Vitiligo is a cutaneous pigmentary disorder caused by selective destruction of melanocytes and is characterized by progressive, patchy loss of pigmentation from skin. Vitiligo is the most common disorder of pigmentation, affecting ∼1–2% of the world’s population, generally occurring during the second decade of life 1. Clinically, there are two types of vitiligo. Localized vitiligo can be segmental and present in a dermatomal distribution or may be focal and present in an asymmetric distribution involving segments of skin. Generalized vitiligo (GV) is characterized by multiple scattered lesions in a symmetrical distribution. The course of vitiligo has an unpredictable pattern, with progressive and stabilized depigmentation phases. Active vitiligo is defined by enlarging lesions or formation of new lesions 2. The impact of vitiligo on patients’ self-esteem and social interactions can be devastating, particularly when milk-white skin lesions appear in darkly pigmented individuals 2.
Before investigating the etiology of vitiligo, investigators first had to prove that melanocytes were absent in the patches of depigmentation in vitiligo. Immunostaining with antibodies against different melanocyte antigens supported the loss of differentiated melanocytes from vitiligo lesional skin. In addition, c-KIT staining for undifferentiated melanocyte stem cells showed that vitiligo lesional skin is also devoid of melanocytic precursors 3.
The ëxact pathophysiology of vitiligo is not fully understood. There are a few major hypotheses for the pathogenesis of vitiligo: (i) Autoimmune pathogenesis is a long-standing and popular hypothesis; (ii) the neural hypothesis suggests that nerve endings release neurochemical substances that can decrease melanin production or damage melanocytes 1; (iii) the biochemical hypothesis implicates the accumulation of toxic intermediate metabolites of melanin synthesis 4 and defective free radical defense 5, and the build-up of excessive quantities of hydrogen peroxide (H2O2) as a cause for destruction of melanocytes 6. Other hypotheses include genetic factors, defects in the structure and function of melanocytes, and deficiency in melanocyte growth factors are playing a role in the depigmentation process.
Not one hypothesis can explain the pathogenesis of vitiligo on its own. The convergence theory proposes that many of the hypotheses play a role in vitiligo. In addition, some hypotheses play a larger role than others in certain manifestations of vitiligo. For example, GV has more supporting evidence for the autoimmune hypothesis, whereas segmental vitiligo has more supporting evidence for the neural hypothesis 7–11.
Vitiligo is a polygenic, multifactorial disorder that involves multiple susceptibility genes 12–14. Epidemiological studies have shown that family clustering of vitiligo occurs frequently. There is also an increased risk of vitiligo in first-degree relatives and high concordance in monozygotic twins 14,15. Approximately one in four vitiligo patients present with additional autoimmune disorders including pernicious anemia, autoimmune thyroid disease, psoriasis, rheumatoid arthritis, adult-onset autoimmune diabetes mellitus, Addison’s disease, and systemic lupus erythematosus 15,16. These same autoimmune disorders also occur at an increased frequency among vitiligo patients’ first-degree relatives, further supporting that increased risk of vitiligo and other autoimmune disorders in these families have a genetic basis.
A number of genes have been implicated in the pathogenesis of vitiligo on the basis of genetic linkage and association studies. A gene encoding a key regulator of the innate immune system 17, NALP1, was previously found to be associated with GV in two different groups of White patients 18,19. In the White population, there were at least two independent risk signals [single-nucleotide polymorphism (SNP) rs6502867 and another SNP tagged by rs2670660 and rs8182352]. In a more recent study that examined Arab patients, two risk signals (SNP rs1008588 and rs2670660) were located upstream in the promoter region of NALP1 and showed a significant association with the vitiligo phenotype. These findings indicate that variation in NALP1 is associated with susceptibility to GV in Arabs and Whites 20.
Numerous genetic studies of biological candidate genes for GV have been published 21. Most candidate genes for GV, besides human leukocyte antigen (HLA) and PTN22, have not been supported consistently because these studies have been limited by false-positive associations. Thus, candidate gene studies have shifted toward genome-wide association studies (GWAS), which control for causes of a false association. Studying, a GWAS of GV was carried out in non-Hispanic White patients, identifying and confirming at least 13 different loci that contribute to the risk of GV, almost all of which have immunoregulatory functions 22. Furthermore, this study carried out one of the largest association studies of previously proposed GV candidate genes by analyzing SNP data for 33 genes in a large White case–control data set. There was evidence for a primary association with GV for only three of the candidate genes tested (FOXP3, TSLP, and XBP1) and the meta-analysis strongly supported XBP1 as a true GV susceptibility locus. A fourth locus, CTLA4, was found to be secondarily associated with GV and primarily associated with the autoimmune diseases that are associated with GV 23.
Alterations to epigenetic DNA methylation patterns contribute toward many autoimmune diseases, including vitiligo. A study carried out by Zhao and colleagues has confirmed a significant increase in global peripheral blood mononuclear cells (PBMCs) DNA methylation in patients with vitiligo compared with control participants. They identified a pattern of aberrant genomic DNA methylation and reduced expression of DNA methyltransferases and methyl-DNA-binding domain proteins that play an active role in de-novo methylation and in regulating DNA methylation, respectively. In addition, there was decreased mRNA transcription of interleukin (IL)-10 in PBMCs that correlated with hypermethylation of the IL-10 enhancer region 24. Decreased IL-10 levels in vitiligo may result in an increased CD8+ T-lymphocyte response and decreased maturation of T regulatory cells, facilitating the development of vitiligo 25–27.
Another study found that transforming growth factor β receptor II (TGFBR2) gene contributes toward the susceptibility for NSV in the Korean population. All three SNPs (rs3773649, rs2005061, and rs3773645) were significantly different between vitiligo patients and controls and haplotype 1 (CG) and haplotype 2 (GA) were also associated with a risk of NSV. Reduced serum TGFBR2 levels, as observed in patients in this study, may contribute toward enhanced cellular immunity. This may facilitate the occurrence of vitiligo by leading to reduced maturation of regulatory T cells, followed by impaired inhibition of inflammation 25.
The HLA is now recognized as a major contributing factor for susceptibility to a variety of autoimmune diseases, and numerous associations with vitiligo have focused on the HLA system. Multiple HLA class II alleles have been found to be associated with vitiligo in different populations, such as DRB4*0101 and DQB1*0303 in Dutch patients 28, DRB1*04 and DRB1*07 in Turkish patients 29, and DRB1*07 in Chinese Han patients 30–33.
Dunston et al. 34 identified DR4 to be associated with a positive family history of autoimmune disease and early onset of disease. Fain et al. 35 reported that HLA class II haplotype DRB1*04–DQB1*0301 contributes toward the risk of familial GV and early onset of disease. Orozco-Topete et al. 36 reported that DRB1*04 is associated with the genetic susceptibility of developing vitiligo with autoimmune thyroid disease.
DRB1*07-positive patients had earlier disease onset and higher frequency of family history compared with DRB1*07-negative patients. A contribution of immune activation toward progressive depigmentation is also supported by abnormal expression of HLA-DR in perilesional epidermis, which is likely relevant in explaining destruction of melanocytes as melanocytes can present antigens in the context of major histocompatibility complex (MHC) class II 37–39. All these mostly indicate that some genetic component of autoimmune susceptibility might play a key role in the pathogenesis of DRB1*07-associated vitiligo 40.
Misri and colleagues 41 carried out a comparative case–control study of clinical features and HLA susceptibility between familial and nonfamilial vitiligo. Family history was associated with HLA A2, A28, A31, and B44 alleles. Early onset of vitiligo (<20 years) was associated significantly with HLA A2, A11, B17, B35, and B44 alleles. The patients with severe affection (>10% area) showed insignificant association with HLA A10 and B8. A family history of vitiligo is associated with an early onset of vitiligo. There is no correlation of family history with the type of vitiligo, stability of lesions, and areas involved.
Silva De Castro and colleagues 42 carried out a combined family-based and case–control association analysis between vitiligo and genetic variants of the discoidin domain receptor (DDR1), which is involved in the regulation of melanocyte attachment to the basement membrane of the epidermis. Adhesion of melanocytes to the basement membrane by integrin CCN3 is mediated through collagen IV receptor DDR1. Allele T (SNP rs4618569) was associated with an increased risk for vitiligo in the family trios. Allele C (SNP rs2267641) was associated with an increased risk for vitiligo in both family-based and case–control populations. The best evidence for an association in the trios was obtained for a haplotype composed of risk alleles of markers rs4618569 and rs2267641. There was an age-dependent enrichment of the rs4618569 T allele and the rs2267641 C allele in early-onset affected individuals. In conclusion, DDR1 has been proposed as a susceptibility gene for vitiligo, possibly implicating a defective cell adhesion 42.
The ultraviolet radiation resistance-associated gene (UVRAG) polymorphism may contribute toward increased susceptibility to NSV in the Korean population and may also be one of many genes confirmed to play a role in polygenic susceptibility to NSV. UVRAG activates the Beclin1–PI(3)KC3 complex, promoting autophagy 43. Autophagy may contribute toward the development and maintenance of autoimmunity by the promotion of the MHC class II presentation of cytosolic antigens and the activation of CD4+ T lymphocyte 44. Two SNPs (rs1458836, rs7933235) showed significant genotypic differences between the NSV patient group and the control group. In addition, the haplotype of the UVRAG polymorphism was associated with NSV. This study suggests a possible association between UVRAG and NSV susceptibility 45.
Jin and colleagues 22 supported the long-standing hypothesis that GV involves genetic susceptibility loci shared with other autoimmune diseases. A GWAS of European-derived White patients with vitiligo was performed and 10 independent susceptibility loci for GV were identified. With the exception of PTPN22 (rs2476601), these associations were observed in both the subgroup of patients with vitiligo only and the subgroup with vitiligo and concomitant autoimmune diseases. There was a significant association between GV and SNPs at several loci associated previously with other autoimmune diseases. These included genes encoding MHC class I molecules and class II molecules, PTPN22, LPP, IL2RA, UBASH3A, and C1QTNF6. There were also associations between GV and SNPs in two additional immune-related loci, RERE and GZMB, and in a locus containing TYR, encoding tyrosinase. Most of these genes encode immune-system proteins involved in biologic pathways that probably influence the development of autoimmunity.
Quan and colleagues 38 carried out a large-scale GWAS of GV in the Chinese Han population, providing new insights into the genetic predisposition to vitiligo. A previously undescribed risk locus at 6q27 (rs2236313) that contains the RNASET2 (encoding RNase T2), FGFR1OP (fibroblast growth factor receptor 1 oncogene partner), and CCR6 (encoding chemokine receptor 6) genes was also identified. Overexpression of RNASET2 hypersensitizes cells to oxidative stress, thus promoting cell death during peroxide exposure and stationary-phase onset 46. It has been postulated that oxidative stress might be the initial pathogenic event in melanocyte destruction in vitiligo 47. FGFR1OP is necessary for cell-cycle progression and survival 48. CCR6 is preferentially expressed by immature dendritic cells and facilitates chemoattraction of immune cells 49. This study also highlighted a potential pathogenic overlap between vitiligo and other autoimmune diseases, especially inflammatory bowel disease (IBD). Notably, a common variant (rs2301436) at 6q27 has been associated with Crohn’s disease and showed a moderate association with vitiligo in the GWAS analysis 38.
SMOC2 is an important novel candidate gene for susceptibility to GV, and perhaps to other autoimmune diseases, in the isolated Romanian founder population. A GWAS for GV was carried out in distantly related affected patients from a unique founder population in an isolated Romanian community with an increased prevalence of GV and healthy controls from surrounding villages. Vitiligo was significantly associated with a SNP in the SMOC2 gene on chromosome 6q27 (rs13208776). This SNP was in close vicinity to IDDM8, a linkage and association signal for type I diabetes mellitus 50–53 and rheumatoid arthritis 54. Type I diabetes mellitus and rheumatoid arthritis are autoimmune diseases that are epidemiologically associated with GV 15,55. In the skin, SMOC2 is mainly present in the basal layer of the epidermis and SMOC2-stimulated attachment of primary keratinocytes in culture 56. These findings are of interest in light of the suggestion that melanocyte loss in vitiligo might result from chronic cell detachment because of defective cell adhesion 57.
MYG1 and tyrosinase mRNA expression levels were found to be affected in the skin of vitiligo patients. MYG1 plays a role in the regulation of pigmentation, which makes it a possible candidate gene of functional importance in the pathogenesis of vitiligo 58,59. In addition, a downregulation of tyrosinase mRNA expression has been documented in lesional skin of vitiligo patients 60. In this study, a statistically significant decrease in mRNA expression of tyrosinase, an essential marker for melanogenesis, was observed in lesional skin of vitiligo patients compared with nonlesional skin of vitiligo patients and healthy control participants. No differences in the expression of MYG1 were detected between different subgroups of vitiligo on the basis of the extent of involvement and progression of the disease. In general, MYG1 mRNA expression was increased in each subgroup of vitiligo and was not dependent on whether the biopsy had been taken from lesional or unaffected skin. Consistent overexpression of MYG1 in the skin of vitiligo patients might suggest a systemic involvement of MYG1 in the pathogenesis of vitiligo 61.
Kingo and colleagues 61 further explored the role of MYG1 in the pathogenesis of vitiligo by examining polymorphisms in the MYG1 gene. Control participants with the −119G promoter allele (rs1465073) showed significantly higher MYG1 mRNA levels than controls with −119C allele. Single-marker association analysis showed that the −119G allele was more frequent in vitiligo patients compared with the controls. Analysis on the basis of the stage of progression of the vitiligo showed that the increased frequency of the −119G allele occurred prevalently in the group of patients with active vitiligo compared with the control group.
The neural hypothesis is supported widely by clinical, ultrastructural, and biochemical findings and can explain the destruction of epidermal pigment cells. Two types of vitiligo have been proposed: type A vitiligo that has a nondermatomal distribution and is believed to arise from autoimmune disease and type B vitiligo that has a dermatomal distribution and arises from the dysfunction of sympathetic nerves that innervate the affected dermatome 62,63. Melanocytes were first believed to be under neural control because melanocytes originate from neural crest cells; thus, the degeneration of nerves and nerve endings was proposed as a possible mechanism for vitiligo 64–66.
Pathologic conditions involving the nervous system support the neural theory in the pathophysiology of vitiligo. In patients with transverse myelitis, there is diffuse hypopigmentation in the areas innervated by nerves originating from the level of the damaged spinal cord; however, hypopigmentation is spared in areas below the spinal cord damage 67. Vitiligo often times follows areas of peripheral nerve injury such as hypopigmentation along the arm when the brachial plexus is injured 68. Neurodysplasias such as neurofibromatosis and tuberous sclerosis can present with both hyperpigmentation and hypopigmentation 69,70. In addition, inflammatory conditions that affect the nervous system such as leprosy, syphilis, pinta, and viral encephalitis can present as cutaneous and hair hypopigmentation or depigmentation. Often times, the arrest of the pathologic condition leads to repigmentation 71,72.
The neural hypothesis is also supported by animal studies. Initially, denervation was found to affect hamster skin pigmentation 73 and acetylcholine, norepinephrine, epinephrine, and melatonin were found to lighten dermal melanocytes in frogs 67. In a rabbit study, there was gradual lightening of the iris color after interrupting the superior cervical ganglion and preganglionic nerve trunk because of inadequate replenishment of melanin within the iris stromal melanocyte. This is consistent with previous reports of direct adrenergic innervation of iris stromal melanocytes 74–76. Further studies showed that stimulation of the α nerve fibers lightens skin color in animals by causing melanosomes to aggregate, whereas opposite stimulation disperses the melanosomes to cause darkening of the skin 77.
Histological studies in patients with vitiligo show inflammatory and degenerative changes in the dermal nerves and nerve endings. Microscopic and ultrastructural studies in humans have shown that there is direct contact between nerve fibers and melanocytes in vitiligo skin 78 and there are dystrophic changes in the nerve trunk and terminals 79. In vitiligo lesions, one study showed evidence of both axonal degeneration and nerve regeneration, most likely in response to previous degeneration. There was a significantly increased thickness in the basement membrane of Schwann cells in the dermal nerves of vitiligo lesion biopsies; however, indicators of regeneration such as increased mitochondria and rough endoplasmic reticulum predominated. This study, however, did not show a major difference in morphology between the central and the marginal parts of the vitiligo lesion, whereas other studies did show more marked changes in morphologies between these two regions 80. The reason for this discrepancy is unclear.
Patients with vitiligo often report that their disease starts after an emotional or a stressful event and/or after physical injury (Koebner phenomenon). Stressful situations are often times associated with catecholamine release from presynaptic cells. Stress is also known to activate neurotransmitter-rich areas of the brain, increase the levels of neuroendocrine hormones 81, and alter the levels of peripheral neurotransmitters in the body 82. One study found that stressful events were a precipitating or an aggravating factor in 65% of patients with vitiligo (odds ratio=6.81). In addition, patients with vitiligo are known to have higher scores for anxiety 83, depression 84, adjustment disorders 85, and obsessive symptoms, and hypochondria 86. In one study, 24 h urine samples were collected from patients with early-phase and stable-phase vitiligo and healthy controls. Early-phase and progressive-phase vitiligo patients showed significantly increased levels of homovanillic acid and vanilmandelic acid than those of controls, whereas stable-phase vitiligo patients and controls showed no difference. The high circulating levels of homovanillic acid and vanilmandelic acid can represent a reaction to a stressful event 87.
The increased levels of catecholamines released from autonomic nerve ending can affect the microenvironment of melanocytes in several ways. Catecholamines and their o-diphenol catabolites can have a direct cytotoxic effect. Phenols can easily undergo oxidation to form toxic quinones, semiquinone radicals, and oxyradicals. Catecholamines can also cause arteriolar vasoconstriction by binding to α receptors, which would lead to local epidermal and dermal hypoxia with the overproduction of toxic radicals. Further supporting the role of the release of catecholamines from autonomic nerve endings and keratinocytes in the pathogenesis of vitiligo is a study 88 that found an increase in cutaneous microcirculation and α and β adrenoreceptor function in segmental-type vitiligo. A laser Doppler flowmetry and iontophoresis were used to assess cutaneous microcirculation in patients with nonsegmental and segmental vitiligo. There was a three-fold increase in cutaneous blood flow in segmental-type vitiligo compared with the normal skin on the contralateral side. Phenylephrine, clonidine, and propranolol were administered and the duration of recovery from vasoconstriction was measured to assess α and β receptor function. In segmental-type vitiligo, there was an increase in cutaneous α and β adrenoreceptor response 88–90.
Descending autonomic nerves can stimulate the release of neuropeptides in the skin 91, and clinical and biochemical studies have suggested that some neuropeptides and neuronal markers are involved in the destruction of melanocytes observed in vitiligo. Al’Abadie et al.92 found a strong increase in neuropeptide Y (NPY) and minimal increases in vasoactive intestinal polypeptide and a general neuronal marker PGP 9.5 in affected vitiligo lesions compared with nonvitiligo control skin. A subsequent study 93 used the immunofluorescence technique to examine neuropeptides (substance P, somatostatin, calcitonin gene-related peptide, and NPY) in lesional, marginal, and nonlesional skin to further characterize the distribution of neuropeptides in the skin. This study found that there was increased immnoreactivity against NPY and to a lesser extent, calcitonin gene-related peptide (CGRP) in the depigmented skin lesions.
There are several proposed mechanisms by which neuropeptides, especially NPY, could lead to vitiligo. NPY could be released by external factors such as physical trauma (Koebner phenomenon) or internal factors such as stress. NPY could consequently trigger a cascade of reactions that affect the immune system. Lymphocytic infiltration is associated with melanocytic destruction in vitiligo lesions and certain neuropeptides such as NPY have the capability to attract lymphocytes to the dermis 94. In addition, the affected areas in vitiligo have been shown to have vasoconstriction and altered levels of sweating 95. NPY has been found to colocalize with norepinephrine in nerves in human skin and often function in parallel as cotransmitters and comodulators 96. NPY was found to be able to exert a local autonomic effect that included vasoconstriction 97.
In addition to the altered balance of neuropeptides in vitiligo skin, there is also a change in the neuronal structural markers in the peripheral nerve fibers in affected and unaffected vitiligo skin compared with normal healthy skin. The number of nerve growth factor receptor immunoreactive (NGFr-IR) and calcitonin gene-related peptide immunoreactive (CGRP-IR) nerve fibers were increased in involved skin compared with control skin. This study also found a reduction in the number of NGFr-IR in basal keratinocytes in involved vitiligo skin compared with control skin. Interestingly, the study found that NGFr-IR nerve fibers connect to these epidermal basal keratinocytes. Other studies have supported this finding by showing similar nerve fiber attachments to keratinocytes and Merkel cells 98.
Although the mechanism of action of NGF is not known, NGF clearly affects melanocyte function. NGF is found in all tissues innervated by sympathetic neurons 99. NGF has been shown to be critical in the development and maintenance of the function of the peripheral sympathetic neurons and neural crest-derived sensory neurons 100,101. NGF is an important survival factor for human melanocytes. In cultured human melanocyte cell lines, NGF has been shown to modulate melanocyte gene expression and exposure to NGF gradient increases the migration of melanocytes and melanocytic dendrites. Nerve endings have been shown to connect to normal and degenerated melanocytes 78,80,102. The observation that NGFr-IR fibers were more numerous and were more intensely labeled in affected vitiligo skin suggests that the release of NGF from peripheral nerves or the overexpression of NGFr-IR by melanocytes may play a role in the destruction of epidermal pigment cells.
In summary, it has been suggested that the overflow of neuropeptides such as NGF, CGRP, NPY, and vasoactive intestinal polypeptide through the NGFr-IR/CGRP-IR nerve fiber pathways can affect the microenvironment of melanocytes and lead to the destruction of melanocytes observed in vitiligo.
Vitiligo is widely considered to have an autoimmune basis; however, the specific trigger and precise nature of the autoimmune response still remain unclear. This hypothesis is broadly supported by the observation that many autoimmune diseases such as autoimmune thyroid disease, pernicious anemia, Addison’s disease, systemic lupus erythematosus, and IBD are associated with vitiligo 15,16. There is a significant association of vitiligo with thyroid dysfunction and thyroid antibodies 102–104, which is the reasoning behind regular thyroid screening in patients with vitiligo.
The initiation and response of vitiligo to various immune-based therapies have also supported an autoimmune hypothesis. Ultraviolet radiation and steroids have an immune-suppressive effect and can cause repigmentation in vitiligo 105–110. Steroids especially have an immune-suppressive and modulatory effect by decreasing the accumulation of lymphocytes and monocytes at sites of inflammation. Cyclosporine, which directly inhibits T-cell-mediated immunity, delays the onset and decreases the incidence and severity of vitiligo 110,111. In contrast to this, various immune-modulating therapies can actually induce vitiligo. For example, interferon (IFN) therapy for hepatitis C can cause the development of vitiligo and there is subsequent repigmentation after discontinuing the therapy 112,113. In addition, allogeneic bone marrow transplantation can cause the transfer of vitiligo through pathogenic lymphocytes 114,115 and lymphocyte infusion from donors without vitiligo to treat relapsed leukemia can also induce GV through new alloreactive lymphocytes 116.
Depigmentation seen in various diseases also supports autoimmunity as a cause of vitiligo. There have been case reports of vitiligo-like depigmentation occurring in patients who had flares in their mycosis fungoides, suggesting that tumor or reactional lymphocytes are cytotoxic against melanocytes or can induce the production of autoantibodies 117.
The autoimmune hypothesis is composed of humoral and cell-mediated components.
Role of humoral response
Studies have shown that patients with vitiligo have an increased concentration of circulating autoantibodies that are specific to melanocytic cytoplasmic and surface antigens compared with normal individuals, suggesting that a humoral response may play a role in the pathogenesis of vitiligo 118–121. The presence of these antibodies is related to the extent of the disease 122. Immunohistologically, IgG and C3 deposits have occasionally been found in the basement membrane zone and in keratinocytes of vitiligo lesions and have fluoresced more brightly in active and extensive disease 118,121,123–125. Some studies, however, suggest that IgA levels better correspond with vitiligo activity 126. There are also a higher percentage of B cells in patients with recent onset of disease, suggesting an involvement of humoral immunity in early vitiligo 127.
Antibodies in vitiligo are directed against specific melanocyte antigens. Some of the proteins are common tissue antigens, whereas others are preferentially expressed on pigment cells 128. Tyrosinase and tyrosinase-related proteins 1 and 2 (TRP-1 and TRP-2) are enzymes localized to melanosomes and are involved in the synthesis of melanin. TRP-1 was found to be expressed on the surface of both melanocytes and melanoma cells. In total, 40–60% of patients with vitiligo patients had autoantibodies to tyrosinase on immunoblotting experiments 129,130. A similarly high percentage (67%) of patients with vitiligo was also found to have anti-TRP-2 antibodies 131. Furthermore, autoantibodies to TRP-1 have been identified in a minority (5%) of individuals with vitiligo 132,133. Other studies did not show results that were as conclusive 134. In addition, tyrosinase antibodies have been found to cross react with TRP-1 and TRP-2, suggesting that a shared epitope may be the target among all of these potential autoantigens 132,133,135. Melanin-concentrating hormone receptor 1 (MCHR1) was also explored as potential melanocyte-specific autoantigens. A sizeable percentage (16.4%) of patients with vitiligo have autoantibodies to MCHR1 and these antibodies are highly disease specific 136. However, the MCHR1 was ultimately not found to be expressed by melanocytes 137.
Whether antibodies are the cause of vitiligo or whether they arise as the result of the disease is currently unclear. In theory, pigment cell antibodies can arise from the following: (i) genetic disposition to immune dysregulation of T cells, (ii) cross-reacting antigens expressed on other target cells or on infecting microorganisms, and (iii) an immune response to damaged pigment cells. In addition, the mechanism of antibody-mediated melanocyte destruction is still unclear. More broadly, the incidence and serum levels of antibodies to melanocyte antigens correlate with disease activity, the extent of depigmentation, and the presence of autoimmune disease 118,122,138. The level of autoantibodies decreases in vitiligo patients who respond to PUVA therapy or following systemic steroid treatment 139,140. On a more microscopic basis, vitiligo antibodies can destroy melanocytes. In-vitro studies have shown that cultured human melanocytes can be damaged by vitiligo sera, which induces complement activation and antibody dependent cellular cytotoxicity 141. In-vitro studies showed melanocyte destruction on skin grafts when nude mice grafted with human tissue were injected with antibodies from vitiligo sera 142. In addition, IgG purified from vitiligo patients can destroy melanoma cells in vivo and in vitro143. In terms of MCHR antibodies in vitiligo patients, they have been shown to be capable of blocking stimulation of the receptor by melanin-concentrating hormone in a heterologous cell line and thus adversely affect pigment cell functioning. In-vitro and in-vivo studies for MCHR antibodies have not been conclusive as yet 144.
Role of cell-mediated response
Histological and immunohistochemical studies in perilesional skin suggest the involvement of cellular immunity in vitiligo. The perilesional skin of patients with GV is consistently accompanied by cellular infiltrates containing CD4+ and CD8+ T lymphocytes and with an increased CD8+/CD4+ T-cell ratio. Occasional T cells remain in lesional skin; however, most migrate to the periphery with the depigmenting epidermal border. Because cytotoxic T cells colocalize with the remaining melanocytes, it has been postulated that T cells are actively cytolytic toward the remaining melanocytes 145. An activation of the T-cell-mediated immune system was confirmed by expression of activation molecules such as soluble IL-2 receptor, HLA-DR, and MHC class II 146–149. Furthermore, serum-soluble IL-2 receptors and lesional tissue IL-2 were found to be significantly increased in generalized, focal, and nondermatomal types of vitiligo and correlated with increased disease activity 150–152. Interestingly, 26% of patients with melanoma responding to IL-2-based immunotherapy also developed vitiligo 153, suggesting that the same T cells that are involved in the regression of melanoma are also involved in the destruction of normal melanocytes. More recently, a positive correlation was found between patient soluble IL-2 receptor (sIL-2R) levels and body surface area involvement in the North American population, which may reflect the increased T-lymphocyte activation and aggressiveness of the disease. This data raise the possibility that serum sIL-2R may serve as an immunological marker to assess the severity and prognosis of disease as well as therapy response for nonsegmental vitiligo (NSV) 154.
The T cells are actively recruited from the peripheral circulation to the affected sites in vitiligo. Depigmentation is accompanied by expression of type 1 cytokines. IFN-γ is increased in lesion and perilesional skin, which enhances T-cell trafficking to the region by increasing ICAM-1 expression 147. In fact, neutralization of IFN-γ with antibody has been shown to prevent CD8+ T-cell accumulation and depigmentation, which highlights the importance of this cytokine and a potential therapeutic target 155. A huge number of T cells express cutaneous lymphocyte antigen (CLA), which is typically expressed by T cells that are ‘en route’ to the skin 39. Skin-homing CLA+/CD8+ T cells may play a significant role in the destruction of melanocytes seen in GV. There was an increased migration of activated skin-homing CLA+/CD8+ T cells and macrophages in the perilesional skin, with CLA+ T cells concentrated around the area of disappearing melanocytes and melanocyte remnants along the dermoepidermal junction. The majority of these interacting T cells were HLA-DR+, indicating their activated state. The T cells also expressed granzyme, indicating that cytotoxicity and apoptosis are mediated by the granzyme/perforin pathway rather than Fas/FasL in vitiligo. Multiple studies have also detected high levels of melan-A-specific CD8+ T cells in the peripheral blood of patients with vitiligo, with their presence correlated with the extent and activity of the disease 156,157.
Both melanocytes and keratinocytes participate in the abnormal local immune reaction by antigen presentation. There is focal HLA-DR expression in the perilesional melanocytes and focal epidermal expression of ICAM-1 and HLA-DR in the perilesional keratinocytes. ICAM-1 and HLA-DR play a role in normal antigen presentation and T-cell activation 37,39,147,158. Keratinocytes may contribute toward the abnormal local immune reaction in vitiligo by presenting melanocytic antigens in a MHC class II-restricted manner after phagocytosis of melanosomes. Melanocytes have been shown to have phagocytic capabilities and can present antigens to MHC class II-restricted T cells in addition to being target cells for T-cell-mediated cytotoxicity 11,159,160. Thus, HLA-DR-positive melanocytes actively participate in the local immune response and become targets for cytotoxic T cells. Keratinocytes can also present melanocyte antigens in a MHC class II-restricted manner after phagocytosis of melanosomes 39. Furthermore, the increased migration of macrophages could be involved in clearing apoptotic melanocytes 161 or direct melanocyte destruction through the nitric oxide pathway 162.
Vitiligo T cells have shown reactivity to antigens recognized previously as target antigens for T cells infiltrating melanoma tumors. Development of vitiligo in melanoma patients is an indicator of effective immunity against the tumor and vitiligo in melanoma patients is considered a positive prognostic factor 163. Melan-A/MART1, gp100, tyrosinase, and TRP-1 and TRP-2 are immunogenic antigens that are commonly expressed by both melanocytes and malignant melanoma. These molecules are located in melanosomes and are required for effective melanization. T-cell-mediated responses to normal melanocyte differentiation antigens (gp100, melan-A/MART1, tyrosinase) were found in vitiligo patients 164. A high frequency of skin-homing, melan-A-specific, CD8+ T cells is detected in the peripheral blood of vitiligo patients 156. Immunotherapy for malignant melanoma can induce T-cell-mediated vitiligo. Melan-A/MART1-specific CD8+ T-cell clones were infused into melanoma patients for treatment and these same T cells were found near normal melanocytes that were undergoing destruction. Melan-A peptides have been used to immunize patients against melanoma. Although there was a regression in the melanoma, melan-A peptides also increased the incidence of vitiligo 165. In addition, melan-A/MART1-specific CD4+ T cells can be generated in patients with melanoma 166.
The role for Langerhans cells (LCs) in the depigmentation process of vitiligo is currently unclear. LCs are found to vary during the course of the disease. There is typically a decrease in LCs during active and repigmenting nonsegmental vitiligo and an increase in LC during stable nonsegmental vitiligo. The LCs may be getting destroyed by local cytotoxic effects or may be migrating to lymph nodes to facilitate antigen presentation 167.
Role of cytokines
There is a growing evidence that cytokines are important in the depigmentation process of vitiligo. Melanocytes and keratinocytes have been hypothesized to have a close relationship where keratinocyte-derived cytokines allow for melanocyte survival and differentiation 168. In nonsegmental, active vitiligo skin, there is a significantly lower expression of granulocyte monocyte colony stimulating factor (GMC-SF), fibroblast growth factor (FGF), stem cell factor (SCF) and a significantly higher expression of IL-6 and tumor necrosis factor α (TNF-α) compared with perilesional, nonlesional, and healthy skin 169. As these cytokines are produced by keratinocytes, it is likely that these epidermal cells are impaired in vitiligo. The exact mechanism of how cytokines affect pigmentation is not fully understood. A few different hypotheses are as follows: (a) GM-CSF, SCF, and bFGF are involved in stimulating melanocytes 170–172 and IL-6 and TNF-α are involved in decreasing the activity of the enzyme tyrosinase and also inhibiting melanocyte proliferation 173,174, and thus a change in cytokine production of epidermal microenvironments may be involved in vitiligo. (b) TNF-α induces IL-1a and promotes B-cell differentiation and immunoglobulin production 175. (c) TNF-α and IL-6 can induce cell surface ICAM-1 on melanocytes, which is necessary for leukocyte–melanocyte adhesion. ICAM-1 can also induce B-cell activation, increasing autoantibody production and may cause melanocyte damage in vitiligo 174. (d) Cytokines such as TNF-α and IL-1a released by keratinocytes, melanocytes, and lymphocytes also have the capacity to induce direct apoptosis in many cell types 175. Also, suppression of TNF-α with the new biological therapies may be associated with the repigmentation of vitiligo 176.
Tacrolimus, a topical immunomodulatory drug, induces repigmentation in vitiligo patients 176–179. Tacrolimus is believed to exert its therapeutic effects by inhibiting T-cell activation. Cytokine skin mRNA expression (TNF-α, IFN-γ, and IL-10) was increased at baseline in the adjacent normal skin of vitiligo patients and was reduced at these sites after tacrolimus treatment. This further supports that cytokine imbalance plays at least some role in the depigmentation process of vitiligo 180.
Reactive oxygen species model
One of the major hypotheses in the pathogenesis of vitiligo is the reactive oxygen species (ROS) model. The ROS model proposes that patients with vitiligo have an imbalanced redox state in their skin, resulting in excess production of ROS such as H2O2. H2O2 can undergo photochemical reduction and produce highly reactive hydroxyl radicals (OH) and hydroxyl ions (OH−) by the Haber–Weiss reaction. Hydroxyl radicals, in turn, are capable of affecting melanocytes by bleaching melanin and causing membrane lysis through lipid peroxidation. Thus, the accumulation of various ROS can exert toxic effects on melanocytes, further contributing toward depigmentation in vitiligo 181–183.
In the epidermis from patients with active vitiligo, an increased production of H2O2 has been reported and is associated with a lower expression and lower activity of the antioxidant enzyme catalase (CAT), suggesting that defective O2 metabolism along the entire epidermis may play a role in the pathogenesis of vitiligo. Furthermore, glutathione reductase was found to be significantly increased in vitiligo compared with controls 6.
In 2006, Ines and colleagues 184 showed that free radicals are increased and the antioxidant systems are insufficient in the blood of patients with vitiligo. This study measured the serum levels in patients with stable and active vitiligo and healthy controls for markers of redox status, which included malondialdehyde (MDA), selenium, vitamin E and A, and the erythrocyte activities of glutathione peroxidase (GPx), superoxide dismutase (SOD), and CAT. MDA is an end product of lipid peroxidation induced by ROS and correlates well with the degree of lipid peroxidation 181,182. Vitamin E is the only lipid-soluble antioxidant that is concentrated mainly in the erythrocyte and plasma membranes and is involved in terminating free radical reactions. Vitamin A is an essential, fat-soluble dietary compound that maintains the immune system and epithelial cell barriers 183. Selenium is incorporated covalently into GPx and thioredoxin reductase and is particularly important for the activity of these antioxidant enzymes. SOD scavenges superoxide radicals by converting O2− to form O2 and H2O2. CAT converts H2O2 into O2 and H2O. GPx is another antioxidant enzyme that converts H2O2 and other peroxides into water. GPx also catalyzes the reduction of hydroperoxides in the presence of glutathione (GSH) to form glutathione disulfide. Erythrocyte SOD activity was increased in respective vitiligo patient groups compared with their controls, which may be an adaptation to the increased oxidative stress that is present in these individuals and can enhance the systemic production of H2O2. MDA and selenium were increased, but most notably in active disease. Erythrocyte CAT activity and serum vitamin A and E levels were not significantly different from those of the controls. However, a significant decrease in GPx activity was observed in respective patient groups compared with their controls. CAT and GPx are downstream enzymes and a significant decrease in GPx was observed in the active vitiligo group, thus leading to further oxidative stress from H2O2 accumulation 184. At present, it remains unclear why serum selenium levels are at the upper normal level or even higher in the vitiligo group. High serum MDA levels may lead to lipid peroxidation in the membrane of melanocytes and may play an important role in the speedy depigmentation of active vitiligo.
Yildirim and colleagues 185 studied GPx activity in tissue of patients with GV and controls and determined that there was a higher level of GPx activity in patients with GV, supporting the more recent findings. A more recent study from Khan et al.186 found that there were lower levels of GPx in circulating erythrocytes, supporting the earlier study. From these results, GPx is increased in tissue, but its activity is decreased in the serum of vitiligo patients. In terms of CAT activity, Schallreuter and colleagues 6 showed epidermal H2O2 accumulation in association with low CAT levels. There was also a study on the CAT gene that associated this gene with susceptibility to vitiligo 187. Other studies have also shown lower CAT activity in vitiliginous melanocytes 188. The discrepancies in the results between different studies could also result from differences in serum and tissue levels, duration and activity of the disease, and differences in lab techniques.
Dell’Anna and colleagues 188 determined that mitochondrial impairment further contributed toward the ROS imbalance in addition to dysfunction of the antioxidant system by examining erythrocytes and PBMCs from patients with active or stable vitiligo and from healthy individuals. The study showed an imbalance in antioxidants (increased SOD and decreased CAT, reduced glutathione, and vitamin E levels) that was associated with higher ROS production with active vitiligo. This study was critical because it showed that the increased production in ROS was because of both the imbalance in antioxidants and mitochondrial impairment. Cyclosporine A, an inhibitor of the permeability transition pores (PTP) opening, reduced ROS production. In addition, an alteration in mitochondrial potential and a higher percentage of apoptotic cells were observed in active vitiligo. This suggests that ROS hyperproduction could be caused by opening of mitochondrial permeability transition pores. Mitochondria are key organelles in the control of intracellular ROS generation. Up to 5% of the oxygen consumed in the respiratory chain is converted into ROS 189–191. A variety of mitochondrial poisons increase mitochondrial superoxide production 191. Free radicals, oxidizing agents, modification of fatty acid pattern of the mitochondrial membrane, catecholamines, and cytokine release such as TNF-α, and increases in Ca2+ concentrations are events that may induce ROS production from mitochondria through PTP opening and alteration of the electron transport, which is consistent with what has been reported previously with active vitiligo 120,156,192. Thus, the mitochondrial dysfunction observed here could represent the possible target of different stimuli and the biochemical basis for the insurgence of the disease.
To better understand whether a defective function of mitochondria is involved in the pathogenesis of vitiligo, Dell’Anna and colleagues 193 determined the intracellular ROS levels and activity of the Krebs cycle enzymes and inhibitors of the electron transport chain in PBMC from a group of patients with active or stable disease and in healthy individuals. The results confirmed that in PBMCs of active vitiligo patients, an increased intracellular ROS production and an alteration in the antioxidant system (decreased CAT and GSH, increased SOD) occur, which is not associated with signs of systemic oxidative stress. The abnormal intracellular ROS generation seems to be an event that occurs upstream the antioxidant alteration and both the increased SOD activity and the reduction of the CAT activity, as well as depletion of GSH, could be considered a result of ROS hyperproduction. Moreover, in the same cells, they found an increased susceptibility to inhibitors of mitochondrial complex I and a marked increase in the expression of mitochondrial malate dehydrogenase activity. These data further suggest an altered functionality of mitochondria in vitiligo patients.
Dell’Anna and colleagues 193 once again provided new insight into the possible pathogenic mechanisms leading to melanocyte loss in vitiligo lesions by providing evidence that the alteration in the lipid components of the mitochondrial membrane could be a possible determining factor in intracellular ROS generation. Here, they studied epidermal primary melanocytes from nonlesional vitiligo skin and they observed altered ROS production, membrane lipoperoxidation, modification of the transmembrane CL distribution, and decreased Cxl activity [assess the electron transport chain (ETC)], as well as increased susceptibility to the chemical oxidant CuH 194. The expression and function of the mitochondrial ETC complex is strictly dependent on the lipid component of the inner mitochondrial membrane and particularly on cardiolipin 195. CL is a dimeric structure consisting of four fatty acyl chains, and is the most representative phospholipid of the inner membrane responsible for the correct housing and activity of the ETC proteins 196,197. An impaired assembly or recycling of CL is associated with mitochondrial defects 196. The structural and functional pattern shown here may represent the initial biochemical impairment occurring in vitiligo and account for the loss of melanocyte viability after oxidant challenge.
The melanin and catecholamine synthesis pathway is tightly controlled by cofactor 6BH4 and is a major target for H2O2 stress in vitiligo. The cofactor 6BH4 is recycled by pterin-4a-carbinolamine (PCD) and dihydropteridine reductase (DHPR) to allow phenylalanine hydroxylase to convert L-phenylalanine into L-tyrosine in the melanin and catecholamine synthesis pathway. Previous studies have shown that the 6BH4 recycling pathway is disrupted in vitiligo 192–199. The disruption of this cycle leads to the excess production of 7BH4, which strongly inhibits phenylalanine hydroxylase 200,201 and also leads to the generation of excess H2O2 instead of L-tyrosine 201. The general excess in H2O2 levels suppresses the enzyme activities of PCD and DHPR 199–202. This is further supported by therapeutically removing epidermal H2O2 with low-dose narrow-band ultraviolet B-activated pseudocatalase (PC-KUS), which returns PCD and DHPR activity back to normal and clinically causes repigmentation. These results prove that the entire 6BH4 recycling system is severely affected in vitiligo 192,199,203. These results also show that patients with vitiligo combat excess H2O2 stress both in the epidermis and in the vascular space, although there seems to be a much higher accumulation of H2O2 in the epidermis 48,199.
Furthermore, the dysregulation of the cholinergic transduction pathway by H2O2 oxidative stress is a novel mechanism that was found to contribute toward the depigmentation process in active vitiligo. Cholinergic involvement was first considered to play a role in vitiligo because the epidermal layer expresses muscarinic and nicotinic receptors and plays an integral role in the synthesis and breakdown of acetylcholine. In addition, the depigmented patches in vitiligo are associated with decreased sweating or cholinergic activity 204. The hydrolysis of acetylcholine is mediated by two key enzymes acetylcholinesterase (AchE) and butyrylcholinesterase (BchE). Both AchE and BchE have been shown to be distributed widely in epidermal tissue and in plasma. Iyengar first showed an association between low AchE levels and vitiligo 205. More recently, AchE and BchE protein expression and enzyme activities have been reported to be severely affected in vitiligo 206. To elucidate the mechanism behind this damage to the cholinergic system in vitiligo, in-situ and in-vitro studies have shown that both AchE and BchE are deactivated by H2O2 through H2O2-mediated oxidation of Met and Trp amino acid residues of both of these proteins in patients with active vitiligo. Topical pseudocatalase (PC-KUS) decreases oxidant burden and reactivates both AchE and BchE, and ultimately allows for repigmentation 195,207. Previous studies have shown that acetylcholine modulates pigment production by inhibiting dopa oxidase activity in melanocytes. Thus, low AchE leads to increased Ach and inhibition of pigmentation in melanocytes, further precipitating vitiligo.
Recent studies 208 have also shown that xanthine oxidase (XO) contributes toward the generation of oxidative stress in vitiligo. XO is an enzyme that facilitates purine degradation by catalyzing the hydroxylation of hypoxanthine to xanthine and finally to uric acid. This reaction produces H2O2 and allantoin is a product formed from the oxidation or uric acid. XO has been shown to be present in epidermal keratinocytes and melanocytes. Furthermore, allantoin, a marker for oxidative stress in purine degradation, was found to be present in both the lesional and the nonlesional skin of all patients with acute vitiligo, whereas it was absent in healthy controls. Thus, this presence of epidermal allantoin further supports the oxidative stress model in vitiligo.
The vast majority of studies involving the ROS model are from patients with GV. Thus, ROS-mediated damage is the most likely pathogenesis of nonsegmental GV.
A more recent hypothesis, melanocytorrhagy, emphasizes that depigmentation is because of chronic detachment of melanocytes. Trauma or repeated friction can contribute toward the detachment of melanocytes over time. The initial lesions in NSV often occur in areas of focal damage such as wound and excoriations and minor trauma such as pressure or repeated friction 209. Gauthier and colleagues 210 explored the consequence of repeated friction in the attachment and survival of melanocytes in nonlesional skin of patients with NSV. After 4 and 24 h of friction in nonlesional NSV skin, they found that many melanocytes had undergone detachment and could be found throughout the suprabasal locations of the epidermis. In addition, the detachment of melanocytes from the basement membrane of normal skin and subsequent migration to the upper layers of the epidermis has been observed after chemical stress and tape stripping 211,212. Although the cause of detachment has not been fully elucidated, severe frictional injury in nonlesional NSV skin has been shown to cause degeneration of keratinocytes, widening of intracellular spaces, and extracellular granular material deposits 213.
Melanocytes in unstable vitiligo have been found to lose the ability to adhere to key surrounding structures. Altered synthesis of extracellular matrix components by damaged keratinocytes may ultimately lead to the disappearance of melanocytes. Tenascin, an extracellular matrix molecule that inhibits adhesion of melanocytes to fibronectin, has been detected in the basal membrane in the papillary dermis and can contribute toward chronic detachment and epidermal loss of melanocytes. The origin of the tenascin deposits is currently unclear. In addition, perilesional skin melanocytes from patients with unstable vitiligo have shown significantly low adhesion to collagen type IV compared with control and stable vitiligo. These same melanocytes are also more susceptible to apoptosis as they expressed increased caspase 3 and annexin V 214.
Structural abnormalities in the dendrite itself may contribute toward melanocyte loss. The melanocyte adhesion system consists of structural junctions such as integrins, which mediate the interaction between melanocytes and the basement membrane and cadherins associated with β-catenin, which mediate the interaction between melanocytes and keratinocytes. In-vitro studies have not found a significant difference between the overall level of expression of integrins and cadherins between control, nonlesional, or lesional NSV skin 210,214. However, independent of the structural junctions, melanocytic dendrites may facilitate attachment of melanocytes to the basal layer of the epidermis and are considered to be a major component of the melanocyte adhesion system. NSV melanocytes were cultured and found to be small, with clubbed ends 215. These dendrites were retracted, whereas the dendrites seen in controls and patients with stable vitiligo were long and normally spread. There was no significant difference in the morphology between control and stable melanocytes. This indicates that dendritic retraction occurs before cell detachment and apoptosis. In addition, when H2O2, epinephrine, or norepinephrine was added to established cultures of vitiligo or normal control melanocytes, there was rapid retraction and loss of dendrites over 24 h 210,216. This finding ties together well the ROS, neural–biochemical, and melanocytorrhagy hypotheses as oxyradicals and catecholamines can adversely affect melanocyte dendricity, which could not only affect melanosome transfer but also adhesion to surrounding structures.
Conflicts of interest
There are no conflicts of interest.
1. Lerner AB. Vitiligo
. Prog Dermatol 1972; 6:1–6.
2. Kent G, Al’Abadie M. Psychologic effects of vitiligo
: a critical incident analysis. J Am Acad Dermatol 1996; 35:895–898.
3. Steel KP, Davidson DR, Jackson IJ. TRP-2/DT, a new early melanoblast marker, shows that steel growth factor (c-kit ligand) is a survival factor. Development 1992; 115:1111–1119.
4. Kemp EH, Waterman EA, Weetman AP. Autoimmune
aspects of vitiligo
. Autoimmunity 2001; 34:65–77.
5. Pawelek J, Korner A, Bergstrom A, Bologna J. New regulators of melanin biosynthesis and the autodestruction of melanoma cells. Nature 1980; 286:617–619.
6. Nordlund JJ, Lerner AB. Vitiligo
. It is important. Arch Dermatol 1982; 118:5–8.
7. Schallreuter KU, Wood JM, Berger J. Low catalase levels in the epidermis of patients with vitiligo
. J Invest Dermatol 1991; 97:1081–1085.
8. Ortonne JP, Bose SK. Vitiligo
: where do we stand? Pigment Cell Res 1993; 6:61–72.
9. Castanet J, Ortonne JP. Pathophysiology of vitiligo
. Clin Dermatol 1997; 15:845–851.
10. Norris DA, Horikawa T, Morelli JG. Melanocyte destruction and repopulation in vitiligo
. Pigment Cell Res 1994; 7:193–203.
11. Le Poole IC, Mutis T, Van den Wijngaard RMJGJ, Westerhof W, Ottenhoff T, De Vries RRP, Das PK. A novel, antigen-presenting function of melanocytes and its possible relationship to hypopigmentary disorders. J Immunol 1993; 151:7284–7292.
12. Majumder PP, Nordlund JJ, Nath SK. Pattern of familial aggregation of vitiligo
. Arch Dermatol 1993; 129:994–998.
13. Nath SK, Majumder PP, Nordlund JJ. Genetic epidemiology of vitiligo
: multilocus recessivity cross-validated. Am J Hum Genet 1994; 55:981–990.
14. Sun X, Xu A, Wei X, Ouyang J, Lu L, Chen M, Zhang D. Genetic epidemiology of vitiligo
: a study of 815 probands and their families from south China. Int J Dermatol 2006; 45:1176–1181.
15. Alkhateeb A, Fain PR, Thody A, Bennett DC, Spritz RA. Epidemiology of vitiligo
and associated autoimmune
diseases in Caucasian probands and their families. Pigment Cell Res 2003; 16:208–214.
16. Laberge G, Mailloux CM, Gowan K, Holland P, Bennett DC, Fain PR, Spritz RA. Early disease onset and increased risk of other autoimmune
diseases in familial generalized vitiligo
. Pigment Cell Res 2005; 18:300–305.
17. Ting JPY, Willingham SB, Bergstralh DT. NLRs at the intersection of cell death and immunity. Nat Rev Immunol 2008; 8:372–379.
18. Jin Y, Birlea SA, Fain PR, Spritz RA. Genetic variations in NALP1 are associated with generalized vitiligo
in a Romanian population. J Invest Dermatol 2007; 127:2558–2562.
19. Jin Y, Mailloux CM, Gowan K, Riccardi SL, Laberge G, Bennett DC, et al.. NALP1 in vitiligo
-associated multiple autoimmune
disease. N Engl J Med 2007; 356:1216–1225.
20. Alkhateeb A, Qarqaz F. Genetic association of NALP1 with generalized vitiligo
in Jordanian Arabs. Arch Dermatol Res 2010; 302:631–634.
21. Spritz RAPicardo M, Taieb A. Genetics
2010. Heidelberg: Springer-Verlag; 155–162.
22. Jin Y, Birlea SA, Fain PR, Gowan K, Riccardi SL, Holland PJ, et al.. Variant of TYR and autoimmunity susceptibility loci in generalized vitiligo
. N Engl J Med 2010; 362:1686–1697.
23. Birlea SA, Jin Y, Bennett DC, Herbstman DM, Wallace MR, McCormack WT, et al.. Comprehensive association analysis of candidate genes for generalized vitiligo
supports XBP1, FOXP3, and TSLP. J Invest Dermatol 2011; 131:371–381.
24. Zhao M, Gao F, Wu X, Tang J, Lu Q. Abnormal DNA methylation in peripheral blood mononuclear cells from patients with vitiligo
. Br J Dermatol 2010; 163:736–742.
25. Basak PY, Adiloglu AK, Ceyhan AM, Tas T, Akkaya VB. The role of helper and regulatory T cells in the pathogenesis
. J Am Acad Dermatol 2009; 60:256–260.
26. Garbelli S, Mantovani S, Palermo B, Giachino C. Melanocyte-specific, cytotoxic T cell responses in vitiligo
: the effective variant of melanoma immunity? Pigment Cell Res 2005; 18:234–242.
27. Sun L, Yi S, O’Connell PJ. IL-10 is required for human CD4+
regulatory T cell-mediated suppression of xenogeneic proliferation. Immunol Cell Biol 2010; 88:477–485.
28. Zamani M, Spaepen M, Sghar SS, Huang C, Westerhof W, Nieuweboer Krobotova L, Cassiman JJ. Linkage and association of HLA class II genes with vitiligo
in a Dutch population. Br J Dermatol 2001; 145:90–94.
29. Taştan HB, Akar A, Orkunoǧlu FE, Arca E, Inal A. Association of HLA class I antigens and HLA class II alleles with vitiligo
in a Turkish population. Pigment Cell Res 2004; 17:181–184.
30. Xiao Y, Zhao YM, Song FJ. Association of HLA-DRB1 alleles with generalized vitiligo
in Chinese Hans in north China. Chinese J Dermatol 2000; 33:5–7.
31. Wang J, Zhao YM, Wang Y, Xiao Y, Wang YK, Chen HD. Association of HLA class I and II alleles with generalized vitiligo
in Chinese Hans in north China. Chinese J Med Genet 2007; 24:221–223.
32. Wang XY, Jiang RH, Zhu MJ. Association of HLA-DRB1 alleles with vitiligo
and psoriasis vulgaris in Chinese Han people of Jilin region. Chinese J Immunol 2008; 28:144–146.
33. Ren Y, Yang S, Xu S, Gao M, Huang W, Gao T, et al.. Genetic variation of promoter sequence modulates XBP1 expression and genetic risk for vitiligo
. PLoS Genet 2009; 5:e1000523.
34. Dunston GM, Halder RM. Vitiligo
is associated with HLA-DR4 in black patients. A preliminary report. Arch Dermatol 1990; 126:56–60.
35. Fain PR, Babu SR, Bennett DC, Spritz RA. HLA class II haplotype DRB1*04-DQB1*0301 contributes to risk of familial generalized vitiligo
and early disease onset. Pigment Cell Res 2006; 19:51–57.
36. Orozco-Topete R, Córdova-López J, Yamamoto-Furusho JK, García-Benitez V, López-Martínez A, Granados J. HLA-DRB1*04 is associated with the genetic susceptibility to develop vitiligo
in Mexican patients with autoimmune
thyroid disease. J Am Acad Dermatol 2005; 52:182–183.
37. Badri AM, Todd PM, Garioch JJ, Gudgeon JE, Stewart DG, Goudie RB. An immunohistological study of cutaneous lymphocytes in vitiligo
. J Pathol 1993; 170:149–155.
38. Quan C, Ren YQ, Xiang LH, Sun LD, Xu AE, Gao XH, et al.. Genome-wide association study for vitiligo
identifies susceptibility loci at 6q27 and the MHC. Nat Genet 2010; 42:614–618.
39. Le Poole IC, van den Wijngaard RM, Westerhof W, Das PK. Presence of T cells and macrophages in inflammatory vitiligo
skin parallels melanocyte disappearance. Am J Pathol 1996; 148:1219–1228.
40. Hu DY, Ren YQ, Zhu KJ, Lv YM, Cheng H, Zhang Z, et al.. Comparisons of clinical features of HLA-DRB1*07 positive and negative vitiligo
patients in Chinese Han population. J Eur Acad Dermatol Venereol 2011; 25:1299–1303.
41. Misri R, Khopkar U, Shankarkumar U, Ghosh K. Comparative case control study of clinical features and human leukocyte antigen susceptibility between familial and nonfamilial vitiligo
. Indian J Dermatol Venereol Leprol 2009; 75:583–587.
42. Silva De Castro CC, Do Nascimento LM, Walker G, Werneck RI, Nogoceke E, Mira MT. Genetic variants of the DDR1 gene are associated with vitiligo
in two independent Brazilian population samples. J Invest Dermatol 2010; 130:1813–1818.
43. Liang C, Feng P, Ku B, Dotan I, Canaani D, Oh BH, Jung JU. Autophagic and tumour suppressor activity of a novel Beclin1-binding protein UVRAG. Nat Cell Biol 2006; 8:688–698.
44. Dengjel J, Schoor O, Fischer R, Reich M, Kraus M, Müller M, et al.. Autophagy promotes MHC class II presentation of peptides from intracellular source proteins. Proc Natl Acad Sci USA 2005; 102:7922–7927.
45. Jeong TJ, Shin MK, Uhm YK, Kim HJ, Chung JH, Lee MH. Association of UVRAG polymorphisms with susceptibility to non-segmental vitiligo
in a Korean sample. Exp Dermatol 2010; 19:e323–e325.
46. Thompson DM, Parker R. The RNase Rny1p cleaves tRNAs and promotes cell death during oxidative stress in Saccharomyces cerevisiae.
J Cell Biol 2009; 185:43–50.
47. Schallreuter KU. Successful treatment of oxidative stress in vitiligo
. Skin Pharmacol Appl Skin Physiol 1999; 12:132–138.
48. Acquaviva C, Chevrier V, Chauvin JP, Fournier G, Birnbaum D, Rosnet O. The centrosomal FOP protein is required for cell cycle progression and survival. Cell Cycle 2009; 8:1217–1227.
49. Le Borgne M, Etchart N, Goubier A, Lira SA, Sirard JC, van Rooijen N, et al.. Dendritic cells rapidly recruited into epithelial tissues via CCR6/CCL20 are responsible for CD8+ T cell crosspriming in vivo. Immunity 2006; 24:191–201.
50. Luo DF, Bui MM, Muir A, Maclaren NK, Thomson G, She JX. Affected-sib-pair mapping of a novel susceptibility gene to insulin-dependent diabetes mellitus (IDDM8) on chromosome 6q25–q27. Am J Hum Genet 1995; 57:911–919.
51. Davies JL, Cucca F, Goy JV, Atta ZA, Merriman ME, Wilson A, et al.. Saturation multipoint linkage mapping of chromosome 6q in type 1 diabetes. Hum Mol Genet 1996; 5:1071–1074.
52. Owerbach D. Physical and genetic mapping of IDDM8 on chromosome 6q27. Diabetes 2000; 49:508–512.
53. Cox NJ, Wapelhorst B, Morrison VA, Johnson L, Pinchuk L, Spielman RS, et al.. Seven regions of the genome show evidence of linkage to type 1 diabetes in a consensus analysis of 767 multiplex families. Am J Hum Genet 2001; 69:820–830.
54. Myerscough A, John S, Barrett JH, Ollier WE, Worthington J. Linkage of rheumatoid arthritis to insulin-dependent diabetes mellitus loci: evidence supporting a hypothesis for the existence of common autoimmune
susceptibility loci. Arthritis Rheum 2000; 43:2771–2775.
55. Birlea SA, Fain PR, Spritz RA. A Romanian population isolate with high frequency of vitiligo
and associated autoimmune
diseases. Arch Dermatol 2008; 144:310–316.
56. Maier S, Paulsson M, Hartmann U. The widely expressed extracellular matrix protein SMOC-2 promotes keratinocyte attachment and migration. Exp Cell Res 2008; 314:2477–2487.
57. Gauthier Y, Cario Andre M, Lepreux S, Pain C, Taïeb A. Melanocyte detachment after skin friction in non lesional skin of patients with generalized vitiligo
. Br J Dermatol 2003; 148:95–101.
58. Clark AJL, Weber A. Adrenocorticotropin insensitivity syndromes. Endocr Rev 1998; 19:828–843.
59. Zuo L, Weger J, Yang Q, Goldstein AM, Tucker MA, Walker GJ, et al.. Germline mutations in the p16(INK4a) binding domain of CDK4 in familial melanoma. Nat Genet 1996; 12:97–99.
60. Machado Filho CD, Almeida FA, Proto RS, Landman G. Vitiligo
: analysis of grafting versus curettage alone, using melanocyte morphology and reverse transcriptase polymerase chain reaction for tyrosinase mRNA. Sao Paulo Med J 2005; 123:187–191.
61. Kingo K, Philips MA, Aunin E, Luuk H, Karelson M, Rätsep R, et al.. MYG1, novel melanocyte related gene, has elevated expression in vitiligo
. J Dermatol Sci 2006; 44:119–122.
62. Koga M. Vitiligo
: a new classification and therapy. Br J Dermatol 1977; 97:255–261.
63. Koga M, Tango T. Clinical features and course of type A and type B vitiligo
. Br J Dermatol 1988; 118:223–228.
64. Kawamura T, Ikeda S, Hori YKawamura T, Fitzpatrick TB, Seiji M. Pigmentary disorders. Asiatics: biology of normal and abnormal melanocytes 1971. Baltimore: University Park Press; 352.
65. Winkelmann RKZelickson AS. Cutaneous nerves. Ultrastruclure of normal and abnormal skin 1967. Philadelphia: Lea and Febiger; 202.
66. Chouchkov CN. On the fine structure of free nerve endings in human digital skin, oral cavity and rectum. Z Mikrosk Anat Forsch 1972; 86:273–288.
67. Lerner AB. Clinical applications of psoralens and related materials: vitiligo
. J Invest Dermatol 1959; 32:285–310.
68. Costea V. Leukoderma patches in the course of traumatic paralysis of the brachial plexus in a subject with insular cavities. Derm Venerol 1961; 2:161–166.
69. Koplon BS, Shapiro L. Poliosis overlying a neurofibroma. Arch Dermatol 1968; 98:631–633.
70. Zvulunov A, Esterly NB. Neurocutaneous syndromes associated with pigmentary skin lesions. J Am Acad Dermatol 1995; 32:915–935.
71. Mosher DB, Fitzpatrick TB, Hori Y, Ortonne JPFitzpatrick TB, Eisen AZ, Wolff K. Disorders of pigmentation. Dermatology in general medicine 1993. New York: McGraw-Hill; 903–995.
72. Nelhaus G. Acquired unilateral vitiligo
and poliosis of the head and subacute encephalitis with partial recovery. Neurology 1970; 20:965–974.
73. Fahian G. The spread of black pigment of the denervated skin of the guinea-pig. Acta Biol Acad Sci Hung 1951; 4:471–480.
74. Ehinger B, Zellforsch Z. Mikrosk. Anat 1971; 116:157–177.
75. Ehinger B, Falck B, Zellforsch Z. Mikrosk. Anat 1970; 105:538–542.
76. Laties AM. Specific neurohistology comes of age: a look back and a look forward the Jonas S. Friedenwald Memorial Lecture. Invest Ophthalmol Vis Sci 1972; 11:555–584.
77. Mosher DB, Fitzpatrick TB, Ortonne JP, Hori YFitzpatrick TB, Eisen AZ, Wolff K, Freedberg IM, Austen K. Disorders in pigmentation. Dermatology in General Medicine 1987. New York: McGraw-Hill Book Company; 794–876.
78. Morohashi M, Hashimoto K, Goodman TF Jr, Newton DE, Rist T. Ultrastructural studies of vitiligo
, Vogt-Koyanagi syndrome and incontinentia pigmenti achromians. Arch Dermatol 1977; 113:755–766.
79. Shao CC. Changes of the neuroreceptor apparatus of the skin in vitiligo
. Vestn Dermatol Venerol 1959; 33:20–24.
80. Breathnach AS, Bor S, Wyllie LM. Electron microscopy of peripheral nerve terminals and marginal melanocytes in vitiligo
. J Invest Dermatol 1966; 47:125–140.
81. Moore RY, Bloom FE. Central catecholamine neuron systems: anatomy and physiology of the norepinephrine and epinephrine systems. Annu Rev Neurosci 1979; 2:113–168.
82. Oehme P, Hecht K, Piesche L, Hilse H, Poppei M, Morgensterm E, Gores E. Substance-P: new aspect of its modulatory function. Acta Biol Med Ger 1980; 39:469–477.
83. Gieler U, Brosig B, Schneider U, Kupfer J, Niemeier V, Stangier U, Küster W. Vitiligo
– coping behavior. Dermatol Psychosom 2000; 1:6–10.
84. Agarwal G. Vitiligo
: an under-estimated problem. Fam Pract 1998; 15Suppl 1S19–S23.
85. Mattoo SK, Handa S, Kaur I, Gupta N, Malhotra R. Psychiatric morbidity in vitiligo
: prevalence and correlates in India. J Eur Acad Dermatol Venereol 2002; 16:573–578.
86. Elgowieni M, Ramadan I, Molukia T. Vitiligo
: its personality proﬁle. Br J Dermatol 2003; 149:92.
87. Morrone A, Picardo M, de Luca C, Terminali O, Passi S, Ippolito F. Catecholamines and vitiligo
. Pigment Cell Res 1992; 5:65–69.
88. Schallreuter KU, Wood JM, Pittelkow MR, Swanson NN, Steinkraus V. Increased in vitro expression of beta2
-adrenoceptors in differentiating lesional keratinocytes of vitiligo
patients. Arch Dermatol Res 1993; 285:216–220.
89. Schallreuter KU, Lemke KR, Pittelkow MR, Wood JM, Korner C, Malik R. Catecholamines in human keratinocyte differentiation. J Invest Dermatol 1995; 104:953–957.
90. Schallreuter KU. Epidermal adrenergic signal transduction as part of the neuronal network in the human epidermis. J Investig Dermatol Symp Proc 1997; 2:37–40.
91. Farber EM, Rein G, Lanigan SW. Stress and psoriasis: psychoneuroimmunologic mechanisms. Int J Dermatol 1991; 30:8–12.
92. Al’Abadie MSK, Gawkrodger DJ, Senior HJ, Bleehen SS. Neuropathological studies in vitiligo
. Br J Dermatol 1992; 127:26.
93. Lazarova R, Hristakieva E, Lazarov N, Shani J. Vitiligo
-related neuropeptides in nerve fibers of the skin. Arch Physiol Biochem 2000; 108:262–267.
94. Covelli V, Jirillo E. Neuropeptides with immunoregulatory functions: current status of investigations. Funct Neurol 1988; 3:253–261.
95. Chanco-Turner ML, Lerner AB. Physiological changes in vitiligo
. Arch Dermatol 1965; 91:390–396.
96. Sundler F, Håkanson R, Ekblad E, Uddman R, Wahlestedt C. Neuropeptide Y in the peripheral adrenergic and enteric nervous systems. Int Rev Cytol 1986; 102:243–269.
97. Lundberg JM, Terenius L, