolonged stimulation under the highest voltage led to significant reduction in wound closure and cell velocity. The reduction of membrane protusions in stimulated cells may be associated with the deleterious effect of DC.

Discussion and Conclusion. Contrary to the authors’ expectations, they found that periodic DC did not promote wound closure, a finding that emphasizes the need to clarify the complex effects of electrical fields on migrating cells. [Godbout C, Frenette J. Periodic direct current does not promote wound closure in an in vitro dynamic model of cell migration. Phys Ther. 2006;86:50–65.]

Key Words: Direct current • Electrical stimulation • Fibroblast • Galvanotaxis • Wound healing


   Introduction
Top
 Abstract
 Introduction
Method
 Results
 Discussion
 Conclusion
 References
 
Mobile cells encounter different signals that can potentially guide their migration. Directional cellular locomotion is thought to be under the guidance of various distinct mechanisms such as chemotaxis, haptotaxis, and galvanotaxis.1 Chemotaxis is a well-characterized mechanism whereby a chemical gradient serves as a directional signal that organizes cell movement.2 Cell motility also can be guided by changes in the composition of the extracellular matrix that forms the substratum for migration. An adhesive molecule, which is present in increasing amounts in the extracellular matrix, generates haptotaxis and can accurately position migrating cells during embryogenesis and tissue regeneration.3 Lastly, electrical fields (EFs) can cause changes in cell shape and polarization and induce directional migration.1, 4–6 The generation of EFs necessary for a galvanotactic response is not limited to externally applied fields under artificial conditions. Previous studies7, 8 have shown that voltage gradients exist in developing embryos. In addition, tissue trauma can create lateral voltage gradients that reach up to 100 mV/mm at the borders of wounds.9 This endogenous potential diminishes to 0 mV/mm at a distance of 3 mm from the site of the injury.10 These electrical potentials result largely from ion fluxes through leaky cell membranes, producing direct currents (DCs) that decay over time.

The presence of an electrical gradient is important for tissue repair, and nullification of endogenous EFs can significantly alter wound healing.11 Clinicians commonly use electrical stimulation because endogenous currents facilitate the normal healing process and the healing potential may be amplified by the application of external currents. Studies using various types of currents have shown that exogenous stimulation could promote the healing of both hard and soft tissue injuries.12–22 For example, high-voltage pulsed current stimulation can accelerate the healing of chronic skin ulcers, increase Ca2+ influx, and promote the rate of protein and DNA synthesis by fibroblasts.14, 23, 24 Moreover, the migration of various cell types such as amoebae, leukocytes, fibroblasts, epithelial cells, bone cells, and neural crest cells is sensitive to DCs.25–33 This response is significantly increased when cells migrate on type I and IV collagens or in the presence of growth factors,34, 35 indicating that external factors can modulate cell migration. Although there is substantial evidence to support the effect of DC in static cells in vitro, the appropriate level of electrical stimulation that must be achieved to trigger the desired effect has never been extensively tested in migrating cells. Because spatial and temporal control of cell migration is of fundamental importance for tissue healing, we hypothesized that electrical currents increase the directionality or rate of migration of fibroblasts in an in vitro dynamic model of wound closure. Thus, the first objective of this study was to set up a model in which fibroblasts migrate under relevant conditions while being electrically stimulated. This model resembles, to some extent, normal clinical situations where an EF is applied to wounded connective tissues. The second objective was to characterize the repair area, cell speed, and cell directionality in a model of wound closure exposed to DC at various voltages. While reaching this objective, we also verified whether unidirectional currents or periodical reversal of current polarity had a greater effect on wound closure. Lastly, to understand how DC may influence cell migration, we evaluated cell membrane protusions under experimental and control conditions.


   Method
Top
 Abstract
 Introduction
 Method
Results
 Discussion
 Conclusion
 References
 
Isolation and Plating of Primary Cultures

To generate primary fibroblast cultures, rat Achilles tendons were cut into small pieces and placed in a 50-mL tube containing a solution of collagenase (1.1 mg/mL) (Sigma-Aldrich*) and trypsin (2.5 mg/mL) (Invitrogen{dagger}) (modified from Freshney36). Following periods of incubation and agitation, tendon-derived fibroblasts were subsequently cultured in Dulbecco’s modified Eagle’s medium (DMEM{dagger}) containing sodium bicarbonate, 10% fetal bovine serum (FBS) (Wisent{ddagger}), and penicillin/streptomycin (100 U/mL and 100 µg/mL, respectively{dagger}). Cells from passages 5 to 9 were removed from the culture flasks by trypsinization 16 to 18 hours before the experiments and were seeded at confluency on sterile glass coverslips (12-mm diameter) in a 24-well plate at a density of 1.0 x 106 cells/well.

