Open Access

The molecular basis of hypertrophic scars

Burns & Trauma20164:2

https://doi.org/10.1186/s41038-015-0026-4

Received: 29 July 2015

Accepted: 30 December 2015

Published: 21 January 2016

Abstract

Hypertrophic scars (HTS) are caused by dermal injuries such as trauma and burns to the deep dermis, which are red, raised, itchy and painful. They can cause cosmetic disfigurement or contractures if craniofacial areas or mobile region of the skin are affected. Abnormal wound healing with more extracellular matrix deposition than degradation will result in HTS formation. This review will introduce the physiology of wound healing, dermal HTS formation, treatment and difference with keloids in the skin, and it also review the current advance of molecular basis of HTS including the involvement of cytokines, growth factors, and macrophages via chemokine pathway, to bring insights for future prevention and treatment of HTS.

Keywords

Hypertrophic scars Animal model Cytokines Growth factors Macrophages Stromal cell-derived factor 1/CXCR4 signaling

Background

Hypertrophic scars (HTS) are considered to be a dermal form of fibroproliferative disorders that are caused by aberrant wound healing due to injuries to the deep dermis, including burn injury, laceration, abrasions, surgery and trauma. HTS are red, raised, rigid and can cause pruritus, pain and joint contracture. HTS formed in the facial area can cause cosmetic disfigurement, which result in psychological and social issues [1, 2] (Fig. 1).
Fig. 1

Patients with HTS. A 24 year-old white man, 11 months after a 21 % TBSA burn. This patient developed HTS, resulting in cosmetic and functional problems that included restricted opening of mouth and tight web spaces of fingers that limited range of motion on hands (From Tredget EE, Levi B, Donelan MB. Biology and principles of scar management and burn reconstruction. Surg Clin North Am. 2014 Aug;94(4):793–815. With permission)

The physiology of wound healing in the skin

Wound healing can be divided into four stages: hemostasis, inflammation, proliferation and tissue remodeling [3]. In these four stages, there are complicated interactions within a complex network of pro-fibrotic and anti-fibrotic molecules, such as growth factors, proteolytic enzymes and extracellular matrix (ECM) proteins [4, 5].

The first stage is hemostasis, which relates to the clotting cascade and the formation of a provisional wound matrix. These changes occur immediately after injury and are completed within hours [6]. Clotting factors from the injured skin (extrinsic system) and aggregation of thrombocytes or platelets after exposure to collagen fibers (intrinsic system) are activated. The exposed collagen also triggers platelets to begin secreting cytokines and growth factors [7]. The provisional wound matrix serves as a scaffold structure for the migration of leukocytes, keratinocytes, fibroblasts and endothelial cells. Platelets induce the vasoconstriction in order to reduce blood loss followed by secretion of a number of inflammatory factors including serotonin, bradykinin, prostaglandins and most importantly histamine, which activate the inflammatory phase.

In the inflammatory phase, polymorphonuclear neutrophils (PMNs) are the first inflammatory cells that are recruited to the inflamed site and are present there for 2–5 days. Several mediators such as tumor necrosis factor-alpha (TNF-α), interleukin-1 beta (IL-1β) and interleukin-6 (IL-6) are released by the neutrophils in order to amplify the inflammatory response [8]. Monocytes are attracted by the inflammatory mediators and differentiate into macrophages soon after they migrate into the wound site. The main functions of macrophages are phagocytosis of pathogens and cell debris as well as the release of growth factors, chemokines and cytokines which will push the wound healing process into the next stage.

The proliferation stage consists of angiogenesis, re-epithelialization, and granulation tissue formation. The process of angiogenesis is commenced by growth factors such as vascular endothelial growth factor (VEGF) released by activated endothelial cells from uninjured blood vessels. The new blood vessels differentiate into arteries and venules by recruitment of pericytes and smooth muscle cells [9]. Re-epithelialization is essential for the re-establishment of tissue integrity, which is ensured by local keratinocytes at the wound edges and epithelial stem cells from skin appendages such as hair follicles or sweat glands [10]. Granulation tissue formation is the last step in the proliferation phase, characterized by accumulation of a high density of fibroblasts, granulocytes, macrophages, capillaries and collagen bundles, which replace the provisional wound matrix formed during the inflammation stage. The predominant cells in this tissue are fibroblasts, which produce types I and III collagen and ECM substances, providing a structural framework for cell adhesion and differentiation [11]. Later, myofibroblasts induce wound contraction by virtue of their multiple attachment points to collagen and help to reduce the surface area of the scar [12].

The remodeling stage is already initiated while the granulation tissue is formed. During the maturation of the wound, type III collagen, which was produced in the proliferation stage, is replaced by the stronger type I collagen which is oriented as small parallel bundles and contributes to the basket-weave collagen formation in normal dermis [13].

HTS formation

The physiological process of normal wound healing will not result in HTS formation. However, if abnormalities occur during the wound healing process, the delicate balance of ECM degradation and deposition will be disrupted. Either insufficient degradation and remodeling of ECM due to an imbalance in expression of matrix metalloproteinases (MMPs) [14] or excessive ECM deposition caused by increased activity of fibroblasts and myofibroblasts [15] might lead to HTS formation. One common mechanism that burn patients often end up with HTS formation is the chronic inflammation or infection due to the severity of the injury, which prolongs the wound healing process and leads to excessive scarring [16]. This prolonged inflammatory phase will lead to HTS formation such as increased vessel and cell number as well as excessive collagen deposition [17].

It is well accepted that fibroblasts and myofibroblasts play essential roles in fibrotic diseases due to their abilities to generate excessive collagen in abnormal wound healing conditions [18, 19]. However, growing evidence suggests that other cells actively participate in scar pathogenesis, for example, keratinocytes and mast cells [20, 21]. When co-cultured with keratinocytes, fibroblasts exhibited significant proliferation activity [22]. The proliferation of dermal fibroblasts can also be stimulated by intercommunication of epidermal keratinocytes while decreasing the collagen production [23]. The activated keratinocytes in HTS tissue showed abnormal epidermal-mesenchymal interactions due to delayed re-epithelialization and prolonged epidermal inflammation, indicating that abnormal wound healing such as severe burn injuries may end up with HTS formation because the regulation of keratinocytes to fibroblasts is impaired [24]. However, independently co-culturing layered fibroblasts and keratinocytes on collagen-glycosaminoglycan scaffolds, aiming to assess the influence of keratinocytes and layered fibroblasts on the characteristics of tissue-engineered skin, showed that keratinocytes reduced fibrotic remodeling of the scaffolds by deep dermal fibroblasts, demonstrating an anti-fibrotic role of keratinocytes on layered fibroblasts in a 3D microenvironment [25]. In addition mast cells appear to activate fibroblasts through gap junction intercellular communication (GJIC), indicating that mast cell-fibroblast GJIC may also play a role in fibrosis [26]. Eliminating the mast cell or its GJIC with fibroblasts may prevent HTS formation or reduce the severity of fibrosis [27]. Mast cells are able to stimulate the proliferation of fibroblasts by releasing biological mediators such as histamine, chymase and tryptase via degranulation, which leads to the promotion of fibrogenesis [28, 29]. Additionally, histamine is able to enhance the effect on fibroblast migration and proliferation in vitro [30]. More histamine was found in HTS mast cells compared to normal skin mast cells after stimulation by a neuropeptide, substance P [31]. In an in vivo experiment, histamine was found significantly elevated in the plasma of patients with HTS compared to age-matched normal volunteers [32]. The elevated histamine can cause vasodilation and itchiness, resulting in the typical pruritic behavior that severely affects patients with HTS [33].

Differences between HTS and keloids

HTS and keloids are both caused by abnormal wound healing and are characterized by pathologically excessive fibrosis in the skin [34]. Sometimes the differentiation between HTS and keloids can be difficult and lead to incorrect identification, which may result in inappropriate treatment [35].

HTS are mostly caused by trauma or burn injury to the deep dermis and do not extend beyond the boundary of the original injury. Keloids can develop after minor injuries and may even spontaneously form on the sternal region without obvious injury, which will project beyond the original wound borders [36, 37]. HTS are red, raised and mostly linear scar occurred in any regions of the body while keloids appear as pink to purple, shiny, rounded protuberances and are commonly seen in sternal skin, shoulder, upper arms and earlobe. HTS usually appear within a few months of injury, regress in one or a few years and can cause contracture when joint regions are affected, whereas, keloids might take years to develop, grow for years and do not cause contracture. Keloids are commonly seen in darker skin population and have never been reported in albino populations [38].

HTS are characterized by abundant alpha-smooth muscle actin (α-SMA) producing myofibroblasts together with more type III collagen than type I collagen. On the contrary, there is no α-SMA producing myofibroblasts and a mixture of type I and type III collagen is found in keloid tissue [34]. The collagen bundles in keloids are thick, large and closely packed random to epidermis, whereas fine, well-organized parallel to epidermis collagen bundles are found in HTS [39]. ATP in keloids remained at higher levels for a long time while ATP level decreased over time in HTS [40]. An investigation of the expression of three proteins of the p53 family in keloids and HTS showed that the level of p53 proteins was higher in keloids compared to HTS. Protein p73 was elevated only in HTS and no difference was found between keloids and HTS of the level of p63 [41]. An in vitro analysis of ECM contraction by fibroblasts isolated from different scars showed that HTS fibroblasts had a consistently higher basal level of fibrin matrix gel contraction than keloid fibroblasts [42]. Despite all these differences, HTS and keloids possess similar features including excessive ECM deposition such as high collagen content and rich proteoglycan levels within the dermis and subcutaneous tissue [43]. The treatment for HTS and keloids are similar but HTS has a better prognosis for surgical excision because keloids have a much higher recurrence rates [16].

Complications of HTS

Complications of HTS include pain, pruritus, immobility of joint region, disfigurement and psychological issues. Pain and pruritus might not be as devastating as other complications, but they are significant complaints for many patients with HTS and they have been shown to persist for decades. The pain patients with HTS experience is often neuropathic pain, which is caused by dysfunction in the peripheral or central nervous system due to the primary injury. The neuropathic pain symptoms complained by patients with HTS are pins and needles, burning, stabbing, shooting or electric sensations [44]. The mechanism of pruritus is not well understood, but it is associated with histamine, which is released by mast cells and implicated as a primary mediator of itchiness [45]. Patients who developed HTS also suffer from reduced functional range of motion due to joint contractures, and disfigurement due to HTS tissue formed in the visible area of the body, which can lead to psychological problems or even social issues. A cross-sectional descriptive study showed that patients with HTS suffered from pain, joint stiffness, handicaps in walking or running up to on average of 17 years since the severe burn injury [46]. With all these complications, patients with HTS have complicated psychiatric disorders, including concern of body image, anxiety, depression, low self-esteem and posttraumatic stress. They have needs for psychological counseling and rehabilitation, especially for those who are economically disadvantaged or with preexisting mental illness [47]. However, a study focused on adolescent with disfiguring burn scars showed that instead of viewing themselves as less personally competent than unburned adolescents, they exhibited a similar or higher degree of self-worth as compared to their peers [48].

Treatments of HTS

The outcome of HTS is quite different because of the varied injured sites, severity of the injuries, and treatments the patients receive which leads to a variety of therapeutic strategies between surgeons and hospitals [49]. The effect of current treatment of HTS is slow and incomplete while being expensive, time consuming and labor intensive. In 2002, Mustoe et al. reported a qualitative overview of the available clinical literature by an international advisory panel of experts and provided evidence-based recommendations on prevention and treatment of HTS, which was considered as an outline for scar management [50]. Surgical excision combined with adjuvant therapies such as steroids, pressure garments and silicone gel is still the most common current management [51]. There are similar studies published in 2014 by Gold et al. [52, 53], which tried to standardize scar management by establishing safe and effective treatment options in order to apply in routine clinical practice. They conducted a comprehensive search of the MEDLINE database over the past 10 years and suggested that the most significant advances were laser therapy [54] and 5-fluorouracil [55]. Emerging therapies for HTS were also reported such as bleomycin [56], onion extract gel [57, 58], and Botulinum toxin A [59].

Review

Molecular basis of HTS

Cytokines in HTS formation

Interleukin-1 alpha (IL-1α) and TNF-α inhibit HTS

Interleukin-1 (IL-1) has two subtypes, IL-1α and IL-1β. IL-1α was found to promote the release of MMPs, activate MMP-1 and stimulate the degradation of ECM [60, 61]. Thus, decreased levels of IL-1α may lead to ECM accumulation and HTS. The expression of IL-1α was found significantly lower in HTS than in normal skin from patients following breast reduction surgery [62]. Quite different from IL-1α, IL-1β is found to be over-expressed in HTS compared to normal skin [63].

TNF-α participates in the early inflammation stage and the ECM remodeling phase. TNF-α is also believed to cause fibrosis together with IL-1β [64]. However, TNF-α expression was shown to be decreased in HTS compared to normal skin, which indicated that TNF-α may be important for wound healing and HTS might be partially a consequence of a decreased amount of TNF-α [65]. Another experiment demonstrated that TNF-α could suppress transforming growth factor beta-1 (TGF-β1)-induced myofibroblasts phenotypic genes such as α-SMA at the mRNA level as well as at the Smad signaling pathway of TGF-β1 [66].

Inappropriate release of IL-6 leads to HTS

IL-6 is also involved in the wound healing process. It is one of the major regulators of cells stimulation, angiogenesis and ECM synthesis [67]. IL-6 could also cause fibrotic diseases such as pulmonary fibrosis and scleroderma [68, 69]. In addition, IL-6 was reported to be highly expressed in fibroblasts from HTS tissue compared to normal fibroblasts, influencing scar formation by modulating fibroblasts [70]. In order to further investigate the function of IL-6, fibroblasts from HTS were treated with IL-6. Results showed an absence of any up-regulation of MMP-1 and MMP-3, indicating that suppression of MMPs may play a role in the excessive accumulation of collagen formed in HTS [71]. In fetal fibroblasts, there was less IL-6 produced compared to adult fibroblasts and the addition of exogenous IL-6 caused scar formation instead of scarless wound healing [72]. However, IL-6 knock-out mice showed delayed wound healing [73].

Interleukin-10 (IL-10) plays an important role in scarless wound healing by regulating pro-inflammatory cytokines

IL-10 is produced by T helper cells and it could mediate the growth or functions of various immune cells including T cells and macrophages. It has been established that IL-10 acts as a key anti-inflammatory cytokine, which could limit or terminate the inflammatory processes [74]. Neutralizing antibodies of IL-10 were administered into incisional wounds in mice and the results demonstrated an inhibited infiltration of neutrophils and macrophages and an over-expression of monocyte chemotactic protein-1 (MCP-1), IL-1β, TNF-α [75] and IL6 [76]. This is supported by another study that IL-10 significantly inhibited lipopolysaccharide (LPS)-induced IL-6 production at a transcriptional level [77]. A study tried to evaluate whether IL-10 could change the innervated conditions of full thickness excisional wounds created on the dorsal surface of CD1 mice. The results showed only temporary changes during the wound healing process but no significant changes at 84 days after treatment. However, wounds treated with IL-10 recovered similarly to normal skin compared to the wounds treated with PBS [78]. Another experiment reported that scar appeared in IL-10 knockout fetal mice compared to scarless wound healing in the control group [79]. A more recent study showed that IL-10 could provide an optimal environment for fetal and postnatal scarless wound healing [80]. A similar study also over-expressed IL-10 but in adult murine wounds. The results showed that increased IL-10 reduced inflammation, collagen deposition and created improved wound healing conditions [81].

Growth factors in HTS formation

Transforming growth factor-β (TGF-β) plays a pivotal role in HTS formation

TGF-β is one of the most important growth factors that regulate tissue regeneration, cell differentiation, embryonic development and regulation of the immune system [8284]. Recent studies showed that TGF-β not only involves in normal wound healing process but also contributes to fibroproliferative disorders such as pulmonary fibrosis [85] and HTS [86]. TGF-β has three isoforms, TGF-β1, transforming growth factor-beta 2 (TGF-β2) and transforming growth factor-beta 3 (TGF-β3) [87]. Shah et al. used the neutralizing antibody to TGF-β1 and TGF-β2 in cutaneous wounds of adult rodents and found reduced cutaneous scarring formation [88]. A subsequent study from Shah reported that exogenous addition of TGF-β3 to cutaneous rat wounds reduced scarring, indicating that TGF-β1 and TGF-β2 were related to cutaneous scarring while TGF-β3 should be considered as a therapeutic agent against scarring [89]. A more recent study treated the rabbit ear wounds with anti-TGF-β1, 2, 3 monoclonal antibodies at different time points of wound healing and early injection of antibodies showed delayed wound healing while the injections of middle or later time points remarkably reduced HTS formation, which implicated the indispensable roles of TGF-β1, 2, 3 in early stage of wound healing [90]. The transcriptional factor forkhead box protein O1 (FOXO1) has recently been found to be important as a regulator in wound healing. It exerts its effect through regulation of TGF-β1 expression from oxidative stress. The absence of FOXO1 reduced TGF-β1 expression and led to impaired re-epithelialization of wounds [91].

Many studies indicate that aberrant TGF-β expression plays a pivotal role in HTS formation. For example, a previous study showed that the serum level of TGF-β1 was up-regulated locally and systemically in burn patients and a significant clinical improvement in scar quality and volume was obtained after interferon-alpha2b (IFN-α2b) therapy, which was associated with normalization of serum TGF-β1 [92]. Treatment of IFN-α2b and interferon-gamma (IFN-γ) to site-matched HTS and normal fibroblasts showed antagonized TGF-β1 protein production, down-regulation of TGF-β1 mRNA levels [93]. Tredget et al. made superficial partial-thickness ear wound and full-thickness back wounds on a transgenic mouse over-expressing TGF-β1 in order to investigate the endogenous derived TGF-β1 on wound re-epithelialization. The findings suggested that over-expression of TGF-β1 speeded the rate of wound closure in partial-thickness wounds; whereas, over-expression of TGF-β1 slowed the rate of wound re-epithelialization in full-thickness wounds [94]. Another study created superficial and deep horizontal dermal scratch experimental wounds on the anterior thigh of adult male patients in order to characterize the related expression of TGF-β1 and TGF-β3. HTS formed after injuries to the deep dermis while superficial wounds healed with minimal or no scarring. Higher TGF-β1 and lower TGF-β3 expression was found in deep wounds compared to superficial wounds, suggesting the pivotal role of TGF-β1 in HTS formation [95].

Connective tissue growth factor (CTGF) acts as a downstream mediator of TGF-β1 signaling pathway and involves in HTS formation

CTGF, also know as CCN2, is a pleiotropic cytokine that is induced by TGF-β1 in dermal fibroblasts and is considered to be a downstream mediator of TGF-β1 [96]. The main role of CTGF is to interact with signaling proteins such as TGF-β1 for the regulation of cell proliferation, differentiation, adhesion, ECM production and granulation tissue formation [97, 98]. This collaboration between CTGF and TGF-β1 has contributed to the pro-fibrotic properties of TGF-β1 confirming the role of CTGF for TGF-β1 induction as a co-factor of gene expression.

