This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
1. Introduction
The wound healing process is a complex series of phases, beginning with hemostasis, followed by the inflammatory, proliferative, and remodeling phases [1]. During hemostasis, a fibrin network is formed, serving as a scaffold to retain blood cells, plasma, and platelets, thereby forming a blood clot. This clot also acts as a source of growth factors and chemokines, recruiting inflammatory cells (ICs) to migrate to the wound site [2–4]. Adhesion molecules initiate their action on endothelial cells, allowing the recruitment and extravasation of neutrophils and macrophages capable of releasing crucial inflammatory mediators that control the healing process, such as interleukins (IL-1, IL-6), tumor necrosis factor-alpha (TNFα), platelet-derived growth factor (PDGF), fibroblast growth factor-2 (FGF2), transforming growth factor-beta (TGFβ), and vascular endothelial growth factor (VEGF) [5–8].
The action of these molecules leads to vascular neoformation (angiogenesis) and the differentiation of forming cells, such as fibroblasts, which proliferate and initiate the deposition of collagen matrix [5–8]. This matrix provides a substrate for the migration of keratinocytes from the wound margins, guiding them at the interface between the clot and the underlying dermis, thus characterizing the proliferative phase. This process culminates in the epithelial closure of the entire wound, referred to as reepithelialization [5–8].
Various critical factors contribute to the unsatisfactory occurrence of the wound healing process, such as vascular insufficiency in the area, systemic disorders like diabetes mellitus, and blood pressure effects, leading to the development of chronic wounds [1]. In response to this challenge, the scientific community has been exploring local or systemic approaches, methods, and treatments aimed at promoting a more regular, uniform, and stable repair process. These encompass photonic therapy, electrical currents, tissue engineering, skin grafting, herbal remedies, stem cells, biomembranes, and ozone therapy.
Ozone is composed of a mixture containing 5% ozone and 95% oxygen that comes in gas or liquid forms (water or oil) [9, 10]. It can be administered either locally, using ozonized water, ozonized oil, or gas, or systemically through methods such as rectal insufflation, intramuscular, subcutaneous, and intraperitoneal injection. The objective is to enhance tissue oxygenation, boost the body’s defense system, and exert a lethal impact on microorganisms in areas affected by infection [11–13]. In dentistry, recent publications have focused on studying its mechanism of action and effects on dental caries [14], canal and cavity disinfection [15], gingivitis [16], implant dentistry [12, 17–19], biofilm control/periodontitis [19–23], and jaw osteonecrosis [24]. Moreover, the use of ozone therapy has been applied to manage postextraction wound healing [25–27] and palatal wound healing [28]. In the medical field, ozone has been tested to accelerate the healing of cutaneous wounds [29–32], dermatological diseases, and pain management [11, 12]. Ozone displays a diverse array of effects including antimicrobial, anti-inflammatory, antioxidant, and antihypoxia effects. These properties of ozone make it suitable to treat cutaneous wounds, but their effects on the healing of skin wounds remain to be elucidated.
The literature on ozone therapy reveals conflicting findings, primarily due to the diverse results encountered and the different methodologies employed (administration via, dosages and regime duration). For instance, a recent study has shown that the application of highly ozonated sunflower oil does not improve palatal wounds in a randomized clinical trial [28]. On the other hand, Patel et al. [33] showed significant improvements in the healing of palatal wounds after gingival graft removal. According to a systematic review [34], wounds treated with topical ozone had a greater reduction in wound size. Wounds treated with ozonated liquids also had a shorter time to tissue healing by approximately 1 week [34]. Furthermore, numerous studies have demonstrated positive outcomes for socket wound healing [25, 26], alveolar osteitis [35], and treating dermal and skin wounds [29–32].
Although several studies have investigated the potential effects of ozone in the dermatology field, the literature is still scarce regarding its effects to heal cutaneous wounds. Therefore, this study is aimed at assessing the impacts of systemic ozone (O3) therapy and the influence of various dosages of the O3/O2 mixture on the healing of open wounds induced on the dorsum of rats. The main hypothesis is that O3 will enhance tissue repair, and the dosage might impact the biological response. The null hypothesis, on the other hand, is that there will be no added benefits from employing O3 in the healing of cutaneous wounds.
2. Materials and Methods
2.1. Animals
Seventy-two male albino Wistar rats (Rattus norvegicus) aged between 3 and 4 months and with an average weight of 335 g, were used in this study. The rats were sourced from the Animal Facility at the School of Dentistry on the Araçatuba UNESP. All animals underwent an acclimatization period for at least 1 week before the start of the experiment. Throughout the entire investigation, they were housed in plastic cages with four animals per cage and maintained in a climate-controlled environment with a temperature of
During the experimental period, the animals were provided with crushed solid food (Presence Feed, Primor S.A. Mill, São Paulo, SP) and had access to water “ad libitum.” The study received approval from the Animal Ethics Committee of the School of Dentistry in Araçatuba, UNESP (Process 0287-2021), and all procedures adhered to ethical principles of animal experimentation and followed the ARRIVE guidelines for the care and use of laboratory animals [36].
2.2. Sample Calculation
The sample size calculation was conducted using the GPower® program, considering an alpha (Type I) and beta (Type II) error of 5% and 80%, respectively, and a medium effect size of
2.3. Surgical Procedures for Wound Creation
For the creation of wounds, animals underwent general anesthesia induced by intramuscular injection of a combination of ketamine hydrochloride (80 mg/kg/bw) and xylazine hydrochloride (10 mg/kg/bw). Then, the dorsal region of the animals was gently shaved, and the area was aseptically prepared with a 10% povidone-iodine topical solution. Subsequently, a single wound was created in the middorsal portion of each animal using a circular punch with an internal diameter of approximately 10 mm, following the animals’ midsagittal plane. To standardize the wound, two markings were made on the outer surface of the punch, representing 50% of its diameter. Next, the skin in the targeted area of the animal was folded along the midsagittal plane and carefully supported on the surgical table. The punch was positioned with the markings aligned at the skin edge, and with the assistance of a surgical hammer, a gentle and single strike was executed, allowing the release of the skin fragment. After tissue release, a circular wound with regular edges was obtained for all animals.