Galvanotactic Chamber and Wound Injury

A culture slide (Nalge Nunc§) was modified to create a galvanotactic chamber with a platinum electrode fixed on opposite sides (Fig. 1). The chamber then was placed on a glass slide and sealed with silicone. Agarose (2%) double bridges were placed at both sides to minimize the introduction of electrolysis products and changes in pH. The coverslip then was transferred to the chamber and fixed with Krazy Glue|| to prevent displacement. A linear wound parallel to the electrodes was produced in the cell layer using a pipette tip. Following 3 washes to remove cellular debris, the galvanotactic chamber was partially filled with culture medium. In order to ensure pH stability, sodium bicarbonate was replaced by 25 mM HEPES.* The chamber was transferred to a microscope stage incubator (CSMI#) (Fig. 1) where the culture medium temperature was kept at 37°C with a probe connected to a controller.# The central well was constantly perfused with a peristaltic pump, at a rate of approximately 225 µL/min, to ensure a consistent replacement with fresh medium. The electrical current was provided by a power supply (Power-Pac 1000**) connected to the electrodes emerging from the chamber.


Figure 1
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[in a new window] 		Figure 1. The CSMI is a versatile microscope stage incubator for cell and tissue cultures that accommodates rectangular disposable chambered slides. A wounded layer of fibroblasts was maintained under appropriate and relevant conditions using this experimental setup. The perfusion tubing is placed to one side, and the temperature controller probe is omitted for clarity.

 
Experimental Protocol

Fibroblast layers were exposed for 8 hours to potential differences of 85, 120, 165, and 215 mV/mm. The cells were stimulated with DC for 10 minutes followed by a 10-minute rest period. The polarity of the electrodes was changed between stimulation periods, creating bidirectional currents. Control cells were submitted to the same experimental conditions except that they were not exposed to any potential difference. Electrical stimulation and pH were periodically monitored. To evaluate the effect of unidirectional current, we stimulated fibroblasts at 120 mV/mm using the same protocol with no change in polarity. However, the perfusion rate was raised to 450 µL/min in order to maintain pH within a physiological range. Lastly, control and bidirectional current experiments at 215 mV/mm were conducted for 5 hours to analyze cell membrane protusions. These parameters were selected because fibroblast velocity peaked after approximately 5 hours, and we hypothesized that these voltages should provide the most detectable differences.

Video Microscopy and Image Analysis

The kinetics of wound closure were calculated over time by quantitative measurements using time-lapse video microscopy. Images of the cells were captured every 30 minutes during the protocol using a 10x objective lens on a Nikon inverted microscope equipped with a CCD CoolSNAP camera.{dagger}{dagger} The images were analyzed using MetaMorph image analysis software.{ddagger}{ddagger} Following acquisition, the images were converted from pixels to micrometers using a standard image. Wound repair area was evaluated by measuring the area covered by migrating cells from both sides of the wound. Fibroblast velocity and directionality were calculated by tracking 6 cells per experiment, 3 from each side of the wound. Cell velocity was evaluated using 3 paradigms: the hour-based velocity, the maximal velocity (defined as the maximal velocity among the hour-based velocities), and the mean velocity (expressed as the total displacement of cells divided by the duration of the experiment). The directionality of the cells also was assessed by measuring a cosine value of the angle between cell position at the beginning and end of the protocol. If a cell migrated at an angle of 90 degrees, parallel to the EF, a cosine value of 0 was assigned. If a cell migrated at an angle of 0 degrees or 180 degrees, perpendicular to the EF, an absolute cosine value of 1 was assigned. Lastly, the area of cell protusions, or lamellipodium area, was measured by capturing images of the wound margins by differential interference contrast microscopy using a 20x objective lens (modified from Galiacy et al37). The lamellipodium area of each cell at the wound margins was measured and expressed as a mean for both sides (Fig. 2).


Figure 2
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[in a new window] 		Figure 2. Schematic representation of a migrating cell closing the wounded area. Lamellipodium area corresponded to the cell membrane protusion contained between the front of the cell and the dotted line.

 
Data Analysis

Hour-based velocity and wound repair area of bidirectional current experiments were statistically evaluated by a 2-way analysis of variance (ANOVA) followed by a t test. Mean and maximal velocities as well as cell directionality were evaluated by a 1-way ANOVA, and post hoc differences were determined using the Fisher least significant difference test. Wound repair area from unidirectional current experiments and lamellipodium area were analyzed in the same manner. The level of significance was set at P<.05. Data are expressed as means ± standard error of the mean (SEM). Statistical analyses were performed using StatView software (SAS§§).