The expression of CTGF was found increased in cultured fibroblasts from HTS, keloids and chronic fibrotic disorders [99]. In addition, cultured fibroblasts from HTS showed an increased expression of CTGF after stimulation by TGF-β [100]. In order to evaluate the role of CTGF in HTS formation, a rabbit animal model was established by Sisco at el. Antisense therapy was used to inhibit the expression of CTGF. Real-time reverse transcription polymerase chained reaction demonstrated an increased expression of CTGF in scar tissue and decreased CTGF expression after the intradermal injection of antisense oligonucleotides. The study showed that inhibition of CTGF in different times in wound healing has a substantial effect on reducing HTS [101]. Another experiment used CTGF small interfering RNA (siRNA) to successfully block the increase in CTGF mRNA levels and the result demonstrated that CTGF could regulate the gene expression of ECM, tissue inhibitor metalloproteinases and partial function of TGF-β1 [102]. In order to elucidate the pathophysiological function of CTGF, CTGF knock-out mice were used in the experiment and those mice died immediately after birth due to malformation of the rib cages. As well, the embryonic fibroblasts from this animal model showed an inability of adhesion and α-SMA formation. All these results suggest that CTGF functions in ECM adhesion and production [103, 104].

Taken together, CTGF acts as a downstream mediator of the TGF-β1 signaling pathway, directly involved in ECM synthesis and assists with TGF-β1 in the pathogenesis of HTS.

Platelet-derived growth factor (PDGF) is essential to wound healing and the over-expression of PDGF is important in the formation of HTS

PDGF has five isoforms, including PDGF-AA, PDGF-AB, PDGF-BB, PDGF-CC and PDGF-DD which function via the activation of three transmembrane receptor tyrosine kinases (RTKs) [105]. PDGF is produced by degranulated platelets in the early phase of the wound healing process and it is also secreted by macrophages during the proliferative phase of wound healing [106]. In wound healing-impaired mice, the expression of PDGF and their receptors decreased [107]. Moreover, PDGF showed reduced expression in chronic human non-healing ulcers compared to the fresh surgically created acute wounds [108]. All these studies support the important role of PDGF in wound healing. However, PDGF also has an important role in several fibrotic diseases including scleroderma, lung and liver fibrosis by promoting the growth and survival of myofibroblasts [109]. PDGF was found to mediate the deposition of collagen in fibroblasts and it was highly over-expressed in both the epidermis and the dermis of HTS. Over-production of collagen was not only related to high levels of TGF-β1, but also with increased expression of PDGF [110]. Another experiment showed that PDGF stimulated myofibroblast formation and increased TGF-β receptor I (TGF-βRI) and TGF-β receptor II (TGF-βRII) expression [111].

Although there are a lot of studies showing that PDGF plays a role in the pathogenesis of HTS, the exact molecular mechanism is still unknown.

Inhibitory effect of basic fibroblast growth factor (bFGF) on HTS via the regulation of collagen production, myofibroblast differentiation and TGF-β receptor expression

Fibroblast growth factors (FGFs) are a large family of growth factors that consist of 22 members with similar structural polypeptide. They have four receptors, which are transmembrane protein tyrosine kinases [112, 113]. Among the growth factors that play roles in wound healing, bFGF is particularly important [114]. bFGF is produced by keratinocytes and is found in the early stages of wound healing. It stimulates growth and differentiation of several types of cells, such as fibroblasts [115]. In a rat model, bFGF was detected in granulation tissue including regenerated epidermis and newborn capillaries [116]. As well, bFGF was found to promote wound healing by stimulating angiogenesis and granulation tissue proliferation [117]. However, bFGF might inhibit the granulation tissue formation by promoting apoptosis [118] and affect tumor growth [119].

Evidence for the importance of bFGF in the pathogenesis of HTS was provided by Tiede et al. that bFGF reduced α-SMA expression by inhibiting myofibroblast differentiation and it also decreased TGF-βRI and TGF-βRII expression [111]. In a rabbit HTS ear model, bFGF was applied everyday for three months and the wounds showed decreased collagen expression and increased MMP-1 expression such that bFGF appeared to have a negative effect on scar formation [120]. In humans bFGF was administered to acute incisional wounds after suturing and the patients remained free from HTS [121]. Hepatocyte growth factor (HGF) and MMP-1 have been demonstrated to have an anti-scarring effect [122]. In a more recent study, the expressions of HGF and MMP-1 were highly regulated in bFGF treated HTS and normal fibroblasts. The highly regulated MMP-1 expression might contribute to the increase of type I and type III collagen degradation, which leads to reduced scar formation. In vitro, bFGF treatment significantly decreased scar weight and the amount of collagen in nude mice that underwent human scar tissue transplantation [123]. Therefore, bFGF can inhibit HTS formation and the mechanism might be related to the regulation of collagen production, myofibroblast differentiation and inhibition of TGF-β receptor expression.

Macrophages involve in HTS formation via Stromal cell-derived factor 1 (SDF-1)/CXCR4 chemokine pathway

Significant more mast cells, fibrocytes and macrophages were found in nude mice that received human split thickness skin graft (STSG) compared to nude mice that received human full thickness skin graft (FTSG) in vivo, where HTS formation was found on both mice 2 months after the grafting with more scar observed in mice that received STSG, suggesting that inflammatory cells and bone marrow-derived fibrocytes might play critical roles in HTS formation in this human HTS-like nude mouse model [124]. A sequent study showed increased grafted skin thickness, increased number of myofibroblasts, decreased decorin and increased biglycan expression, positive staining of human leukocyte antigen in STSG grafted skin that formed persistent scars, which showed morphologic, histologic and immunohistochemical consistency with human HTS [125]. This animal model provides a means to study HTS and test new novel treatment options. Although there is not an ideal animal model that can be directly translated into human subjects to clearly explain the molecular basis of HTS formation, the human HTS-like nude mouse model is closer to the perfect animal model because the survived human skin grafts possess the genetic and histological properties of human HTS.

SDF-1 is found to be a potent chemokine that attracts lymphocytes and monocytes by binding exclusively to its receptor, CXCR4 [126128]. Studies focused on the functions of SDF-1/CXCR4 signaling have suggested that it involves not only in the tumor metastasis and vascularization but also in the pathogenesis of fibroproliferative diseases [129, 130]. Recent studies found up-regulated SDF-1 expression in the HTS tissue and serum of the burn patients as well as increased number of CD14+ CXCR4+ cells in the peripheral blood mononuclear cells, which suggested that SDF-1/CXCR4 signaling could recruit these CXCR4+ cells such as monocytes to the prolonged inflamed injured site and contribute to HTS formation [131]. In order to further verify the role of SDF-1/CXCR4 signaling in HTS formation, the CXCR4 antagonist CTCE-9908 was used to inhibit the SDF-1/CXCR4 effect on the human HTS-like nude mouse model. The study showed that CTCE-9908 significantly attenuated scar formation and contraction, reduced the number of macrophages in the tissue, which was differentiated and replenished by CXCR4 expressing monocytes in the circulation [132]. These findings support the role of SDF-1/CXCR4 in HTS formation and suggest an important role of macrophages in HTS formation.

Macrophages were first discovered by a Russian scientists, Élie Metchnikoff, in 1884 [133]. They are differentiated from newly recruited monocytes from the circulation. They are considered to play a vital role in the whole wound healing process because recent studies showed that impaired wound healing was associated with decreased number of macrophage infiltration at the injured site [134, 135]. However, pathological functioning of macrophages in the abnormal wound healing process can lead to disordered wound healing, including the formation of HTS [136]. Macrophages have two phenotypes, classically activated macrophages or the so called M1 macrophages and alternatively activated macrophages or the so called M2 macrophages [137]. Mahdavian et al. reported that M1 and M2 macrophages have distinct opposite functions in the wound healing process [136]. M1 macrophages can induce MMP-1 secretion and promote ECM degradation while M2 macrophages can secret large amount of TGF-β1, which can stimulate myofibroblast transformation and lead to ECM deposition. It is also hypothesized that prolong inflammatory phase will attract more macrophages and those macrophages will initially be more pro-inflammatory M1 phenotype and then switch to a more pro-fibrotic M2 phenotype due to more intense stimuli from the microenvironment [138]. The most distinct difference between M1 and M2 macrophages is that in M1 macrophages the arginine metabolism is shifted to nitric oxide and citrulline while in M2 macrophages it is shifted to ornithine and polyamines [139]. Growing evidence suggests that M2 macrophages are not constituted by a uniform population but can be further subdivided into M2a, M2b and M2c subsets [140]. M2a macrophages are induced by IL-4 and IL-13, which are involved in the anti-parasitic immune response and are considered to be pro-fibrotic. M2b macrophages are induced by IL-1β, LPS and immune complexes while M2c macrophages are induced by IL-10, TGF-β and glucocorticoids [141]. The fourth type, M2d macrophages, are characterized by switching from a M1 phenotype into an angiogenic M2-like phenotype, which termed M2d by Leibovich et al. [142].