2.4. Randomization and Treatment Assignment
The animals were randomly assigned using a table generated by the website Randomization.com (http://www.randomization.com/) to four groups that were divided according to the type of treatment, as follows: Control group (C): The animals received no systemic treatment. OZ0.3 group: The animals received an intraperitoneal dose of 0.3 mg/kg/bw of O3, with a concentration of 15 μg/ml of O3 released by the generator; OZ0.7 group: The animals received an intraperitoneal dose of 0.7 mg/kg/bw of O3, with a concentration of 35 μg/ml of O3 released by the generator; and OZ1.0 group: The animals received an intraperitoneal dose of 1.0 mg/kg/bw of O3, with a concentration of 50 μg/ml of O3 released by the generator. The dosage of ozone was based on previous published studies [37–39].
The ozone generator Philozon Medplus V (Santa Catarina, Brazil) was employed, featuring the following specifications: a safety valve with a vacuum system, automatic regulation of oxygen flow, automatic internal catalyzer, and adjustable flow of oxygen concentrations ranging from 5 to 60 μg/ml. These concentrations were set and displayed on the equipment panel. A single examiner performed the intraperitoneal injections.
2.5. Sample Collection and Histological Processing
Six animals from each group underwent euthanasia through an overdose of anesthetic (150 mg/kg, Thiopental®) after 7, 14, and 21 days following the operative procedures and treatments. Soft tissue specimens from the dorsal region encompassing the entire extent of the surgical wound were removed using a Bard-Parker scalpel with a 15C blade and blunt-tipped straight scissors. Care was taken to include a safety margin around the entire wound and cover the entire extent of the subcutaneous connective tissue in the area. The anatomical specimens obtained were labeled, placed in plastic cassettes, and immersed in a 4% formaldehyde solution in 0.1 M phosphate buffer (pH 7.4), remaining for a minimum period of 48 h. After this period, they were directed to the histological processing.
The removed specimens underwent routine laboratory procedures (washed for 24 h in running water, dehydrated in increasing concentrations of ethanol, cleared in xylene, and embedded in paraffin blocks). Semiserial sections with a thickness of 5 μm were made, with sections stained using the hematoxylin and eosin (H & E), and also submitted to immunohistochemical (IHC) analysis.
2.6. Clinical Assessment of Wounds
All wounds from each group were clinically evaluated by an experienced and blinded examiner (JMC). The following clinical conditions were observed: wound contraction, presence of visible blood vessels in the wound area, presence and characteristics of crust, edema, bleeding, and signs of inflammation and infection.
2.7. Descriptive Histology and Histometric Evaluation
Descriptive histological and histometric analyses from each group and time points were assessed by an expert histologist (EE) blinded to the experimental groups. Photographs of all wounds from each animal and group were taken at all experimental time points (0 h, 7, 14, and 21 days). Each animal was positioned in a ventral decubitus on a surgical table, ensuring that the wound remained as parallel as possible to the ground. A digital camera (Canon 60D, Japan) with a Macro 100 lens (Canon 60D, Japan) fixed on a stand was used to maintain a standardized distance between the wound surface and the camera lens. The images of each animal group were then processed using publicly available image processing software, Image J® (National Institutes of Health, USA). The wound area measurement was conducted by outlining the entire wound perimeter with the mouse cursor on the computer screen, and the obtained measurement was expressed in millimeters (mm).
For the histometric analysis, the characteristics of the epithelium were evaluated to quantitate the area occupied by epithelial tissue (AET) in the surgical wound. Subsequently, the surface occupied by the epithelial tissue (SET) was assessed. In the connective tissue in the central area of the wound, the amount of ICs expressed in percentage of the total area and the area occupied by collagen fibers (ACF) were quantitated. These measurements were performed by two experienced examiners (EMM and JMC) blinded to the experimental groups.
2.8. IHC Assessment
Tissue sections underwent the indirect immunoperoxidase method for the identification of VEGF and TGFβ-1 proteins. Antigen retrieval was achieved by immersing histological slides in a specific buffer (Diva decloaker®, Biocare Medical, Concord, CA, USA) in a pressurized chamber at 95°C for 20 min. Then, the histological slides were washed in 0.1 M phosphate-buffered saline (pH 7.4) and were immersed in 3% hydrogen peroxide (H2O2) for 1 h and in 1% bovine serum albumin. The slides were incubated with anti-VEGF anti-TGFβ-1 antibodies for 24 h. The revelation was performed using 3,3
2.9. Statistical Analysis
For the statistical analysis of clinical wound data, the IBM program (International Business Machines Corp., Armonk, New York, USA) was utilized. The Kolmogorov–Smirnov and Shapiro–Wilk tests were employed to assess data normality and sample distribution. Subsequently, a one-way analysis of variance (ANOVA) was applied, followed by the Games–Howell posttest for multiple parametric comparisons in the presence of heterogeneous variances, with a significance level set at 5% (
3. Results
3.1. Clinical Analysis
The wound evaluation followed a standardized approach, documenting all clinical evidence of repair, including contraction, visible blood vessels, presence and characteristics of crust, edema, bleeding, inflammation, and signs of infection (Figure 1). Immediately after creating the cutaneous wounds, all wounds showed sharp edges and a regular size, close to the diameter of the circular punch, demonstrating the effectiveness of the method for their creation. Capillaries were observed in the dermis, and there were no signs of significant bleeding, infection, or inflammation within the wounds (Figure 1). At 7 days postoperatively, wounds in the C group exhibited the presence of a dry crust, with signs of partial displacement at the wound margin along its entire length. Wounds in the OZ0.3, OZ0.7, and OZ1.0 groups showed a wet crust, with some areas beginning to detach at the margin, which was similar in clinical characteristics in these groups. By day 14 postoperatively, all wounds in all groups showed complete displacement of the crust; however, those in the C group exhibited a lower degree of contraction compared to the other groups. In the comparative analysis among wounds in all groups, those in the OZ0.7 and OZ1.0 groups demonstrated a more advanced state of healing, with greater contraction and a smaller area of redness in their central portion. At 21 days, all wounds showed a total absence of the crust, but wounds in the C and OZ0.3 groups still displayed a slightly reddish area in their central portion compared to those in the OZ0.7 and OZ1.0 groups, which demonstrated complete closure of the wound with scar formation. There were no wounds presenting with undesirable side effects such as bleeding or infection throughout the course of the study.
