Effect of microamperage stimulation on the rate of wound healing in rats: a histological study.

[Leffman DJ, Arnall DA Holmgren PR, Cornwall MW. Effect of microamperage stimulation on the rate of wound healing in rats: a histological study. Phys Ther 1994;74 15)5-200.]

Key Words: Electrical stimulation, Microamperage, Wound healing.

In recent years electrical stimulation of very low amplitude and frequency modulation has become an increasingly popular treatment modality. This form of stimulation has been referred to as microelectric neuromuscular stimulation (MENS),[1] microcurrent,[2] and microamperage stimulation MS).[2] Some authors[1,2] have suggested that these forms of stimulation are more effective for the treatment of a wide variety of conditions, including wound healing, than more conventional modes of electrical stimulation. Unfortunately, the efficacy of low-amplitude stimulation has not been demonstrated.

The use of electrical stimulation, in general, to enhance wound healing is not new, yet its efficacy is still somewhat equivocal. Alvarez et al[3] reported increased healing rates compared with a control group of skin wounds in pigs following use of continuous direct current of positive polarity that ranged between 50 and 300 [mu]A over a 24-hour period. After 7 days of treatment, an increase in the rate of collagen synthesis was shown for these animals as compared with untreated animals. Other investigators have reported similar results with the use of continuous low-intensity direct current (LIDC)[4-6] and high-voltage pulsed galvanic stimulation.[7-10] Other investigators[8,11,12] have shown no improvement in wound strength or healing rate with other forms of low-intensity electrical stimulation. The greatest difficulty when evaluating the efficacy of electrical stimulation for enhancement of wound healing is the tremendous variety of stimulation variables cited in the literature. The equivocal results, therefore, could be the result of the different methods and models used in each study.

Although many of the treatment protocols described utilized low-amplitude currents, these levels are not generally considered to be in the microamperage range.[13] For the purposes of our study, we will define microamperage stimulation as being the use of low-intensity (< 1 mA/[cm.sup.2]), low pulse rate (<3 Hz), monophasic current for the achievement of a therapeutic effect. Although this definition excludes some currently available stimulators because they lack the ability for current variables to be modified beyond these limits, we believe our definition is sufficiently broad for the majority of MS applications and takes into account the size of electrode used. Using our definition, we could find no published studies that used this type of current to increase the rate of wound healing in rats.

Much of the theoretical basis for the use of MS is based on the work of Becker.[14-16] He proposed that minute direct current flows through the perineural cells of the body via their semiconducting properties.[15] Ingvar[17] had previously proposed that this current greatly influences such things as health, growth, perception of pain, and healing. Becker also proposed that this current is altered in reaction to injury. This altered current flow is termed the "current of injury."[14,16] Further studies have shown that a small negative-polarity current is a key feature in dedifferentiation of cells necessary for regeneration.[18] Application of this current in nonregenerating animals (ie, rats and frogs) has resulted in the initiation of regeneration.[19] Blocking the current in regenerating species has retarded limb growth.[19]

Despite the theoretical basis for using MS to treat cutaneous wounds, controlled, experimental studies that demonstrate its effectiveness have not been performed. In addition, the stimulation variables to be used have not been established. Because of the growing enthusiasm for MS, and the paucity of supporting evidence for its use, our investigation was undertaken. Our null hypotheses were (1) that the wounds of electrically stimulated animals would not show a significantly different change in size over time compared with the wounds of control group animals, which received no treatment, and (2) that no significant change in fibroblast density, blood vessel density, or epithelial thickness would be seen in the electrically stimulated animals compared with the controls.

Method

Animal Care and Handling

Twelve female Sprague-Daweley rats from the same litter mean weight [[+ or -]SD], 300[+ or -]21 g; mean age [[+ or -]SD], 5[+ or -]0 months) were selected for this study. A nonhuman model was chosen to better control variables and to reduce time and costs. We felt that the use of small mammals such as the rat would fulfill these objectives. In addition, their physiological response to a wide variety of perturbations is sufficiently similar to that of humans to make them a good animal model to extrapolate the results and conclusions of studies to humans.[20]

The animals were handled and familiarized with the treatment procedures and research personnel for 4 weeks before the beginning of the experiment. The rats were fed standard rat chow and water ad libitum.

Surgery

On the day of surgery, the animals were brought to stage 4 anesthesia induced by ether inhalation. Anesthesia was maintained and respiration and heart rate were monitored continuously. A 10- x 6-cm rectangular area was shaved over each scapula for the placement of a 0.41-cm square electrode next to the skin. An 8-mm Keyes dermal punch was applied to the area and lightly twisted, producing a circular incision through the dermis. The skin within the incision was resected back with mouse-tooth forceps, and its underlying connective tissue was cut with surgical scissors.

The thickness of the removed skin section was measured using a ratchet and friction thimble outside micrometer.(*) The micrometer was accurate to within 0.0254 mm (0.001 in), as specified by the manufacturer. After the rat was removed from the ether anesthesia, the diameter of the circular wound was measured with a transparent hole template. The wound area was sponged clean with a sterile gauze pad, and the animal was observed until it had recovered fully from the anesthesia. Recovery took less than 15 minutes. The wounds were not covered or bandaged. There were no observable complications from surgery or anesthesia.

Group Assignment

Using the tissue samples acquired during surgery, pairs of animals were matched according to skin section thickness. One animal from each pair was randomly assigned to the treatment and control groups. Treatment was initiated on all animals 24 hours after surgery,

Treatment Session Handling

During all treatment sessions, the rats were lightly restrained in a clear plastic cone-shaped tube into which the animal crawled and rested comfortably. The small end of the cone provided an air hole through which the rat's nose projected; the large end of the cone was loosely secured with a plastic tie so that the animal could not back out. An opening was cut in the side of the plastic tube for access to the treatment area. All treatments were given in the sleep phase of the rat's 12:12-hour on:off diurnal light cycle. The rats were also housed separately to prevent the wound area from being injured by another rat.

Procedure

Both the treatment and nontreatment electrodes consisted of 0.41- x 0.41-cm carbonized rubber. Wound treatment began by inserting the electrodes through the window in the cone and placing them over saline-soaked gauze pads.

The treatment protocol used in this study was a modification of the method described by Carley and Wainapel.[4] First, we secured the negative electrode on sterile moist gauze placed directly over the wound. The positive electrode was then placed on the shaved area contralateral to the wound. Skin contact for the positive electrode was made by placing a sterile-saline soaked gauze under the entire area of the pad. A current intensity of 100 [mu]A at 0.3 Hz on a 50% duty cycle was applied through the electrodes to the wound for a duration of 2 hours, once a day, using a commercially available stimulator.[dagger] The shape of the applied waveform was monophasic and rectangular. Because of the lack of research concerning the use of MS for wound healing, the particular stimulation variables were chosen based on recommendations by the manufacturer of the stimulator used in this study (Picker RI, personal correspondence, 1991).[2,21]

After the initial 3 days of treatment, the polarity of the treatment and nontreatment electrodes was reversed and remained in this configuration for the duration of the experiment (total of 14 days) for the experimental animals. The reversal of polarity is similar to the protocol used by Wolcott et al[6] in their investigation of skin ulcer treatment using LIDC. The control group animals underwent the same electrode placement and handling as the treatment group animals, with the exception that no current was delivered through the electrodes. The wounds were left uncovered except during treatment each day. The nontreatment electrode site was shaved every 4 days to ensure good contact.

At the beginning of the study, we decided that when 50% of the animals had achieved wound closure, the study would be terminated. If the wound areas of the control and treatment group animals had healed completely before the wound samples were obtained for histological staining, wound remodeling might have taken place such that any differences attributable to the electrical stimulation between the two groups of animals would no longer be evident.

Data Reduction and Analysis

Wound size measurement. The wound area for all animals was inspected daily for general condition and infection. Infection was judged to be absent if there was no yellow or brown exudate, no red inflamed areas, and no odiferous discharge coming from the wound. Wound size was measured immediately after each treatment using a transparent hole template with concentric circular markings. The template was placed over the wound, and the circular ring nearest to the size of the wound was noted. The resolution of the template was 1.58 mm. This process was repeated a minimum of three times for each measurement, with the value obtained two or more times being reported. The same person performed all measurements and was blind with respect to the animal's treatment assignment.

Histologic preparation. At the conclusion of the 2-week treatment period, the animals were brought to stage 4 anesthesia with anhydrous ethyl ether. A 10- x 10-mm sample of the wound area was excised and fixed in formaldehyde. The tissue sample included normal cutaneous and subcutaneous tissue peripheral to the wound and was to a depth that included the muscular layers. Following tissue sampling, the animals were sacrificed by carbon dioxide inhalation using an anesthetic box. The fixed tissue sections, including the wound area and the adjacent normal skin, were embedded in paraffin and cut into 7.0-[mu]m-thick serial sections. For each rat tissue sample, 2 serial sections were prepared using hematoxylin and eosin (H&E) stains, 2 serial sections were prepared using Milligan's trichrome stain, and 2 serial sections were prepared using Ledrum's fibrin stain. Six slides, therefore, were prepared for each control and treatment group rat.

All slide preparations were examined under a x 100 light microscope for epithelial thickness, vascular density, and fibroblast density. Epithelial thickness was measured by a standard microscope ocular micrometer. Capillaries and fibroblasts were counted in five predetermined areas of each wound from the H&E slides. The areas consisted of the center as well as points corresponding to 12, 3, 6, and 9 on a clock. Again, the same individual performed all measurements and was blinded to the group from which the tissue samples were obtained.

Data Analysis

A two-way analysis of variance (ANOVA) design for repeated measures on one factor (time) but not on the other factor (group) was used to test for differences in wound diameter. Statistical differences between the two groups concerning epithelial thickness, vascular density, and fibroblast density were tested using two-tailed independent t tests. An alpha level of .05 was used for all tests of statistical significance.

Results

The ANOVA results for wound diameter are presented in Table 1. As expected. a significant (P<.05) reduction in wound diameter occurred over the 14 days of treatment in this study. There was no significant (P>.05) difference, however, between the two groups with respect to wound diameter. In addition, no significant (P> 05) interaction was found between the two groups and time. Figure 1 shows the average percentage of wound closure over the 14 days of treatment, The t tests showed no significant difference between the two groups with respect to epithelial thickness (t= -1.80, df= 10, P =. 103), fibroblast density (t= -1.12, df= 10, P=.289), or vascular density (t= -0.86, df= 10, P=.411). Table 2 shows the means and standard deviations for each of these variables as well as the results of the t tests. Figure 2 depicts a histological sample from a treated wound and an untreated wound.

Table 1. Analysis-of-Variance Results for Wound Diameter
Source         df   SS         MS         F
Group                  74.23      74.23     2.92
Error          12   304 81        25.40
Time           13   118764.00   9135.69   872.67(a)
Group x time   13   137.96        10.61     1.01
Error          155  1622.64       10.47
(a)P< .05.

[TABULAR DATA 2 OMITTED]

Discussion

Electrical stimulation as applied in this study had no discernable effect on the healing rate of acute dermal punch wounds in rats. In addition, the t tests prevented us from also rejecting our second null hypothesis. Therefore, MS as performed in this study did not increase fibroblast density, blood vessel density, or epithelial thickness.

Other possible factors that might have affected our results include (1) leaving the wound uncovered, (2) wound contraction, (3) sample size, and (4) the treatment protocol used. The wounds were left uncovered primarily because of the impracticality of attempting to cover them. Bandages were not secured to the wounds because the bandages were either not removable for daily wound inspection and treatment or their removal would have been traumatic to the animals.

Measurements of wound size were taken after each treatment session. Because the wound sites were left uncovered between stimulation sessions, a scab formed over each of the wounds. Although the formation of a scab during healing is normal, its presence can retard epidermal cell migration.[22,23] In this study, the scab was moistened and made soft by the sterile saline solution used as the conducting medium during the 2-hour treatment. In almost all instances, the scab was sufficiently moistened such that it sloughed off during handling of the rat after treatment. Only on two separate occasions was it necessary to gently roll aside the edges of the scab to visualize the wound for measurement purposes. At no time was any bleeding noted from any of the wounds during treatment or during measurement of the wound. Because the formation and removal of the scab occurred in both treatment and control group animals, we do not believe that it affected our results. Previous research[22] indicates that the removal of the scabs may actually have hastened healing. The presence of the scab during electrical stimulation, however, may have inhibited current penetration into the wound site. Although we believe that this is highly unlikely, its possible influence should be investigated in future studies of this kind.

A large portion of the decrease in wound size observed in this study was due to myofibroblast activity causing wound contracture. Peacock and Van Winkle[24] noted that 5 to 7 days after injury, movement of the skin edges proceeds at a rate of 0.6 to 0.75 mm per day. Although this aspect could not be controlled, we believe that it was a constant feature for both groups and therefore had little or no effect on the results.

Although our study was conducted with a relatively small sample, the efficacy of electrical stimulation in the healing of dermal wounds has been demonstrated in as few as 11 subjects.3 The inclusion of additional rats would certainly have improved the statistical power of our results. Post hoc power analysis[25] revealed that as many as 235 additional rats would be needed to show a statistically significant difference between the groups on the variables tested in this study and the effect size observed. Although additional subjects would have improved the statistical power, it is unlikely that a larger sample size would have altered our results. Given the "trend" in our data, a larger sample would possibly have resulted in significantly better wound reduction in the control group compared with the treatment group.

The stimulation variables chosen for this study may not have been optimal, and enhanced wound healing could be found with different settings. We attempted to select stimulation settings that were commonly used and clinically relevant. The stimulation settings selected for this study were modifications of those used by other investigators[4-16,12,26] and were recommended by the manufacturer of the stimulator we used (Wing TV, personal correspondence, 1991). Some studies using other forms of stimulation had longer treatment durations (6 hours per day)[5,6,8] and higher stimulation intensities (300-700 [mu]A).[4-6] We stimulated the rats' wounds for only 2 hours a day with a maximum current intensity of 100 [mu]A. Although this protocol is commonly found and advocated for clinical use, it may not be optimal for enhancing wound healing. Additional, controlled studies dealing with MS currents should be conducted to determine the correct stimulation settings required for wound healing.

An additional limitation of our study is the use of an animal model. Although the rat has been shown to be an excellent model for research into a wide variety of physiological actions and mechanisms, differences exist between rodent and human skin.20 We still believe, however, that the results of our investigation can be extrapolated to humans with confidence.

The clinical implications of our result!; are that, given the limitations of the study, MS is not indicated to accelerate the natural healing process of acute dermal wounds, including superficial lacerations, abrasions, and postsurgical incisions. Because our study did not investigate chronic wounds, which are frequently seen in physical therapy, the use of MS in the treatment of these types of wounds cannot be commented on. Further research should be conducted to determine the efficacy of MS in the treatment of chronic wounds.

Conclusion

The purpose of this study was to examine the efficacy of MS for wound healing. The results of our study indicate that rate of wound closure, fibroblast density, vascular density, and epithelial thickness were not significantly enhanced. Within the limitations of this study, we believe that the use of MS to accelerate acute superficial tissue healing is not supported by current research.

Acknowledgments

We thank Doug Van Cleave and Peggy Campbell at the small animals research facility at Northern Arizona University for their help and practical advice.

References

[1] Picker RI. Micro electrical neuromuscular stimulation. Network-Electrix. March 1987:S72-S74. [2] Picker RI. Current trends: low-volt pulsed microamp stimulation, part 1. Clinical Management in Physical Therapy. 1988;9(2):10-14. [3] Alvarez OM, Mertz PM, Smerbeck RV, et al. The healing of superficial skin wounds is stimulated by external electrical current. J Invest Dermatol. 1983;81:144-148. [4] Carley PJ, Wainapel SF. Electrotherapy for acceleration of wound healing: low intensity direct current. Arch Phys Med Rehabil. 1985;66: 443-446. [5] Gault WR, Gatens PF. Use of low intensity direct current in management of ischemic skin ulcers, Phys Ther. 1976;56:265-269. [6] Wolcott LE, Wheeler PC, Hardwicke HM, et al. Accelerated healing of skin ulcers by electrotherapy; preliminary clinical results. South Med J. 1969;62:795-801. [7] Brown M, Gogia PP. Effects of high voltage stimulation on cutaneous wound healing in rabbits. Phys Ther. 1987;67:662-667. [8] Brown M, McDonnell MK, Menton DN. Polarity effects on wound healing using electrical stimulation in rabbits. Arch Phys Med Rehabil. 1989;70:624-628 [9] Cupshan M. High-voltage galvanic stimulation: an aid to post-operative healing. Current Pod. May 1981:110-114. [10] Cruz N, Bayron F, Suarez A. Accelerated healing of full-thickness burns by the use of high-voltage pulsed galvanic stimulation in the pig. Ann Plast Surg. 1989;28:49-55. [11] Deuland R, Hoffer RE, Seleen WA, et al. The effects of low voltage current on healing of thermal third degree wounds. Cornell Vet. 1977;68;51-59. [12] Wu KT, Go N, Dennis C, et al. Effects of electric currents and interfascial potentials on wound healing. J Surg Res. 1967;7:122-128. [13] Alon G. Principles of electrical stimulation. In: Nelson RM, Currier DP, eds. Clinical Electrotherapy. East Norwalk, Conn: Appleton & Lange; 1991:35-103. [14] Becker RO. The Body Electric. New York, NY: William Morrow & Co Inc; 1985. [15] Becker RO. The significance of bioelectric potentials. Biochem Biogen. 1974;1:187-199. [16] Becker RO. The electrical control of growth processes, Med Times. 1967;95:657-659. [17] Ingvar S. Reaction of cells to the galvanic current in tissue cultures. Proc Soc Exp Biol Med 1920;17:198-199. [18] Becker RO, Murray DG. A method for producing cellular dedifferentiation by means of very small electical currents. Trans NY Acad Sci. 1967;29:606-615. [19] Vanable JW, Hearson LL, McGinnis ME. The role of endogenous electrical fields in limb regeneration. Prog Clin Biol Res. 1983; 110: 587-596. [20] Gill TJ, Smith GJ, Wissler RW, et al. The rat as an experimental animal. Science. 1989;245: 269-276. [21] Picker RI. Current trends: low-volt pulsed microamp stimulation, part II. Clinical Management in Physical Therapy. 1988;9(3):28-33. [22] Rovee PT, Kurowsky CA. Labun J. Effect of local wound environment in epidermal healing. In: Maibach HI, Rovee DT, eds. Epidermal Wound Healing. Chicago, Ill: Year Book Medical Publishers; 1972:159-181. [23] Feedar JA, Kloth LC. Conservative management of chronic wounds. In: Kloth LC, McCulloch JM, Feedar JA, eds. Wound Healing: Alternatives in Management. Philadelphia, Pa: FA Davis Co; 1990:135-172. [24] Peacock EE, Van Winkle E. Surgery and Biology of Repair. Philadelphia, Pa: W]B Saunders Co; 1970. [25] Keppel GT. Design and Analysis: A Researcher's Handbook. Englewood Cliffs, NJ: Prentice-Hall Inc; 1973. [26] Assimacopoulos D. Wound healing promotion and the use of negative electric current. Am Surg. 1968;34:423-431.

DJ Leffmann, PT, is Staff Physical Therapist, Group Health Co-operative-Central Hospital, 200 15th East, Seattle, WA 98112. Mr Leffmann was a graduate student, Department of Physical Therapy, Northern Arizona University, at the time this study was completed.

DA Arnall, PhD, PT, is Associate Professor, Department of Physical Therapy, Northern Arizona Unive PO Box 15105, Flagstaff, AZ 86011.

PR Holmgren, PhD, PT, is Associate Professor, Department of Biology, Northern Arizona University.

MW Conrwall, PhD, PT, is Associate Professor, Department of Physical Therapy, Northern Arizona University, PO Box 15105, Flagstaff, AZ 86011 (USA). Address all correspondence to Dr Cornwall.

This study was approved by the Institution Animal Care and Use Committee of Northern Arizona University.

This article was submitted February 25, 1993, and was accepted September 5, 1993.