Although studies suggest a close relationship between SDF-1/CXCR4 signaling and macrophage infiltration in the formation of HTS, more studies on the interaction between the two is still needed. Meanwhile, the roles of macrophage phenotypes in different phases of abnormal wound healing, like HTS-like nude mouse model, are to be investigated. Here we hypothesize that the monocytes, CXCR4 expressing cells in the circulation, will be attracted to the injured site via the SDF-1/CXCR4 signaling pathway due to concentration difference between the circulation and local tissue as well as the chemotactic effect of SDF-1. The monocytes then differentiate into M1 macrophages (NF-κB and STAT1 signaling pathways) and M2 macrophages (STAT3 and STAT6 signaling pathways) [143]. M1 macrophages secret pro-inflammatory cytokines such as IFN-γ, IL-1β, TNF-α, IL-6, IL-8 and generate reactive oxygen and nitric oxide through the activation of nitric oxide synthase 2 (NOS2). On the other hand, M2 macrophages inhibit the NOS2 activity via the activation of arginase-1. The distinct opposite and complementary functions of M1 and M2 macrophages will eventually lead to normal wound healing. However, in prolonged inflammatory environment such as wounds from a patient who suffered from severe thermal injury, large amounts of TGF-β1 can be produced together with increased myofibroblast proliferation, which will result in ECM deposition and finally HTS formation (Fig. 2).
Fig. 2

The roles of monocytes and polarized macrophages in HTS formation. We hypothesize that monocytes in the blood are recruited to the injured site via the SDF-1/CXCR4 signaling pathway and differentiate into polarized macrophages. The polarized M1 and M2 macrophages then exert their functions via various signaling pathways and involve in wound healing and HTS formation

Conclusions

In this review, four phases of normal wound healing are discussed before outlining the pathogenesis of HTS, illustrating the delicate balance of ECM deposition and degradation which influences the outcome of the wound healing process. Differentiating HTS from keloids is also important because the clinical and molecular mechanisms are different leading to distinct therapeutic outcomes. HTS formation is a dynamic, complex process that involves interactions between multiple factors such as inflammatory cells, cytokines, growth factors, and chemokines. Keratinocytes and mast cells are considered to be involved in HTS formation. The role of cytokines such as IL-1, TNF-α, IL-6 and IL-10 as well as growth factors such as TGF-β, CTGF, PDGF and bFGF in HTS formation were discussed. Despite the complexity of HTS, more attentions are drawn to the molecular and cellular mechanism of HTS for technological and scientific advances such as the establishment of new animal models and in vitro techniques. Growing studies are focusing on the roles of polarized macrophages in HTS formation and it is suggested that polarized macrophages actively participate in HTS formation via the SDF-1/CXCR4 signaling pathway. A preliminary experiment conducted by our laboratory confirmed potential roles of M2 macrophages in HTS formation. A subsequent study of specific depletion of M2 macrophages by Cre-LoxP technology on our human HTS-like nude mouse model together with the study of the roles of molecular precursors mentioned above might provide novel findings and potential new treatment and prevention of HTS.

Abbreviations

α-SMA: 

Alpha-smooth muscle actin

bFGF: 

Basic fibroblast growth factor

CTGF: 

Connective tissue growth factor

ECM: 

Extracellular matrix

FGFs: 

Fibroblast growth factors

FOXO1: 

Forkhead box protein O1

FTSG: 

Full thickness skin graft

GJIC: 

Gap junction intercellular communication

HGF: 

Hepatocyte growth factor

HTS: 

Hypertrophic scars

IFN-α2b: 

Interferon-alpha2b

IFN-γ: 

Interferon-gamma

IL-1: 

Interleukin-1

IL-1α: 

Interleukin-1 alpha

IL-1β: 

Interleukin-1 beta

IL-6: 

Interleukin-6

IL-10: 

Interleukin-10

LPS: 

Lipopolysaccharide

MCP-1: 

Monocyte chemotactic protein-1

MMPs: 

Matrix metalloproteinases

NOS2: 

Nitric oxide synthase 2

PDGF: 

Platelet-derived growth factor

PMNs: 

Polymorphonuclear neutrophils

RTKs: 

Receptor tyrosine kinases

SDF-1: 

Stromal cell-derived factor 1

siRNA: 

Small interfering RNA

STSG: 

Split thickness skin graft

TGF-β: 

Transforming growth factor-beta

TGF-β1: 

Transforming growth factor-beta 1

TGF-β2: 

Transforming growth factor-beta 2

TGF-β3: 

Transforming growth factor-beta 3

TGF-βRI: 

TGF-β receptor I

TGF-βRII: 

TGF-β receptor II

TNF-α: 

Tumor necrosis factor-alpha

VEGF: 

Vascular endothelial growth factor

Declarations

Acknowledgements

Authors gratefully acknowledge the support from Li Ka Shine Sino-Canadian Exchange Program (ZZ). This work was also funded by University Hospital foundation from University of Alberta and the Firefighters’ Burn Trust Fund.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

(1)
Wound Healing Research Group, Division of Plastic and Reconstructive Surgery, University of Alberta
(2)
Division of Plastic Surgery, Department of Surgery, University of Alberta
(3)
Department of Burn and Reconstructive Surgery, 2nd Affiliated Hospital of Shantou University Medical College