[figure(s) omitted; refer to PDF]
Regarding the degree of wound contraction, no statistically significant difference was detected among the experimental groups (Figure 1). At 14 and 21 days, the degree of contraction of surgical wounds was greater than at 7 days in all experimental groups (Figure 1).
3.2. Descriptive Histological Analysis
At 7 days postoperatively, the surgical wounds exhibited stratified squamous epithelium proliferating from their edges towards the center. A thin crust was present on the surface of the wound. The connective tissue underlying the epithelium/crust was intensely vascularized, with a large number of ICs and some fibroblasts, supported by a delicate network of collagen fibers (Figure 2).
[figure(s) omitted; refer to PDF]
By day 14 postoperatively, the epithelial tissue covered a large part of the wound. Remnants of the crust were still present in some specimens. The underlying connective tissue was intensely vascularized, with some ICs and abundant fibroblasts, supported by a moderate amount of collagen fibers (Figure 3).
[figure(s) omitted; refer to PDF]
At 21 days postoperatively, the epithelial tissue covered the entire surgical wound. The underlying connective tissue still showed significant vascularity, albeit with few ICs and numerous fibroblasts associated with a large amount of collagen fibers. At this stage, the initiation of the reorganization of cutaneous appendages, such as hair and glands, was observed (Figure 4).
[figure(s) omitted; refer to PDF]
When comparing groups within their respective experimental periods, it was noted that histological characteristics were very similar; however, the OZ1.0 group exhibited some histological aspects consistent with a more accelerated tissue repair process, such as the thickness and cellular pattern of the epidermis and the amount of collagen fibers in the dermis.
3.3. Histometric Analysis
Initially, the degree of reliability between the evaluators was assessed, obtaining an intraclass correlation coefficient (ICC) of 0.960. The intergroup analysis showed a statistically significant increase in the percentage of AET for the OZ0.7 and OZ1.0 groups compared to the C and OZ0.3 groups (Figure 5(a)). The OZ1.0 group showed the most promising effects, meaning that the O3 effects are dose-dependent (Figure 5(a)). Regarding the quantitation of SET, there were no differences among the experimental groups, but the epithelial coverage increased over time for all groups (Figure 5(b)). Furthermore, the quantification of ICs showed a significant increase in the initial periods of evaluation for all groups. The percentage of ICs decreased for all groups in the later periods of evaluation especially at 21 days. There were no differences regarding the percentage of ICs among groups (Figure 5(c)). The ACF was also evaluated. Our findings showed that the OZ1.0 group showed the most promising results regarding the amount of collagen fibers compared to the other groups (Figure 5(d)).
[figure(s) omitted; refer to PDF]
3.4. IHC Analysis
The IHC analysis of TGFβ1 revealed that in groups C, OZ0.3, and OZ0.7 at 21 days, there was higher immunolabeling compared to the 7-day period in the same group. In the OZ0.7 group at 14 days and the OZ1.0 group at 7 and 14 days, there was higher immunolabeling than in the C group during the same period. Additionally, in the OZ1.0 group at 7 and 14 days, there was higher immunolabeling than in the OZ0.3 group during the same period (Figure 6).
[figure(s) omitted; refer to PDF]
The IHC analysis of VEGF revealed higher immunolabeling in groups C and OZ0.3 at 14 and 21 days, compared to the 7-day period in the same group. Additionally, there was higher immunolabeling of VEGF in the OZ0.7 group at 7 days and the OZ1.0 group at 7 and 14 days, compared to the C group during the same period (Figure 7).
[figure(s) omitted; refer to PDF]
4. Discussion
The present study assessed, from clinical, histological, histometric, and IHC perspectives, the effects of systemically administered ozone and the influence of different dosages on the wound repair in surgical defects created in rats. The results of this study suggested a beneficial action of the O3/O2 mixture in stimulating wound repair in a dose-dependent manner, confirming our study hypothesis. From a clinical perspective, our data reveal that wounds treated with ozone displayed a dry crust, exhibiting greater contraction than the control wounds across all experimental time points, particularly notable in the OZ0.7 and OZ1.0 groups at 14 and 21 days. These clinical observations find support in histometric results, where wounds treated with the highest ozone dosage at 14 and 21 days demonstrated more significant contraction compared to the other groups.
The most commonly used method of ozone application is topical, whether using ozonized water, ozonized oil, gel formulations, or gas [33, 34, 42, 43]. Systemic administration is achieved through intramuscular injection [44], subcutaneous, and intravenous routes [45], with rectal insufflation being the preferred method in various studies [46]. However, intraperitoneal administration has been employed in the medical field for treating various pathologies [29, 47, 48]. Therefore, we choose to treat the animals with systemic administration of ozone to stimulate biological events in skin tissue repair.