   Results
Top
 Abstract
 Introduction
 Method
 Results
Discussion
 Conclusion
 References
 
To study the effect of DC in this model of wound closure, we first stimulated fibroblast layers with bidirectional currents at various voltages for 8 hours. The electrical current was applied for 10 minutes followed by a 10-minute rest period. The electrode polarity was changed between periods of stimulation. Time-lapse video analyses showed that the EF had no effect on wound closure until hour 7 of the experiment. From this time point, we observed a significant reduction of wound repair area with cells stimulated at 215 mV/mm, when compared with control fibroblasts at 7 and 8 hours of stimulation or those under a potential difference of 120 mV/mm at 8 hours (Figs. 3 and 4a). Indeed, wound closure was generally gradual and progressive, but the closure rate of fibroblast layers stimulated with the highest potential difference had a tendency to slow down and even regress during the last hour of the experiment.


Figure 3
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[in a new window] 		Figure 3. Micrographs of wound closure captured after 0 hours (A and B), 4 hours (C and D), and 8 hours (E and F). The left and right columns represent wounded cell layers exposed to control conditions and bidirectional current at a voltage of 215 mV/mm, respectively. Bar=65 µm.

 

Figure 4
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[in a new window] 		Figure 4. (a) Wound repair area with control conditions and bidirectional currents at voltages of 85, 120, 165, and 215 mV/mm. Asterisk (*) indicates significantly different from cells stimulated at 215 mV/mm (P<.05). The error bars were omitted for the sake of clarity. Values in the table are expressed as means ± SEM x103 µm2 (n=7 for the control group, n=5 for the experimental groups). (b) Velocity of fibroblasts with control conditions and bidirectional currents at voltages of 85, 120, 165, and 215 mV/mm. Fibroblasts stimulated at 120 mV/mm (a), 165 mV/mm (b), and 215 mV/mm (c) were significantly different from control cells (P<.05). The error bars were omitted for the sake of clarity. Values in the table are expressed as means ± SEM µm/h (n=5 for all groups). Six cells were tracked for each experiment.

 
The measurements of fibroblast directionality at the edge of the wound indicated that cells migrated quite parallel to the EF. However, the electrical stimulation had no influence on the orientation of fibroblast locomotion, as control cells migrated identically (Table). To evaluate the effect of bidirectional currents at the level of individual cells, we measured the hour-based, maximal, and mean velocities of fibroblasts at the wound margins. Significant differences between control and experimental groups were observed among hour-based velocities, as illustrated in Figure 4b. Interestingly, after 5 hours of stimulation and until the end of the experiment, results obtained with fibroblasts at a voltage of 215 mV/mm were significantly lower than those of control cells. Furthermore, the maximal velocity of individual fibroblasts did not differ between groups, whereas the mean cell velocities of the control and 120-mV/mm groups were significantly higher than that of the 215-mV/mm group (TableGo).


View this table:
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[in a new window] 		Table. Effects of Bidirectional Current Stimulation on Cell Migration Parametersa

 
Another set of experiments with unidirectional currents was performed to ascertain that the periodic reversal of polarity did not inhibit cell migration. In this dynamic model of fibroblast migration, a unidirectional current of 120 mV/mm did not increase the closure rate on either the cathodal or anodal side of the wound, suggesting that DC cannot potentiate the migration or spread of migrating cells (Fig. 5). Because membrane protusion is required for cell locomotion, we also quantified the area of the lamellipodia in slow (stimulated at 215 mV/mm) and fast (control) migrating fibroblasts. The area of the fibroblast lamellipodia exposed to an EF was reduced by roughly 15% compared with the control group (Fig. 6). This reduction in the area of the lamellipodia may be associated with the delay in wound closure observed with electrical stimulation.


Figure 5
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[in a new window] 		Figure 5. Wound repair area in layers stimulated for 8 hours with bidirectional current and unidirectional current at a potential difference of 120 mV/mm. Results are expressed as percentages of their respective controls (0 mV/mm). Values are expressed as means ± SEM. There was no difference between groups (n=5 for all groups).

 

Figure 6
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[in a new window] 		Figure 6. Lamellipodium area of cells at wound margins. Control and bidirectional current experiments at voltage of 215 mV/mm were performed for 5 hours. Values are expressed as means ± SEM. Asterisk (*) indicates significantly different from control cells (P<.05, n=5 for both groups).

 

   Discussion
Top
 Abstract
 Introduction
 Method
 Results
 Discussion
Conclusion
 References
 
The clinical practice of physical therapy and of orthopedics has created a variety of biophysical environments in an attempt to optimize tissue repair. These environments include physical exercise, stretching, manual therapy ultrasound, electrical stimulation, and other physical modalities. Although human or animal studies may provide very important information, in vitro studies are essential to evaluate the effects of external stimulations at the cellular or molecular level. The model of wound closure is well accepted to mimic wound healing in vitro37–43 where cells will migrate without external stimulation at least by contact inhibition of movement.1, 44 Alternatively, the presence of endogenous DCs at site of the wound certainly explain why the vast majority of in vitro studies were performed with DC to understand the effects of electrical current on cell migration. In the present study, we first tested the influence of alternating periods of stimulation and rest along with polarity reversal. The rationale for selecting these parameters in this model of wound closure was that continuous DC stimulation without a change in polarity causes temperature and pH perturbations, leading to cell damage and chemical burns.45 These parameters, therefore, were chosen in an attempt to minimize the adverse effects in an in vitro model that resembles skin, ligament, and tendon injuries in which fibroblasts migrate to both ruptured ends.

The present findings demonstrated that bidirectional current did not speed up but even slowed down cell migration in vitro. Furthermore, the experiments with unidirectional current indicated that reversal of polarity was not responsible for the lack of migration and that the application of unidirectional current did not significantly accelerate or inhibit the rate of wound closure. It is important to point out that cell migration generally implies a polarization of the cell that includes a redistribution of cell membrane receptors or downstream mediators/effectors and modifications in actin organization.46 This process cannot be reversed in few minutes,47 and reversing the polarity every 10 minutes would not simply annihilate the effects of EF because cells are generally forced to migrate to fill the empty space. Our results also are consistent with the observation that wound epithelization rates are diminished when cells are exposed to field strengths that are significantly higher than normal.48 Thus, the machinery of migrating cells may already function at a nearly optimal level, and the EFs might interfere with cellular and molecular signals involved in cell locomotion.

Cell directionality did not appear to be influenced by DC in our experiments. We speculate that cell contacts might force them to migrate quite parallel to the wound even without any external stimulation. Indeed, a recent study from Green et al38 demonstrated that all wound margin cells spread into the wound gap perpendicularly to the wound long axis during the first 120 minutes after wounding. In addition, previous studies showing that cell directionality was significantly improved under an electrical stimulation were performed at relatively low cell density where cell contacts were minimized.49–52 It is also interesting that migration velocity and directionality appear as 2 distinct properties of cell locomotion and that the EFs influence mainly the directionality of the cells.35, 47, 51, 53 Nishimura and colleagues47 observed that isolated human keratinocytes migrated at similar rate with or without electrical stimulation, but their directionality was altered under EFs. However, these observations were different in the model of wound closure where fibroblasts are in contact with each other and move in a relatively directed manner.

Our results also demonstrated that a voltage of 215 mV/mm inhibits wound closure. Consistent with these results, the membrane protusion area of cells at the wound margins had smaller lamellipodia in the stimulated cells compared with the control cells. Although the signaling pathway by which an EF initiates lamellipodium formation and cell migration remains unclear, it appears likely that smaller lamellipodia are at least a direct consequence of high-voltage stimulation. We hypothesize that DC may influence rhythmic stretching and squeezing cycles of lamellipodia by perturbing [Ca2+]i in migrating fibroblasts. This hypothesis is based on the fact that migrating cells segregate [Ca2+]i, resulting in an increase in [Ca2+]i at the rear and causing the cells to contract, whereas protrusion formation occurs at the front.46 Actin polymerization/depolymerization and actomyosin contractility are likely the 2 most obvious Ca2+-dependent mechanisms that may dictate cell contraction or protrusion.54 The EF thus may perturb cell activity at the leading edge of moving cells, decreasing lamellipodium size, forcing smaller locomotion steps, and ultimately reducing cell velocity and wound closure rate.

Our results are, to some extent, in contradiction to recent work by Finkelstein et al,55 who used a similar model of wound closure and found that the stimulation of NIH-3T3 fibroblasts with DC at 2.0 V/cm for 1.5 hours induced significant migration toward the cathode when compared with control cells. They also observed that cathodal-facing cells migrated faster than anodal-facing cells at 0.6 and 2.0 V/cm. These discrepancies may be attributable to several factors, including cell type, stimulation parameters, and experimental conditions. For example, melanocytes56 and human dermal fibroblasts,57 unlike many cell types, exhibit neither directional migration nor increased velocity when exposed to DC. The significance of Finkelstein and colleagues’ results was weakened by the fact that EFs greater than 2.0 V/cm are deleterious for cells and that pH measurements and culture medium perfusion were not performed. Previous studies58–70 showed that pH is of critical importance for cell metabolism, survival, or migration and that pH gradients can greatly influence wound healing. In our experimental model, the increases in buffer concentration or perfusion rate were sufficient to support bidirectional currents of 2.15 V/cm for up to 12 hours or 1.2 V/cm of continuous DC stimulation for at least 4 hours. Furthermore, Finkelstein et al performed another set of experiments that included an additional 2.5 hours of incubation before the exposition of injured cell layers to DC. This preincubation, which is even longer than the migration assay itself, prevented an increase of cathodal-directed migration. In these experiments, the unstimulated cells migrated at a speed that was roughly 5 to 24 times higher than those measured without preincubation. These results, however, were consistent with our observations that the application of DC under these conditions does not promote wound closure, suggesting that DC may have different effects on static or mobile cells.

The findings of our study did not appear to result from extensive cell death or inappropriate level of stimulation. Indeed, the addition of Trypan blue, a nontoxic staining that penetrates porous cell membrane (dead cells), showed a relatively low and nonsignificant number of dead cells with or without electrical stimulation. In addition, we observed cell movements at the end of the experiment, indicating that cells were still alive. Otherwise, the voltages applied during our experiments were in the same range as those of endogenous currents and similar to those of previous reports that investigated the effects of electrical stimulation on cell migration.9, 30, 34, 49, 55, 71 Conceivably, electrical stimulation as well as drugs may be represented by a bell-shape curve in which "medium current" would be the most effective dose. Further studies are needed to determine the window of opportunity for EFs in various connective tissue pathologies.


   Conclusion
Top
 Abstract
 Introduction
 Method
 Results
 Discussion
 Conclusion
References
 
In some circumstances where cells are already migrating to the site of injury, DC may not speed up cell migration. It would then be interesting to evaluate the effect of EFs in a chronic wound environment. The construction of a combined in vitro chemotactic and galvanotactic chamber would be helpful in determining the participation of chemical agents and electrical currents in cell migration. We hope that these results will pave the way to determining the best time to use DC and will help to refine the clinical decision-making process of physical therapists using electrotherapy.


   Footnotes
 
Both authors provided concept/idea/research design, writing, and data collection and analysis. Dr Frenette provided project management, fund procurement, and facilities/equipment. The authors thank Dr Claude H Côté for helpful discussions, David Marsolais for isolating the fibroblasts, and Gilles Chabot for the figures.

This research was presented, in part, at the Canadian Physiotherapy Association Congress, May 27–30, 2004, Quebec City, Canada, and the 14th International Congress of the World Confederation for Physical Therapy, June 7–12, 2003, Barcelona, Spain.

Financial support for this study was provided by the National Sciences and Engineering Research Council in Canada, the Fonds de la Recherche en Santé du Québec, the Canadian Institutes for Health Research, and the Fondation Docteur Georges Phénix.

* Sigma-Aldrich Canada Ltd, 2149 Winston Park Dr, Oakville, Ontario, Canada L6H 6J8. Back

{dagger} Invitrogen Canada Inc, 2270 Industrial St, Burlington, Ontario, Canada L7P 1A1. Back

{ddagger} Wisent Inc, PO Box 131, St-Bruno, Quebec, Canada J3V 1Y0. Back

§ Nalge Nunc International, 75 Panorama Creek Dr, Rochester, NY 14625. Back

|| Elmer’s Products Inc, 180 E Broad St, Columbus, OH 43215. Back

# Harvard Apparatus Inc, 84 October Hill Rd, Holliston, MA 01746-1388. Back

** Bio-Rad Laboratories (Canada) Ltd, 5671 McAdam Rd, Mississauga, Ontario, Canada L4Z 1N9. Back

{dagger}{dagger} Nikon Canada, 1366 Aerowood Dr, Mississauga, Ontario, Canada L4W 1C1. Back

{ddagger}{ddagger} Universal Imaging Corp, 402 Boot Rd, Downingtown, PA 19335. Back

§§ SAS Institute Inc, 100 SAS Campus Dr, Cary, NC 27513-2414. Back


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Top
 Abstract
 Introduction
 Method
 Results
 Discussion
 Conclusion
 References
 

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