References

  1. Engrav LH, Garner WL, Tredget EE. Hypertrophic scar, wound contraction and hyper-hypopigmentation. J Burn Care Res official publ Am Burn Assoc. 2007;28:593–7.View ArticleGoogle Scholar
  2. Bombaro KM, Engrav LH, Carrougher GJ, Wiechman SA, Faucher L, Costa BA, et al. What is the prevalence of hypertrophic scarring following burns? Burns. 2003;29:299–302.PubMedView ArticleGoogle Scholar
  3. Reinke JM, Sorg H. Wound repair and regeneration. Eur surg res Eur chirurgische Forschung Recherches chirurgicales eu. 2012;49:35–43. doi:https://doi.org/10.1159/000339613.View ArticleGoogle Scholar
  4. Miller MC, Nanchahal J. Advances in the modulation of cutaneous wound healing and scarring. BioDrugs. 2005;19:363–81.PubMedView ArticleGoogle Scholar
  5. Werner S, Grose R. Regulation of wound healing by growth factors and cytokines. Physiol Rev. 2003;83:835–70. doi:https://doi.org/10.1152/physrev.00031.2002.PubMedGoogle Scholar
  6. Robson MC, Steed DL, Franz MG. Wound healing: biologic features and approaches to maximize healing trajectories. Curr Probl Surg. 2001;38:72–140. doi:https://doi.org/10.1067/msg.2001.111167.PubMedView ArticleGoogle Scholar
  7. Midwood KS, Williams LV, Schwarzbauer JE. Tissue repair and the dynamics of the extracellular matrix. Int J Biochem Cell Biol. 2004;36:1031–7. doi:https://doi.org/10.1016/j.biocel.2003.12.003.PubMedView ArticleGoogle Scholar
  8. Eming SA, Krieg T, Davidson JM. Inflammation in wound repair: molecular and cellular mechanisms. J Invest Dermatol. 2007;127:514–25. doi:https://doi.org/10.1038/sj.jid.5700701.PubMedView ArticleGoogle Scholar
  9. Bauer SM, Bauer RJ, Velazquez OC. Angiogenesis, vasculogenesis, and induction of healing in chronic wounds. Vasc Endovasc Surg. 2005;39:293–306.View ArticleGoogle Scholar
  10. Lau K, Paus R, Tiede S, Day P, Bayat A. Exploring the role of stem cells in cutaneous wound healing. Exp Dermatol. 2009;18:921–33. doi:https://doi.org/10.1111/j.1600-0625.2009.00942.x.PubMedView ArticleGoogle Scholar
  11. Barker TH. The role of ECM proteins and protein fragments in guiding cell behavior in regenerative medicine. Biomaterials. 2011;32:4211–4. doi:https://doi.org/10.1016/j.biomaterials.2011.02.027.PubMedView ArticleGoogle Scholar
  12. Profyris C, Tziotzios C, Do VI. Cutaneous scarring: pathophysiology, molecular mechanisms, and scar reduction therapeutics part I. The molecular basis of scar formation. J Am Acad Dermatol. 2012;66:1–10. doi:https://doi.org/10.1016/j.jaad.2011.05.055. quiz 1–2.PubMedView ArticleGoogle Scholar
  13. Greenhalgh DG. The role of apoptosis in wound healing. Int J Biochem Cell Biol. 1998;30:1019–30.PubMedView ArticleGoogle Scholar
  14. Ghahary A, Shen YJ, Nedelec B, Wang R, Scott PG, Tredget EE. Collagenase production is lower in post-burn hypertrophic scar fibroblasts than in normal fibroblasts and is reduced by insulin-like growth factor-1. J Invest Dermatol. 1996;106:476–81.PubMedView ArticleGoogle Scholar
  15. Brown JJ, Bayat A. Genetic susceptibility to raised dermal scarring. Br J Dermatol. 2009;161:8–18. doi:https://doi.org/10.1111/j.1365-2133.2009.09258.x.PubMedView ArticleGoogle Scholar
  16. Gauglitz GG, Korting HC, Pavicic T, Ruzicka T, Jeschke MG. Hypertrophic scarring and keloids: pathomechanisms and current and emerging treatment strategies. Mol Med. 2011;17:113–25. doi:https://doi.org/10.2119/molmed.2009.00153.PubMedPubMed CentralView ArticleGoogle Scholar
  17. Tredget EE, Nedelec B, Scott PG, Ghahary A. Hypertrophic scars, keloids, and contractures. The cellular and molecular basis for therapy. Surg Clin North Am. 1997;77:701–30.PubMedView ArticleGoogle Scholar
  18. Scott PG, Ghahary A, Tredget EE. Molecular and cellular aspects of fibrosis following thermal injury. Hand Clin. 2000;16:271–87.PubMedGoogle Scholar
  19. Nedelec B, Shankowsky H, Scott PG, Ghahary A, Tredget EE. Myofibroblasts and apoptosis in human hypertrophic scars: the effect of interferon-alpha2b. Surgery. 2001;130:798–808. doi:https://doi.org/10.1067/msy.2001.116453.PubMedView ArticleGoogle Scholar
  20. Smith CJ, Smith JC, Finn MC. The possible role of mast cells (allergy) in the production of keloid and hypertrophic scarring. J Burn Care Rehabil. 1987;8:126–31.PubMedView ArticleGoogle Scholar
  21. Huang C, Murphy GF, Akaishi S, Ogawa R. Keloids and hypertrophic scars: update and future directions. Plast Reconstr Surg Glob Open. 2013;1:e25. doi:https://doi.org/10.1097/GOX.0b013e31829c4597.PubMedPubMed CentralView ArticleGoogle Scholar
  22. Funayama E, Chodon T, Oyama A, Sugihara T. Keratinocytes promote proliferation and inhibit apoptosis of the underlying fibroblasts: an important role in the pathogenesis of keloid. J Invest Dermatol. 2003;121:1326–31. doi:https://doi.org/10.1111/j.1523-1747.2003.12572.x.PubMedView ArticleGoogle Scholar
  23. Garner WL. Epidermal regulation of dermal fibroblast activity. Plast Reconstr Surg. 1998;102:135–9.PubMedView ArticleGoogle Scholar
  24. Machesney M, Tidman N, Waseem A, Kirby L, Leigh I. Activated keratinocytes in the epidermis of hypertrophic scars. Am J Pathol. 1998;152:1133–41.PubMedPubMed CentralGoogle Scholar
  25. Varkey M, Ding J, Tredget EE. Fibrotic remodeling of tissue-engineered skin with deep dermal fibroblasts is reduced by keratinocytes. Tissue Eng Part A. 2014;20:716–27. doi:https://doi.org/10.1089/ten.TEA.2013.0434. Epub 2013 Nov 9.PubMedGoogle Scholar
  26. Moyer KE, Saggers GC, Ehrlich HP. Mast cells promote fibroblast populated collagen lattice contraction through gap junction intercellular communication. Wound Repair Regen. 2004;12:269–75.PubMedView ArticleGoogle Scholar
  27. Foley TT, Ehrlich HP. Through gap junction communications, co-cultured mast cells and fibroblasts generate fibroblast activities allied with hypertrophic scarring. Plast Reconstr Surg. 2013;131:1036–44. doi: 10.97/PRS.0b013e3182865c3f.PubMedView ArticleGoogle Scholar
  28. Gruber BL, Kew RR, Jelaska A, Marchese MJ, Garlick J, Ren S, et al. Human mast cells activate fibroblasts: tryptase is a fibrogenic factor stimulating collagen messenger ribonucleic acid synthesis and fibroblast chemotaxis. J Immunol. 1997;158:2310–7.PubMedGoogle Scholar
  29. Kofford MW, Schwartz LB, Schechter NM, Yager DR, Diegelmann RF, Graham MF. Cleavage of type I procollagen by human mast cell chymase initiates collagen fibril formation and generates a unique carboxyl-terminal propeptide. J Biol Chem. 1997;272:7127–31.PubMedView ArticleGoogle Scholar
  30. Kupietzky A, Levi-Schaffer F. The role of mast cell-derived histamine in the closure of an in vitro wound. Inflammation research. 1996;45:176–80.PubMedView ArticleGoogle Scholar
  31. Chen L, Liu S, Li SR, Cong L, Wu JL, Wang ZX. [Influence of substance P on the release of histamine in the human hypertrophic scar tissue]. Zhonghua shao shang za zhi = Zhonghua shaoshang zazhi =. Chinese J of burns. 2006;22:192–4.Google Scholar
  32. Tredget EE, Iwashina T, Scott PG, Ghahary A. Determination of plasma Ntau-methylhistamine in vivo by isotope dilution using benchtop gas chromatography–mass spectrometry. J Chromatogr B Biomed Sci Appl. 1997;694:1–9.PubMedView ArticleGoogle Scholar
  33. Eishi K, Bae SJ, Ogawa F, Hamasaki Y, Shimizu K, Katayama I. Silicone gel sheets relieve pain and pruritus with clinical improvement of keloid: possible target of mast cells. J Dermatolog Treat. 2003;14:248–52.PubMedView ArticleGoogle Scholar
  34. Arno AI, Gauglitz GG, Barret JP, Jeschke MG. Up-to-date approach to manage keloids and hypertrophic scars: a useful guide. Burns. 2014;40:1255–66. doi:https://doi.org/10.1016/j.burns.2014.02.011.PubMedPubMed CentralView ArticleGoogle Scholar
  35. Tritto M, Kanat IO. Management of keloids and hypertrophic scars. J Am Podiatr Med Assoc. 1991;81:601–5. doi:https://doi.org/10.7547/87507315-81-11-601.PubMedView ArticleGoogle Scholar
  36. Slemp AE, Kirschner RE. Keloids and scars: a review of keloids and scars, their pathogenesis, risk factors, and management. Curr Opin Pediatr. 2006;18:396–402.PubMedView ArticleGoogle Scholar
  37. Murray JC. Keloids and hypertrophic scars. Clin Dermatol. 1994;12:27–37.PubMedView ArticleGoogle Scholar
  38. Halim AS, Emami A, Salahshourifar I, Kannan TP. Keloid scarring: understanding the genetic basis, advances, and prospects. Arch Plast Surg. 2012;39:184–9. doi:https://doi.org/10.5999/aps.2012.39.3.184.EpubMay10.PubMedPubMed CentralView ArticleGoogle Scholar
  39. Ehrlich HP, Desmouliere A, Diegelmann RF, Cohen IK, Compton CC, Garner WL, et al. Morphological and immunochemical differences between keloid and hypertrophic scar. Am J Pathol. 1994;145:105–13.PubMedPubMed CentralGoogle Scholar
  40. Ueda K, Furuya E, Yasuda Y, Oba S, Tajima S. Keloids have continuous high metabolic activity. Plast Reconstr Surg. 1999;104:694–8.PubMedView ArticleGoogle Scholar
  41. Tanaka A, Hatoko M, Tada H, Iioka H, Niitsuma K, Miyagawa S. Expression of p53 family in scars. J Dermatol Sci. 2004;34:17–24.PubMedView ArticleGoogle Scholar
  42. Younai S, Venters G, Vu S, Nichter L, Nimni ME, Tuan TL. Role of growth factors in scar contraction: an in vitro analysis. Ann Plast Surg. 1996;36:495–501.PubMedView ArticleGoogle Scholar
  43. Ledon JA, Savas J, Franca K, Chacon A, Nouri K. Intralesional treatment for keloids and hypertrophic scars: a review. Dermatol Surg. 2013;39:1745–57. doi:https://doi.org/10.1111/dsu.12346.PubMedView ArticleGoogle Scholar
  44. Schneider JC, Harris NL, El Shami A, Sheridan RL, Schulz 3rd JT, Bilodeau ML, et al. A descriptive review of neuropathic-like pain after burn injury. J Burn Care Res. 2006;27:524–8. doi:https://doi.org/10.1097/01.BCR.0000226019.76946.5D.PubMedView ArticleGoogle Scholar
  45. Wilgus TA, Wulff BC. The importance of mast cells in dermal scarring. Adv Wound Care. 2014;3:356–65. doi:https://doi.org/10.1089/wound.2013.0457.View ArticleGoogle Scholar
  46. Holavanahalli RK, Helm PA, Kowalske KJ. Long-Term Outcomes in Patients Surviving Large Burns: The Musculoskeletal System. Journal of burn care & research : official publication of the American Burn Association. 2015. doi:https://doi.org/10.1097/BCR.0000000000000257
  47. Stoddard Jr FJ, Ryan CM, Schneider JC. Physical and psychiatric recovery from burns. Surg Clin North Am. 2014;94:863–78. doi:https://doi.org/10.1016/j.suc.2014.05.007.PubMedView ArticleGoogle Scholar
  48. Robert R, Meyer W, Bishop S, Rosenberg L, Murphy L, Blakeney P. Disfiguring burn scars and adolescent self-esteem. Burns. 1999;25:581–5.PubMedView ArticleGoogle Scholar
  49. Zhu Z, Ding J, Shankowsky HA, Tredget EE. The molecular mechanism of hypertrophic scar. J cell commun signal. 2013;7:239–52. doi:https://doi.org/10.1007/s12079-013-0195-5.PubMedPubMed CentralView ArticleGoogle Scholar
  50. Mustoe TA, Cooter RD, Gold MH, Hobbs FD, Ramelet AA, Shakespeare PG, et al. International clinical recommendations on scar management. Plast Reconstr Surg. 2002;110:560–71.PubMedView ArticleGoogle Scholar
  51. Kwan P, Hori K, Ding J, Tredget EE. Scar and contracture: biological principles. Hand Clin. 2009;25:511–28. doi:https://doi.org/10.1016/j.hcl.2009.06.007.PubMedView ArticleGoogle Scholar
  52. Gold MH, Berman B, Clementoni MT, Gauglitz GG, Nahai F, Murcia C. Updated international clinical recommendations on scar management: part 1--evaluating the evidence. Dermatol Surg. 2014;40:817–24. doi:https://doi.org/10.1111/dsu.0000000000000049.PubMedGoogle Scholar
  53. Gold MH, McGuire M, Mustoe TA, Pusic A, Sachdev M, Waibel J, et al. Updated international clinical recommendations on scar management: part 2--algorithms for scar prevention and treatment. Dermatol Surg. 2014;40:825–31. doi:https://doi.org/10.1111/dsu.0000000000000050.PubMedGoogle Scholar
  54. Lee KK, Mehrany K, Swanson NA. Surgical revision. Dermatol Clin. 2005;23:141–50. doi:https://doi.org/10.1016/j.det.2004.08.006. vii.PubMedView ArticleGoogle Scholar
  55. Manuskiatti W, Fitzpatrick RE. Treatment response of keloidal and hypertrophic sternotomy scars: comparison among intralesional corticosteroid, 5-fluorouracil, and 585-nm flashlamp-pumped pulsed-dye laser treatments. Arch Dermatol. 2002;138:1149–55.PubMedView ArticleGoogle Scholar
  56. Aggarwal H, Saxena A, Lubana PS, Mathur RK, Jain DK. Treatment of keloids and hypertrophic scars using bleom. J Cosmet Dermatol. 2008;7:43–9. doi:https://doi.org/10.1111/j.473-2165.008.00360.x.PubMedView ArticleGoogle Scholar
  57. Draelos ZD. The ability of onion extract gel to improve the cosmetic appearance of postsurgical scars. J Cosmet Dermatol. 2008;7:101–4. doi:https://doi.org/10.1111/j.1473-2165.2008.00371.x.PubMedView ArticleGoogle Scholar
  58. Chanprapaph K, Tanrattanakorn S, Wattanakrai P, Wongkitisophon P, Vachiramon V. Effectiveness of onion extract gel on surgical scars in asians. Dermatol Res Pract. 2012;2012:212945.:10.1155/2012/212945. Epub 2012 Aug 8Google Scholar
  59. Gassner HG, Brissett AE, Otley CC, Boahene DK, Boggust AJ, Weaver AL, et al. Botulinum toxin to improve facial wound healing: a prospective, blinded, placebo-controlled study. Mayo Clin Proc. 2006;81:1023–8. doi:https://doi.org/10.4065/81.8.1023.PubMedView ArticleGoogle Scholar
  60. Heckmann M, Adelmann-Grill BC, Hein R, Krieg T. Biphasic effects of interleukin-1 alpha on dermal fibroblasts: enhancement of chemotactic responsiveness at low concentrations and of mRNA expression for collagenase at high concentrations. J Invest Dermatol. 1993;100:780–4.PubMedView ArticleGoogle Scholar
  61. Elias JA, Gustilo K, Baeder W, Freundlich B. Synergistic stimulation of fibroblast prostaglandin production by recombinant interleukin 1 and tumor necrosis factor. J Immunol. 1987;138:3812–6.PubMedGoogle Scholar
  62. Niessen FB, Andriessen MP, Schalkwijk J, Visser L, Timens W. Keratinocyte-derived growth factors play a role in the formation of hypertrophic scars. J Pathol. 2001;194:207–16. doi:https://doi.org/10.1002/path.853.PubMedView ArticleGoogle Scholar
  63. Salgado RM, Alcantara L, Mendoza-Rodriguez CA, Cerbon M, Hidalgo-Gonzalez C, Mercadillo P, et al. Post-burn hypertrophic scars are characterized by high levels of IL-1beta mRNA and protein and TNF-alpha type I receptors. Burns. 2012;38:668–76. doi:https://doi.org/10.1016/j.burns.2011.12.012.PubMedView ArticleGoogle Scholar
  64. Furuzawa-Carballeda J, Krotzsch E, Barile-Fabris L, Alcala M, Espinosa-Morales R. Subcutaneous administration of collagen-polyvinylpyrrolidone down regulates IL-1beta, TNF-alpha, TGF-beta1, ELAM-1 and VCAM-1 expression in scleroderma skin lesions. Clin Exp Dermatol. 2005;30:83–6. doi:https://doi.org/10.1111/j.1365-2230.2004.01691.x.PubMedView ArticleGoogle Scholar
  65. Castagnoli C, Stella M, Berthod C, Magliacani G, Richiardi PM. TNF production and hypertrophic scarring. Cell Immunol. 1993;147:51–63. doi:https://doi.org/10.1006/cimm.1993.1047.PubMedView ArticleGoogle Scholar
  66. Goldberg MT, Han YP, Yan C, Shaw MC, Garner WL. TNF-alpha suppresses alpha-smooth muscle actin expression in human dermal fibroblasts: an implication for abnormal wound healing. J Invest Dermatol. 2007;127:2645–55. doi:https://doi.org/10.1038/sj.jid.5700890.PubMedPubMed CentralView ArticleGoogle Scholar
  67. Witte MB, Barbul A. General principles of wound healing. Surg Clin North Am. 1997;77:509–28.PubMedView ArticleGoogle Scholar
  68. O’Donoghue RJ, Knight DA, Richards CD, Prele CM, Lau HL, Jarnicki AG, et al. Genetic partitioning of interleukin-6 signalling in mice dissociates Stat3 from Smad3-mediated lung fibrosis. EMBO Mol Med. 2012;4:939–51. doi:https://doi.org/10.1002/emmm.201100604.PubMedPubMed CentralView ArticleGoogle Scholar
  69. O’Reilly S, Ciechomska M, Cant R, Hugle T, van Laar JM. Interleukin-6, its role in fibrosing conditions. Cytokine Growth Factor Rev. 2012;23:99–107. doi:https://doi.org/10.1016/j.cytogfr.2012.04.003.PubMedView ArticleGoogle Scholar
  70. Xue H, McCauley RL, Zhang W, Martini DK. Altered interleukin-6 expression in fibroblasts from hypertrophic burn scars. J Burn Care Rehabil. 2000;21:142–6.PubMedView ArticleGoogle Scholar
  71. Dasu MR, Hawkins HK, Barrow RE, Xue H, Herndon DN. Gene expression profiles from hypertrophic scar fibroblasts before and after IL-6 stimulation. J Pathol. 2004;202:476–85.PubMedView ArticleGoogle Scholar
  72. Liechty KW, Adzick NS, Crombleholme TM. Diminished interleukin 6 (IL-6) production during scarless human fetal wound repair. Cytokine. 2000;12:671–6. doi:https://doi.org/10.1006/cyto.1999.0598.PubMedView ArticleGoogle Scholar
  73. McFarland-Mancini MM, Funk HM, Paluch AM, Zhou M, Giridhar PV, Mercer CA, et al. Differences in wound healing in mice with deficiency of IL-6 versus IL-6 receptor. J Immunol. 2010;184:7219–28. doi:https://doi.org/10.4049/jimmunol.0901929.PubMedView ArticleGoogle Scholar
  74. Moore KW, de Waal MR, Coffman RL, O’Garra A. Interleukin-10 and the interleukin-10 receptor. Annu Rev Immunol. 2001;19:683–765. doi:https://doi.org/10.1146/annurev.immunol.19.1.683.PubMedView ArticleGoogle Scholar
  75. Sato Y, Ohshima T, Kondo T. Regulatory role of endogenous interleukin-10 in cutaneous inflammatory response of murine wound healing. Biochem Biophys Res Commun. 1999;265:194–9.PubMedView ArticleGoogle Scholar
  76. Fortunato SJ, Menon R, Swan KF, Lombardi SJ. Interleukin-10 inhibition of interleukin-6 in human amniochorionic membrane: transcriptional regulation. Am J Obstet Gynecol. 1996;175:1057–65.PubMedView ArticleGoogle Scholar
  77. Dagvadorj J, Naiki Y, Tumurkhuu G, Noman AS, Iftekar EKI, Koide N, et al. Interleukin (IL)-10 attenuates lipopolysaccharide-induced IL-6 production via inhibition of IkappaB-zeta activity by Bcl-3. Innate Immun. 2009;15:217–24. doi:https://doi.org/10.1177/1753425909103738.PubMedView ArticleGoogle Scholar
  78. Henderson J, Ferguson MW, Terenghi G. The reinnervation and revascularization of wounds is temporarily altered after treatment with interleukin 10. Wound Repair Regen. 2011;19:268–73. doi:https://doi.org/10.1111/j.524-475X.2011.00667.x.PubMedView ArticleGoogle Scholar
  79. Liechty KW, Kim HB, Adzick NS, Crombleholme TM. Fetal wound repair results in scar formation in interleukin-10-deficient mice in a syngeneic murine model of scarless fetal wound repair. J Pediatr Surg. 2000;35:866–72. doi:https://doi.org/10.1053/jpsu.2000.6868. discussion 72–3.PubMedView ArticleGoogle Scholar
  80. Gordon A, Kozin ED, Keswani SG, Vaikunth SS, Katz AB, Zoltick PW, et al. Permissive environment in postnatal wounds induced by adenoviral-mediated overexpression of the anti-inflammatory cytokine interleukin-10 prevents scar formation. Wound Repair Regen. 2008;16:70–9. doi:https://doi.org/10.1111/j.1524-475X.2007.00326.x.PubMedView ArticleGoogle Scholar
  81. Peranteau WH, Zhang L, Muvarak N, Badillo AT, Radu A, Zoltick PW, et al. IL-10 overexpression decreases inflammatory mediators and promotes regenerative healing in an adult model of scar formation. J Invest Dermatol. 2008;128:1852–60. doi:https://doi.org/10.1038/sj.jid.5701232.PubMedView ArticleGoogle Scholar
  82. Alexandrow MG, Moses HL. Transforming growth factor beta and cell cycle regulation. Cancer Res. 1995;55:1452–7.PubMedGoogle Scholar
  83. McGee GS, Broadley KN, Buckley A, Aquino A, Woodward SC, Demetriou AA, et al. Recombinant transforming growth factor beta accelerates incisional wound healing. Curr Surg. 1989;46:103–6.PubMedGoogle Scholar
  84. Roberts AB, Sporn MB, Assoian RK, Smith JM, Roche NS, Wakefield LM, et al. Transforming growth factor type beta: rapid induction of fibrosis and angiogenesis in vivo and stimulation of collagen formation in vitro. Proc Natl Acad Sci U S A. 1986;83:4167–71.PubMedPubMed CentralView ArticleGoogle Scholar
  85. Broekelmann TJ, Limper AH, Colby TV, McDonald JA. Transforming growth factor beta 1 is present at sites of extracellular matrix gene expression in human pulmonary fibrosis. Proc Natl Acad Sci U S A. 1991;88:6642–6.PubMedPubMed CentralView ArticleGoogle Scholar
  86. Ghahary A, Shen YJ, Scott PG, Tredget EE. Immunolocalization of TGF-beta 1 in human hypertrophic scar and normal dermal tissues. Cytokine. 1995;7:184–90.PubMedView ArticleGoogle Scholar
  87. Bock O, Yu H, Zitron S, Bayat A, Ferguson MW, Mrowietz U. Studies of transforming growth factors beta 1–3 and their receptors I and II in fibroblast of keloids and hypertrophic scars. Acta Derm Venereol. 2005;85:216–20.PubMedGoogle Scholar
  88. Shah M, Foreman DM, Ferguson MW. Neutralising antibody to TGF-beta 1,2 reduces cutaneous scarring in adult rodents. J Cell Sci. 1994;107(Pt 5):1137–57.PubMedGoogle Scholar
  89. Shah M, Foreman DM, Ferguson MW. Neutralisation of TGF-beta 1 and TGF-beta 2 or exogenous addition of TGF-beta 3 to cutaneous rat wounds reduces scarring. J Cell Sci. 1995;108(Pt 3):985–1002.PubMedGoogle Scholar
  90. Lu L, Saulis AS, Liu WR, Roy NK, Chao JD, Ledbetter S, et al. The temporal effects of anti-TGF-beta1, 2, and 3 monoclonal antibody on wound healing and hypertrophic scar formation. J Am Coll Surg. 2005;201:391–7.PubMedView ArticleGoogle Scholar
  91. Hameedaldeen A, Liu J, Batres A, Graves GS, Graves DT. FOXO1, TGF-beta regulation and wound healing. Int J Mol Sci. 2014;15:16257–69. doi:https://doi.org/10.3390/ijms150916257.PubMedPubMed CentralView ArticleGoogle Scholar
  92. Tredget EE, Shankowsky HA, Pannu R, Nedelec B, Iwashina T, Ghahary A, et al. Transforming growth factor-beta in thermally injured patients with hypertrophic scars: effects of interferon alpha-2b. Plast Reconstr Surg. 1998;102:1317–28. discussion 29–30.PubMedView ArticleGoogle Scholar
  93. Tredget EE, Wang R, Shen Q, Scott PG, Ghahary A. Transforming growth factor-beta mRNA and protein in hypertrophic scar tissues and fibroblasts: antagonism by IFN-alpha and IFN-gamma in vitro and in vivo. J Interf Cytokine Res. 2000;20:143–51. doi:https://doi.org/10.1089/107999000312540.View ArticleGoogle Scholar
  94. Tredget EB, Demare J, Chandran G, Tredget EE, Yang L, Ghahary A. Transforming growth factor-beta and its effect on reepithelialization of partial-thickness ear wounds in transgenic mice. Wound Repair Regen. 2005;13:61–7. doi:https://doi.org/10.1111/j.1067-1927.2005.130108.x.PubMedView ArticleGoogle Scholar
  95. Honardoust D, Varkey M, Marcoux Y, Shankowsky HA, Tredget EE. Reduced decorin, fibromodulin, and transforming growth factor-beta3 in deep dermis leads to hypertrophic scarring. J Burn Care Res. 2012;33:218–27. doi:https://doi.org/10.1097/BCR.0b013e3182335980.PubMedView ArticleGoogle Scholar
  96. Mori T, Kawara S, Shinozaki M, Hayashi N, Kakinuma T, Igarashi A, et al. Role and interaction of connective tissue growth factor with transforming growth factor-beta in persistent fibrosis: a mouse fibrosis model. J Cell Physiol. 1999;181:153–9. doi:https://doi.org/10.1002/(SICI)1097-4652(199910)181%3A1%3C153%3A%3AAID-JCP16%3E3.0.CO%3B2-K.PubMedView ArticleGoogle Scholar
  97. Frazier K, Williams S, Kothapalli D, Klapper H, Grotendorst GR. Stimulation of fibroblast cell growth, matrix production, and granulation tissue formation by connective tissue growth factor. J Invest Dermatol. 1996;107:404–11.PubMedView ArticleGoogle Scholar
  98. Shi-wen X, Stanton LA, Kennedy L, Pala D, Chen Y, Howat SL, et al. CCN2 is necessary for adhesive responses to transforming growth factor-beta1 in embryonic fibroblasts. J Biol Chem. 2006;281:10715–26. Epub 2006 Feb 16.PubMedView ArticleGoogle Scholar
  99. Igarashi A, Nashiro K, Kikuchi K, Sato S, Ihn H, Fujimoto M, et al. Connective tissue growth factor gene expression in tissue sections from localized scleroderma, keloid, and other fibrotic skin disorders. J Invest Dermatol. 1996;106:729–33.PubMedView ArticleGoogle Scholar
  100. Colwell AS, Phan TT, Kong W, Longaker MT, Lorenz PH. Hypertrophic scar fibroblasts have increased connective tissue growth factor expression after transforming growth factor-beta stimulation. Plast Reconstr Surg. 2005;116:1387–90. discussion 91–2.PubMedView ArticleGoogle Scholar
  101. Sisco M, Kryger ZB, O’Shaughnessy KD, Kim PS, Schultz GS, Ding XZ, et al. Antisense inhibition of connective tissue growth factor (CTGF/CCN2) mRNA limits hypertrophic scarring without affecting wound healing in vivo. Wound Repair Regen. 2008;16:661–73. doi:https://doi.org/10.1111/j.524-475X.2008.00416.x.PubMedView ArticleGoogle Scholar
  102. Wang JF, Olson ME, Ma L, Brigstock DR, Hart DA. Connective tissue growth factor siRNA modulates mRNA levels for a subset of molecules in normal and TGF-beta 1-stimulated porcine skin fibroblasts. Wound Repair Regen. 2004;12:205–16. doi:https://doi.org/10.1111/j.1067-1927.2004.012113.x.PubMedView ArticleGoogle Scholar
  103. Ivkovic S, Yoon BS, Popoff SN, Safadi FF, Libuda DE, Stephenson RC, et al. Connective tissue growth factor coordinates chondrogenesis and angiogenesis during skeletal development. Development. 2003;130:2779–91.PubMedPubMed CentralView ArticleGoogle Scholar
  104. Chen Y, Abraham DJ, Shi-Wen X, Pearson JD, Black CM, Lyons KM, et al. CCN2 (connective tissue growth factor) promotes fibroblast adhesion to fibronectin. Mol Biol Cell. 2004;15:5635–46. doi:https://doi.org/10.1091/mbc.E04-06-0490.PubMedPubMed CentralView ArticleGoogle Scholar
  105. Heldin CH, Eriksson U, Ostman A. New members of the platelet-derived growth factor family of mitogens. Arch Biochem Biophys. 2002;398:284–90. doi:https://doi.org/10.1006/abbi.2001.2707.PubMedView ArticleGoogle Scholar
  106. Mori R, Shaw TJ, Martin P. Molecular mechanisms linking wound inflammation and fibrosis: knockdown of osteopontin leads to rapid repair and reduced scarring. J Exp Med. 2008;205:43–51. doi:https://doi.org/10.1084/jem.20071412.PubMedPubMed CentralView ArticleGoogle Scholar
  107. Beer HD, Longaker MT, Werner S. Reduced expression of PDGF and PDGF receptors during impaired wound healing. J Invest Dermatol. 1997;109:132–8.PubMedView ArticleGoogle Scholar
  108. Pierce GF, Tarpley JE, Tseng J, Bready J, Chang D, Kenney WC, et al. Detection of platelet-derived growth factor (PDGF)-AA in actively healing human wounds treated with recombinant PDGF-BB and absence of PDGF in chronic nonhealing wounds. J Clin Invest. 1995;96:1336–50. doi:https://doi.org/10.1172/JCI118169.PubMedPubMed CentralView ArticleGoogle Scholar
  109. Bonner JC. Regulation of PDGF and its receptors in fibrotic diseases. Cytokine Growth Factor Rev. 2004;15:255–73. doi:https://doi.org/10.1016/j.cytogfr.2004.03.006.PubMedView ArticleGoogle Scholar
  110. Tan EM, Qin H, Kennedy SH, Rouda S, Fox JW, Moore Jr JH. Platelet-derived growth factors-AA and -BB regulate collagen and collagenase gene expression differentially in human fibroblasts. Biochem J. 1995;310(Pt 2):585–8.PubMedPubMed CentralView ArticleGoogle Scholar
  111. Tiede S, Ernst N, Bayat A, Paus R, Tronnier V, Zechel C. Basic fibroblast growth factor: a potential new therapeutic tool for the treatment of hypertrophic and keloid scars. Annals of anatomy. Anat Anz. 2009;191:33–44. doi:https://doi.org/10.1016/j.aanat.2008.10.001.View ArticleGoogle Scholar
  112. Ornitz DM, Itoh N. Fibroblast growth factors. Genome biology. 2001;2:REVIEWS3005.Google Scholar
  113. Johnson DE, Williams LT. Structural and functional diversity in the FGF receptor multigene family. Adv Cancer Res. 1993;60:1–41.PubMedView ArticleGoogle Scholar
  114. Nissen NN, Polverini PJ, Gamelli RL, DiPietro LA. Basic fibroblast growth factor mediates angiogenic activity in early surgical wounds. Surgery. 1996;119:457–65.PubMedView ArticleGoogle Scholar
  115. Chen GJ, Forough R. Fibroblast growth factors, fibroblast growth factor receptors, diseases, and drugs. Recent Pat Cardiovasc Drug Discov. 2006;1:211–24.PubMedView ArticleGoogle Scholar
  116. Kibe Y, Takenaka H, Kishimoto S. Spatial and temporal expression of basic fibroblast growth factor protein during wound healing of rat skin. Br J Dermatol. 2000;143:720–7.PubMedView ArticleGoogle Scholar
  117. Klingbeil CK, Cesar LB, Fiddes JC. Basic fibroblast growth factor accelerates tissue repair in models of impaired wound healing. Prog Clin Biol Res. 1991;365:443–58.PubMedGoogle Scholar
  118. Akasaka Y, Ono I, Yamashita T, Jimbow K, Ishii T. Basic fibroblast growth factor promotes apoptosis and suppresses granulation tissue formation in acute incisional wounds. J Pathol. 2004;203:710–20. doi:https://doi.org/10.1002/path.1574.PubMedView ArticleGoogle Scholar
  119. Agasse F, Nicoleau C, Petit J, Jaber M, Roger M, Benzakour O, et al. Evidence for a major role of endogenous fibroblast growth factor-2 in apoptotic cortex-induced subventricular zone cell proliferation. Eur J Neurosci. 2007;26:3036–42. doi:https://doi.org/10.1111/j.1460-9568.2007.05915.x.PubMedView ArticleGoogle Scholar
  120. Xie JL, Bian HN, Qi SH, Chen HD, Li HD, Xu YB, et al. Basic fibroblast growth factor (bFGF) alleviates the scar of the rabbit ear model in wound healing. Wound Repair Regen. 2008;16:576–81. doi:https://doi.org/10.1111/j.1524-475X.2008.00405.x.PubMedView ArticleGoogle Scholar
  121. Ono I, Akasaka Y, Kikuchi R, Sakemoto A, Kamiya T, Yamashita T, et al. Basic fibroblast growth factor reduces scar formation in acute incisional wounds. Wound Repair Regen. 2007;15:617–23. doi:https://doi.org/10.1111/j.1524-475X.2007.00293.x.PubMedView ArticleGoogle Scholar
  122. Jinnin M, Ihn H, Mimura Y, Asano Y, Yamane K, Tamaki K. Effects of hepatocyte growth factor on the expression of type I collagen and matrix metalloproteinase-1 in normal and scleroderma dermal fibroblasts. J Invest Dermatol. 2005;124:324–30. doi:https://doi.org/10.1111/j.0022-202X.2004.23601.x.PubMedView ArticleGoogle Scholar
  123. Eto H, Suga H, Aoi N, Kato H, Doi K, Kuno S, et al. Therapeutic potential of fibroblast growth factor-2 for hypertrophic scars: upregulation of MMP-1 and HGF expression. Lab Invest. 2012;92:214–23. doi:https://doi.org/10.1038/labinvest.2011.127.PubMedView ArticleGoogle Scholar
  124. Wang J, Ding J, Jiao H, Honardoust D, Momtazi M, Shankowsky HA, et al. Human hypertrophic scar-like nude mouse model: characterization of the molecular and cellular biology of the scar process. Wound repair and regeneration : official publication of the wound healing society [and] the European tissue repair. Society. 2011;19:274–85. doi:https://doi.org/10.1111/j.1524-475X.2011.00672.x.Google Scholar
  125. Momtazi M, Kwan P, Ding J, Anderson CC, Honardoust D, Goekjian S, et al. A nude mouse model of hypertrophic scar shows morphologic and histologic characteristics of human hypertrophic scar. Wound Repair Regen. 2013;21:77–87. doi:https://doi.org/10.1111/j.524-475X.2012.00856.x. Epub 2012 Nov 5.PubMedView ArticleGoogle Scholar
  126. Bleul CC, Fuhlbrigge RC, Casasnovas JM, Aiuti A, Springer TA. A highly efficacious lymphocyte chemoattractant, stromal cell-derived factor 1 (SDF-1). J Exp Med. 1996;184:1101–9.PubMedView ArticleGoogle Scholar
  127. Hamada T, Mohle R, Hesselgesser J, Hoxie J, Nachman RL, Moore MA, et al. Transendothelial migration of megakaryocytes in response to stromal cell-derived factor 1 (SDF-1) enhances platelet formation. J Exp Med. 1998;188:539–48.PubMedPubMed CentralView ArticleGoogle Scholar
  128. Choi WT, An J. Biology and clinical relevance of chemokines and chemokine receptors CXCR4 and CCR5 in human diseases. Exp Biol Med. 2011;236:637–47. doi:https://doi.org/10.1258/ebm.2011.010389.View ArticleGoogle Scholar
  129. Balkwill F. Cancer and the chemokine network. Nat Rev Cancer. 2004;4:540–50. doi:https://doi.org/10.1038/nrc1388.PubMedView ArticleGoogle Scholar
  130. Xu J, Mora A, Shim H, Stecenko A, Brigham KL, Rojas M. Role of the SDF-1/CXCR4 axis in the pathogenesis of lung injury and fibrosis. Am J Respir Cell Mol Biol. 2007;37:291–9. Epub 2007 Apr 26.PubMedPubMed CentralView ArticleGoogle Scholar
  131. Ding J, Hori K, Zhang R, Marcoux Y, Honardoust D, Shankowsky HA, et al. Stromal cell-derived factor 1 (SDF-1) and its receptor CXCR4 in the formation of postburn hypertrophic scar (HTS). Wound Repair Regen. 2011;19:568–78. doi:https://doi.org/10.1111/j.524-475X.2011.00724.x.PubMedView ArticleGoogle Scholar
  132. Ding J, Ma Z, Liu H, Kwan P, Iwashina T, Shankowsky HA, et al. The therapeutic potential of a C-X-C chemokine receptor type 4 (CXCR-4) antagonist on hypertrophic scarring in vivo. Wound Repair Regen. 2014;22:622–30. doi:https://doi.org/10.1111/wrr.12208.PubMedView ArticleGoogle Scholar
  133. Zalkind SIA. Ilya Mechnikov, his life and work. Moscow: Foreign Languages Pub. House; 1959.Google Scholar
  134. Nagaoka T, Kaburagi Y, Hamaguchi Y, Hasegawa M, Takehara K, Steeber DA, et al. Delayed wound healing in the absence of intercellular adhesion molecule-1 or L-selectin expression. Am J Pathol. 2000;157:237–47. doi:https://doi.org/10.1016/S0002-9440(10)64534-8.PubMedPubMed CentralView ArticleGoogle Scholar
  135. Eming SA, Werner S, Bugnon P, Wickenhauser C, Siewe L, Utermohlen O, et al. Accelerated wound closure in mice deficient for interleukin-10. Am J Pathol. 2007;170:188–202. doi:https://doi.org/10.2353/ajpath.2007.060370.PubMedPubMed CentralView ArticleGoogle Scholar
  136. Mahdavian Delavary B, van der Veer WM, van Egmond M, Niessen FB, Beelen RH. Macrophages in skin injury and repair. Immunobiology. 2011;216:753–62. doi:https://doi.org/10.1016/j.imbio.2011.01.001.PubMedView ArticleGoogle Scholar
  137. Wynn TA. Fibrotic disease and the T(H)1/T(H)2 paradigm. Nat Rev Immunol. 2004;4:583–94. doi:https://doi.org/10.1038/nri1412.PubMedPubMed CentralView ArticleGoogle Scholar
  138. Song E, Ouyang N, Horbelt M, Antus B, Wang M, Exton MS. Influence of alternatively and classically activated macrophages on fibrogenic activities of human fibroblasts. Cell Immunol. 2000;204:19–28. doi:https://doi.org/10.1006/cimm.2000.1687.PubMedView ArticleGoogle Scholar
  139. Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM. M-1/M-2 macrophages and the Th1/Th2 paradigm. J Immunol. 2000;164:6166–73.PubMedView ArticleGoogle Scholar
  140. Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M. The chemokine system in diverse forms of macrophage activation and polarization. Trends Immunol. 2004;25:677–86. doi:https://doi.org/10.1016/j.it.2004.09.015.PubMedView ArticleGoogle Scholar
  141. Martinez FO, Sica A, Mantovani A, Locati M. Macrophage activation and polarization. Front Biol. 2008;13:453–61.View ArticleGoogle Scholar
  142. Grinberg S, Hasko G, Wu D, Leibovich SJ. Suppression of PLCbeta2 by endotoxin plays a role in the adenosine A(2A) receptor-mediated switch of macrophages from an inflammatory to an angiogenic phenotype. Am J Pathol. 2009;175:2439–53. doi: 10.353/ajpath.009.090290. Epub 2009 Oct 22.PubMedPubMed CentralView ArticleGoogle Scholar
  143. Sica A, Mantovani A. Macrophage plasticity and polarization: in vivo veritas. J Clin Invest. 2012;122:787–95. doi:https://doi.org/10.1172/JCI59643.PubMedPubMed CentralView ArticleGoogle Scholar

Copyright

© The Author(s) 2016

Advertisement