Wound contraction is determined by specialized fibroblasts known as myofibroblasts, containing actin filaments, myosin, and smooth muscle actin (α-SMA) in their cytoplasm [49, 50]. Myofibroblasts possess twice the contractile strength of normal fibroblasts [51]. The differentiation of fibroblasts into myofibroblasts in the lesion area is primarily attributed to the activity of TGF-β1 [52], which is released in large quantities by platelets during bleeding from the surgical procedure. In the wound repair process, TGF-β1 is generated by leukocytes, macrophages, fibroblasts, and keratinocytes, influencing these cells to stimulate the infiltration of ICs, fibroplasia, matrix deposition, and angiogenesis in the area. The results of our study demonstrated a higher staining of TGF-β1 in both OZ0.7 (14 days) and OZ1.0 (7 and 14 days) groups compared to the C group. These findings suggest that the employed O3O2 in these groups was suitable to promote greater cellular differentiation, in a dosage-dependent manner.
The disruption of the vascular network during tissue injury and the high oxygen consumption by metabolically active cells in the area create an environment with reduced oxygen levels (hypoxia). In this hypoxic environment, important cytokines are produced, such as TNF-α, PDGF, TGF-β, and VEGF, which are crucial promoters of cell proliferation, migration, chemotaxis, and angiogenesis in wound healing [53, 54]. Our data revealed that the immunolabeling of VEGF was higher in the wounds of the OZ0.7 (7 days) and OZ1.0 (7 and 14 days) groups compared to the C group. This finding indicates the ability of the O3/O2 mixture used in these groups to promote greater angiogenesis in the injured area, resulting in benefits in tissue repair, especially in the early stages of the healing process. Studies have shown an increase in VEGF in the early stages of repair [55], paralleling observations from another preclinical study that observed an increase in VEGF and TGF-β1 between 5 and 7 days after topical application of ozonized oil in cutaneous wounds surgically created in the guinea pig [42].
The histological findings of this study revealed that the histological characteristics of the groups treated with O3 were very similar to each other. However, the OZ1.0 group showed some histological aspects consistent with a faster tissue repair process, such as the thickness and pattern of cellularity in the epidermis and the quantity of collagen fibers in the dermis. The area occupied by the epithelial tissue (AET) in the OZ0.7 (21 days) and OZ1.0 groups was increased when compared to the C group. This observation suggested that O3 has positive effects on cell differentiation and epithelial migration. Additionally, the area in the connective tissue occupied by collagen fibers was larger in the OZ0.7 group (14 days) and the OZ1.0 group (14 and 21 days) compared to the C group. The higher ozone dosage was capable of reducing the inflammatory infiltrate and activating collagen formation, resulting in differentiated epithelial migration. Our findings align with other studies demonstrating that ozone therapy induces biological effects at doses ranging between 20 and 50 μg/ml [56, 57].
Recent studies have developed membranes loaded with curcumin, chitosan, and sodium alginate using microwave technology to enhance wound-healing processes [58–60]. The findings of these studies indicated a notable increase in reepithelialization (
While our study methodology does not allow inferences regarding the mechanisms of action involved in this favorable biological response, it is known that O3 reacts rapidly with tissue fluids such as water and polyunsaturated fatty acids, resulting in the formation of ozonized lipid products and H2O2 [61]. In addition to the formation of the important free radical, when in contact with blood, O3 decomposes, giving rise to other reactive oxygen species (ROS) such as superoxide anion radical (O2-), hydroxyl radical (OH-), and nitric oxide (NO), which, in low concentrations, act as vasodilators and stimulate important endogenous growth factors [62]. Studies have also demonstrated that O3, upon reaching the cell membrane, forms ROS that activate the nuclear factor kappa-B (NF-κB) pathways, crucial signaling proteins for the cellular nucleus, consequently promoting the activation of intracellular pro and anti-inflammatory cytokine pathways [63].
It is important to bear in mind that our study has some limitations. First, we do not evaluate the toxicity of systemic administration of ozone. Secondly, systemic administration of ozone does not guarantee the same concentration at the local wound. Therefore, local treatment with ozone should be implemented in future studies. Finally, there is no description about the biological mechanisms by which ozone accelerates wound healing. Nevertheless, further research, particularly controlled randomized clinical studies with a substantial sample size, are warranted for the development of clinical usage protocols.
In conclusion, our findings suggest that the systemic use of O3 enhanced tissue healing of cutaneous wounds. The dosage plays a significant role in the biological response in a dose-dependent manner. Furthermore, the results of this study implicate the use of O3 for the treatment of skin wounds aiming at improving the healing process over time. Our findings suggest the use of O3 as a viable alternative to enhance wound healing and repair. Future research is indicated in the field and might be expanded to use O3 for treating palatal wounds after removal of connective tissue graft for root covering procedures, after tooth extraction to improve soft tissue healing and also to manage mucositis and peri-implantitis.
Author Contributions
J.M.C., E.E., E.M.M., L.H.T., L.E.M.P., E.L.P., R.S.d.M., and V.G.G.: conceptualization, investigation, editing, review, data curation, and formal analysis. R.S.d.M.: writing of original draft; V.G.G.: supervision. All authors read and approved the final version of the manuscript to be published.
Funding
Rafael Scaf de Molon is currently supported by a grant provided by the Sao Paulo Research Foundation–Fundacao de Amparo a Pesquisa do Estado de Sao Paulo (FAPESP) award #2023/15750-7.
Acknowledgments
Rafael Scaf de Molon is currently supported by a grant provided by the Sao Paulo Research Foundation–Fundacao de Amparo a Pesquisa do Estado de Sao Paulo (FAPESP) award #2023/15750-7.
[1] L. E. Lindley, O. Stojadinovic, I. Pastar, M. Tomic-Canic, "Biology and biomarkers for wound healing," Plastic and Reconstructive Surgery, vol. 138 no. 3S, pp. 18S-28S, DOI: 10.1097/PRS.0000000000002682, 2016.
[2] P. Martin, "Wound healing--aiming for perfect skin regeneration," Science, vol. 276 no. 5309, pp. 75-81, DOI: 10.1126/science.276.5309.75, 1997.
[3] A. J. Singer, R. A. Clark, "Cutaneous wound healing," The New England Journal of Medicine, vol. 341 no. 10, pp. 738-746, DOI: 10.1056/NEJM199909023411006, 1999.
[4] L. Uccioli, V. Izzo, M. Meloni, E. Vainieri, V. Ruotolo, L. Giurato, "Non-healing foot ulcers in diabetic patients: general and local interfering conditions and management options with advanced wound dressings," Journal of Wound Care, vol. 24, pp. 35-42, DOI: 10.12968/jowc.2015.24.Sup4b.35, 2015.
[5] A. S. Colwell, S. R. Beanes, C. Soo, C. Dang, K. Ting, M. T. Longaker, J. B. Atkinson, H. P. Lorenz, "Increased angiogenesis and expression of vascular endothelial growth factor during scarless repair," Plastic and Reconstructive Surgery, vol. 115 no. 1, pp. 204-212, DOI: 10.1097/01.PRS.0000138252.51581.22, 2005.
[6] S. A. Eming, "Evolution of immune pathways in regeneration and repair: recent concepts and translational perspectives," Seminars in Immunology, vol. 26 no. 4, pp. 275-276, DOI: 10.1016/j.smim.2014.09.001, 2014.
[7] S. A. Eming, P. Martin, M. Tomic-Canic, "Wound repair and regeneration: mechanisms, signaling, and translation," Science Translational Medicine, vol. 6 no. 265, article 265sr6,DOI: 10.1126/scitranslmed.3009337, 2014.
[8] S. Werner, R. Grose, "Regulation of wound healing by growth factors and cytokines," Physiological Reviews, vol. 83 no. 3, pp. 835-870, DOI: 10.1152/physrev.2003.83.3.835, 2003.
[9] I. Filipovic-Zore, Z. Divic, R. Duski, N. Gnjatovic, N. Galic, D. Prebeg, "Impact of ozone on healing after alveolectomy of impacted lower third molars," Saudi Medical Journal, vol. 32 no. 6, pp. 642-644, 2011.
[10] C. G. Nogales, P. H. Ferrari, E. O. Kantorovich, J. L. Lage-Marques, "Ozone therapy in medicine and dentistry," The Journal of Contemporary Dental Practice, vol. 9 no. 4, pp. 75-84, DOI: 10.5005/jcdp-9-4-75, 2008.
[11] O. A. El Meligy, N. M. Elemam, I. M. Talaat, "Ozone therapy in medicine and dentistry: a review of the literature," Dentistry Journal, vol. 11 no. 8,DOI: 10.3390/dj11080187, 2023.
[12] C. J. Randi, C. M. C. Heiderich, R. V. Serrano, S. Morimoto, L. O. C. de Moraes, L. Campos, L. F. Palma, "Use of ozone therapy in implant dentistry: a systematic review," Oral and Maxillofacial Surgery, vol. 28, pp. 39-49, DOI: 10.1007/s10006-023-01149-3, 2024.
[13] Y. Suh, S. Patel, R. Kaitlyn, J. Gandhi, G. Joshi, N. L. Smith, S. A. Khan, "Clinical utility of ozone therapy in dental and oral medicine," Medical Gas Research, vol. 9 no. 3, pp. 163-167, DOI: 10.4103/2045-9912.266997, 2019.
[14] A. A. AlMogbel, M. I. Albarrak, S. F. AlNumair, "Ozone therapy in the management and prevention of caries," Cureus, vol. 15 no. 4, article e37510,DOI: 10.7759/cureus.37510, 2023.
[15] S. S. El-Desouky, S. M. M. Omer, R. F. Ghouraba, R. Latif, I. A. Kabbash, S. M. Hadwa, "Zinc oxide-ozonated olive oil as a new root canal filling material in primary molars: a clinical randomized controlled trial," Clinical Oral Investigations, vol. 27 no. 12, pp. 7395-7405, DOI: 10.1007/s00784-023-05329-z, 2023.
[16] R. R. Talasani, S. P. Potharaju, B. Vijaya Lakshmi, Y. Durga Bai, R. K. Chintala, V. Mahankali, P. Koppolu, A. R. S. AlGhamdi, "Efficacy of ozonated water over chlorhexidine mouth rinse in chronic gingivitis patients - a comparative clinical study," The Saudi Dental Journal, vol. 34 no. 8, pp. 738-743, DOI: 10.1016/j.sdentj.2022.09.004, 2022.
[17] A. Butera, M. Pascadopoli, S. Gallo, C. Pérez-Albacete Martínez, J. E. Maté Sánchez de Val, L. Parisi, A. Gariboldi, A. Scribante, "Ozonized hydrogels vs. 1% chlorhexidine gel for the clinical and domiciliary management of peri-implant mucositis: a randomized clinical trial," Journal of Clinical Medicine, vol. 12 no. 4,DOI: 10.3390/jcm12041464, 2023.
[18] F. D’Ambrosio, M. Caggiano, A. Acerra, M. Pisano, F. Giordano, "Is ozone a valid adjuvant therapy for periodontitis and peri-implantitis? A systematic review," Journal of Personalized Medicine, vol. 13 no. 4,DOI: 10.3390/jpm13040646, 2023.
[19] L. F. Palma, C. Joia, L. Chambrone, "Effects of ozone therapy on periodontal and peri-implant surgical wound healing: a systematic review," Quintessence International, vol. 54 no. 2, pp. 100-110, DOI: 10.3290/j.qi.b3512007, 2023.
[20] A. Alsakr, K. Gufran, A. S. Alqahtani, M. Alasqah, B. Alnufaiy, H. G. Alzahrani, A. A. Alahmari, F. K. Alhumaidani, R. K. Alhumaidani, M. J. Althobiti, "Ozone therapy as an adjuvant in the treatment of periodontitis," Journal of Clinical Medicine, vol. 12 no. 22,DOI: 10.3390/jcm12227078, 2023.
[21] B. Rapone, E. Ferrara, E. Qorri, F. Inchingolo, G. Isola, P. Dongiovanni, G. M. Tartaglia, A. Scarano, "Research efficacy of gaseous ozone therapy as an adjuvant to periodontal treatment on oxidative stress mediators in patients with type 2 diabetes: a randomized clinical trial," BMC Oral Health, vol. 23 no. 1,DOI: 10.1186/s12903-023-02985-1, 2023.
[22] A. Scribante, S. Gallo, M. Pascadopoli, M. Frani, A. Butera, "Ozonized gels vs chlorhexidine in non-surgical periodontal treatment: a randomized clinical trial," Oral Diseases,DOI: 10.1111/odi.14829, 2023.
[23] G. Tete, T. D'Amicantonio, E. Polizzi, "Efficacy ozone therapy in reducing periodontal disease," Materials, vol. 16 no. 6,DOI: 10.3390/ma16062375, 2023.
[24] T. J. Lima Neto, L. A. Delanora, M. E. Sa Simon, K. H. Carmo Ribeiro, M. A. Matsumoto, M. J. Quirino Louzada, J. A. Shibli, E. Ervolino, L. P. Faverani, "Ozone improved bone dynamic of female rats using zoledronate," Tissue Engineering. Part C, Methods, vol. 30 no. 1,DOI: 10.1089/ten.tec.2023.0159, 2024.
[25] L. J. Varghese, B. Lahiri, N. V. Penumatsa, C. R. Soans, A. Sekar, F. A. Nasyam, "Effectiveness of topical ozone gel application in the management of postextraction wound healing: an in vivo study," The Journal of Contemporary Dental Practice, vol. 24 no. 11, pp. 887-890, DOI: 10.5005/jp-journals-10024-3557, 2023.
[26] F. Erdemci, Y. Gunaydin, M. Sencimen, I. Bassorgun, M. Ozler, S. Oter, A. Gulses, A. Gunal, S. Sezgin, G. R. Bayar, N. Dogan, I. K. Gider, "Histomorphometric evaluation of the effect of systemic and topical ozone on alveolar bone healing following tooth extraction in rats," International Journal of Oral and Maxillofacial Surgery, vol. 43 no. 6, pp. 777-783, DOI: 10.1016/j.ijom.2013.12.007, 2014.
[27] E. M. Miyasawa, E. Ervolino, J. M. Cardoso, L. H. Theodoro, G. R. C. Silveira, R. S. Molon, L. Levin, V. G. Garcia, L. E. Padovan, "Effects of systemic ozone administration on the fresh extraction sockets healing: a histomorphometric and immunohistochemical study in rats," Journal of Applied Oral Science, vol. 32, article e20230412,DOI: 10.1590/1678-7757-2023-0412, 2024.
[28] B. B. Loureiro, P. Juber, A. A. Souza, E. M. Cezario, J. C. Lima-Junior, K. Fontes, I. F. Soares, E. P. Zuza, "Application of highly ozonated sunflower oil does not improve palatal wound healing: a randomized controlled clinical trial," Journal of Periodontology, vol. 94 no. 7, pp. 858-867, DOI: 10.1002/JPER.22-0693, 2023.
[29] A. Hesham, M. Abass, H. Abdou, R. Fahmy, M. M. Rashad, A. A. Abdallah, W. Mossallem, I. F. Rehan, A. Elnagar, F. Zigo, S. Ondrašovičová, A. F. Abouelnaga, A. Rizk, "Ozonated saline intradermal injection: promising therapy for accelerated cutaneous wound healing in diabetic rats," Frontiers in Veterinary Science, vol. 10, article 1283679,DOI: 10.3389/fvets.2023.1283679, 2023.
[30] G. Y. Ozturk, B. F. Kocyigit, "Healing refractory livedoid vasculopathy-related skin ulcers by ozone therapy: a case-based review," Rheumatology International, vol. 44 no. 2, pp. 369-377, DOI: 10.1007/s00296-023-05504-1, 2024.
[31] A. Roth, A. Krishnakumar, R. Rahimi, "Ozone as a topical treatment for infected dermal wounds," Frontiers in Bioscience-Elite, vol. 15 no. 2,DOI: 10.31083/j.fbe1502009, 2023.
[32] J. Y. Lu, X. Q. Wang, Z. B. Fu, L. H. Gao, H. Mannam, Y. P. Xiang, Y. Y. Joo, J. R. Zeng, D. Wang, A. S. Paller, "Topical ozone accelerates diabetic wound healing by promoting re-epithelialization through the activation of IGF1R-EGFR signaling," The Journal of Investigative Dermatology, vol. 143 no. 12, pp. 2507-2514.e6, DOI: 10.1016/j.jid.2023.05.015, 2023.
[33] P. V. Patel, S. Kumar, G. D. Vidya, A. Patel, J. C. Holmes, V. Kumar, "Cytological assessment of healing palatal donor site wounds and grafted gingival wounds after application of ozonated oil: an eighteen-month randomized controlled clinical trial," Acta Cytologica, vol. 56 no. 3, pp. 277-284, DOI: 10.1159/000336889, 2012.
[34] D. J. Romary, S. A. Landsberger, K. N. Bradner, M. Ramirez, B. R. Leon, "Liquid ozone therapies for the treatment of epithelial wounds: a systematic review and meta-analysis," International Wound Journal, vol. 20 no. 4, pp. 1235-1252, DOI: 10.1111/iwj.13941, 2023.
[35] D. Torul, M. M. Omezli, T. Avci, "Investigation of the clinical efficacy of CGF and ozone in the management of alveolar osteitis: a randomized controlled trial," Clinical Oral Investigations, vol. 27 no. 8, pp. 4521-4529, DOI: 10.1007/s00784-023-05074-3, 2023.
[36] C. Kilkenny, W. J. Browne, I. C. Cuthill, M. Emerson, D. G. Altman, "Improving bioscience research reporting: the ARRIVE guidelines for reporting animal research," PLoS Biology, vol. 8 no. 6, article e1000412,DOI: 10.1371/journal.pbio.1000412, 2010.
[37] E. K. Simsek, F. Sahinturk, E. Gul, M. Tepeoglu, C. Araz, B. Haberal, "Effect of ozone therapy on epidural fibrosis in rats," World Neurosurgery, vol. 175, pp. e296-e302, DOI: 10.1016/j.wneu.2023.03.075, 2023.
[38] K. Uslu, H. D. Tansuker, A. Tabaru, S. E. Egeren, K. K. Kulahci, P. Bulut, F. Emre, M. F. Oktay, "Investigation of the effects of thrombocyte-rich plasma, systemic ozone and hyperbaric oxygen treatment on intraoral wound healing in rats: experimental study," European Archives of Oto-Rhino-Laryngology, vol. 277 no. 6, pp. 1771-1777, DOI: 10.1007/s00405-020-05872-5, 2020.
[39] B. Kan, M. Sencimen, G. R. Bayar, P. Korkusuz, A. T. Coskun, A. Korkmaz, E. Bahador, T. Zerener, "Histomorphometric and microtomographic evaluation of the effects of hyperbaric oxygen and systemic ozone, used alone and in combination, on calvarial defect healing in rats," Journal of Oral and Maxillofacial Surgery, vol. 73 no. 6, pp. 1231.e1-1231.e10, DOI: 10.1016/j.joms.2015.02.018, 2015.
[40] Y. J. Kim, R. S. de Molon, V. C. da Silva, M. C. A. da Veiga Conrado, L. C. Spolidorio, M. C. A. Roque-Barreira, J. A. Cirelli, "Topical application of lectin Artin M improves wound healing in defects created in the palatal mucosa: an in vivo study in dogs," Odontology, vol. 108 no. 4, pp. 560-568, DOI: 10.1007/s10266-020-00495-y, 2020.
[41] E. Ervolino, C. Statkievicz, L. F. Toro, J. M. de Mello-Neto, T. P. Cavazana, J. P. M. Issa, R. C. M. Dornelles, J. M. de Almeida, M. J. H. Nagata, R. Okamoto, C. A. Casatti, V. G. Garcia, L. H. Theodoro, "Antimicrobial photodynamic therapy improves the alveolar repair process and prevents the occurrence of osteonecrosis of the jaws after tooth extraction in senile rats treated with zoledronate," Bone, vol. 120, pp. 101-113, DOI: 10.1016/j.bone.2018.10.014, 2019.
[42] H. S. Kim, S. U. Noh, Y. W. Han, K. M. Kim, H. Kang, H. O. Kim, Y. M. Park, "Therapeutic effects of topical application of ozone on acute cutaneous wound healing," Journal of Korean Medical Science, vol. 24 no. 3, pp. 368-374, DOI: 10.3346/jkms.2009.24.3.368, 2009.
[43] A. Materni, C. Pasquale, E. Longo, M. Frosecchi, S. Benedicenti, M. Bozzo, A. Amaroli, "Prevention of dry socket with ozone oil-based gel after inferior third molar extraction: a double-blind split-mouth randomized placebo-controlled clinical trial," Gels, vol. 9 no. 4,DOI: 10.3390/gels9040289, 2023.
[44] A. de Sire, N. Marotta, M. Ferrillo, F. Agostini, C. Sconza, L. Lippi, S. Respizzi, A. Giudice, M. Invernizzi, A. Ammendolia, "Oxygen-ozone therapy for reducing pro-inflammatory cytokines serum levels in musculoskeletal and temporomandibular disorders: a comprehensive review," International Journal of Molecular Sciences, vol. 23 no. 5,DOI: 10.3390/ijms23052528, 2022.
[45] R. J. Rowen, "Ozone therapy in conjunction with oral antibiotics as a successful primary and sole treatment for chronic septic prosthetic joint: review and case report," Medical Gas Research, vol. 8 no. 2, pp. 67-71, DOI: 10.4103/2045-9912.235139, 2018.
[46] Q. Wen, D. Liu, X. Wang, Y. Zhang, S. Fang, X. Qiu, Q. Chen, "A systematic review of ozone therapy for treating chronically refractory wounds and ulcers," International Wound Journal, vol. 19 no. 4, pp. 853-870, DOI: 10.1111/iwj.13687, 2022.
[47] F. Mete, H. Tarhan, O. Celik, I. Akarken, K. Vural, R. G. Ekin, I. Aydemir, Y. O. Ilbey, "Comparison of intraperitoneal and intratesticular ozone therapy for the treatment of testicular ischemia-reperfusion injury in rats," Asian Journal of Andrology, vol. 19 no. 1, pp. 43-46, DOI: 10.4103/1008-682X.171570, 2017.
[48] S. Y. Lin, J. Ma, J. X. An, X. Y. Qian, Y. Wang, D. K. Cope, J. P. Williams, "Ozone inhibits APP/A β production and improves cognition in an APP/PS1 transgenic mouse model," Neuroscience, vol. 418, pp. 110-121, DOI: 10.1016/j.neuroscience.2019.07.027, 2019.
[49] J. F. Reger, M. K. Dabbous, "A comparative fine structure study on myofibroblasts from a cultured human and an in-situ rat tumor source," Journal of Submicroscopic Cytology and Pathology, vol. 20 no. 3, pp. 501-508, 1988.
[50] G. Majno, G. Gabbiani, B. J. Hirschel, G. B. Ryan, P. R. Statkov, "Contraction of granulation tissue in vitro: similarity to smooth muscle," Science, vol. 173 no. 3996, pp. 548-550, DOI: 10.1126/science.173.3996.548, 1971.
[51] L. K. Wrobel, T. R. Fray, J. E. Molloy, J. J. Adams, M. P. Armitage, J. C. Sparrow, "Contractility of single human dermal myofibroblasts and fibroblasts," Cell Motility and the Cytoskeleton, vol. 52 no. 2, pp. 82-90, DOI: 10.1002/cm.10034, 2002.
[52] Y. Tai, E. L. Woods, J. Dally, D. Kong, R. Steadman, R. Moseley, A. C. Midgley, "Myofibroblasts: function, formation, and scope of molecular therapies for skin fibrosis," Biomolecules, vol. 11 no. 8,DOI: 10.3390/biom11081095, 2021.
[53] P. G. Rodriguez, F. N. Felix, D. T. Woodley, E. K. Shim, "The role of oxygen in wound healing: a review of the literature," Dermatologic Surgery, vol. 34 no. 9, pp. 1159-1169, DOI: 10.1111/j.1524-4725.2008.34254.x, 2008.
[54] S. Schreml, R. M. Szeimies, L. Prantl, S. Karrer, M. Landthaler, P. Babilas, "Oxygen in acute and chronic wound healing," The British Journal of Dermatology, vol. 163 no. 2, pp. 257-268, DOI: 10.1111/j.1365-2133.2010.09804.x, 2010.
[55] M. Detmar, L. F. Brown, B. Berse, R. W. Jackman, B. M. Elicker, H. F. Dvorak, K. P. Claffey, "Hypoxia regulates the expression of vascular permeability factor/vascular endothelial growth factor (VPF/VEGF) and its receptors in human skin," The Journal of Investigative Dermatology, vol. 108 no. 3, pp. 263-268, DOI: 10.1111/1523-1747.ep12286453, 1997.
[56] S. V. Naik, K. Rajeshwari, S. Kohli, S. Zohabhasan, S. Bhatia, "Ozone- a biological therapy in dentistry-reality or myth?????," The Open Dentistry Journal, vol. 10 no. 1, pp. 196-206, DOI: 10.2174/1874210601610010196, 2016.
[57] P. T. Pires, J. C. Ferreira, S. A. Oliveira, M. J. Silva, P. R. Melo, "Effect of ozone gas on the shear bond strength to enamel," Journal of Applied Oral Science, vol. 21 no. 2, pp. 177-182, DOI: 10.1590/1678-7757201302362, 2013.
[58] H. A. Albarqi, A. A. Alqahtani, I. Ullah, N. R. Khan, H. M. Basit, T. Iftikhar, A. Wahab, M. Ali, M. Badar, "Microwave-assisted physically cross-linked chitosan-sodium alginate hydrogel membrane doped with curcumin as a novel wound healing platform," AAPS PharmSciTech, vol. 23 no. 2,DOI: 10.1208/s12249-022-02222-y, 2022.
[59] H. Rostami, R. Mohammadi, S. Asri-Rezaei, A. Tehrani, "Evaluation of application of chitosan/nano selenium biodegradable film on full thickness excisional wound healing in rats," Iranian Journal Of Veterinary Surgery, vol. 13 no. 1, 2018.
[60] Z. Wang, Z. Zhao, N. R. Khan, Z. Hua, J. Huo, Y. Li, "Microwave assisted chitosan-polyethylene glycol hydrogel membrane synthesis of curcumin for open incision wound healing," Pharmazie, vol. 75 no. 4, pp. 118-123, DOI: 10.1691/ph.2020.9179, 2020.
[61] V. Bocci, C. Aldinucci, "Biochemical modifications induced in human blood by oxygenation-ozonation," Journal of Biochemical and Molecular Toxicology, vol. 20 no. 3, pp. 133-138, DOI: 10.1002/jbt.20124, 2006.
[62] D. Roy, P. K. Wong, R. S. Engelbrecht, E. S. Chian, "Mechanism of enteroviral inactivation by ozone," Applied and Environmental Microbiology, vol. 41 no. 3, pp. 718-723, DOI: 10.1128/aem.41.3.718-723.1981, 1981.
[63] S. L. Manoto, M. J. Maepa, S. K. Motaung, "Medical ozone therapy as a potential treatment modality for regeneration of damaged articular cartilage in osteoarthritis," Saudi Journal of Biological Sciences, vol. 25 no. 4, pp. 672-679, DOI: 10.1016/j.sjbs.2016.02.002, 2018.
You have requested "on-the-fly" machine translation of selected content from our databases. This functionality is provided solely for your convenience and is in no way intended to replace human translation. Show full disclaimer
Neither ProQuest nor its licensors make any representations or warranties with respect to the translations. The translations are automatically generated "AS IS" and "AS AVAILABLE" and are not retained in our systems. PROQUEST AND ITS LICENSORS SPECIFICALLY DISCLAIM ANY AND ALL EXPRESS OR IMPLIED WARRANTIES, INCLUDING WITHOUT LIMITATION, ANY WARRANTIES FOR AVAILABILITY, ACCURACY, TIMELINESS, COMPLETENESS, NON-INFRINGMENT, MERCHANTABILITY OR FITNESS FOR A PARTICULAR PURPOSE. Your use of the translations is subject to all use restrictions contained in your Electronic Products License Agreement and by using the translation functionality you agree to forgo any and all claims against ProQuest or its licensors for your use of the translation functionality and any output derived there from. Hide full disclaimer
Copyright © 2024 Jânderson de Medeiros Cardoso et al. This is an open access article distributed under the Creative Commons Attribution License (the “License”), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. Notwithstanding the ProQuest Terms and Conditions, you may use this content in accordance with the terms of the License. https://creativecommons.org/licenses/by/4.0/
Details
; Ervolino, Edilson 2
; Miyasawa, Erton Massamitsu 1 ; Theodoro, Leticia Helena 3
; Marques Padovan, Luis Eduardo 1
; Estevão Lopes Pereira 2
; Rafael Scaf de Molon 3
; Valdir Gouveia Garcia 1
1 Department of Implantology Latin American Institute of Dental Research and Teaching (ILAPEO) Curitiba PR Brazil
2 Department of Basic Sciences School of Dentistry São Paulo State University (UNESP) Araçatuba São Paulo Brazil
3 Department of Diagnostic and Surgery School of Dentistry São Paulo State University (UNESP) Araçatuba São Paulo Brazil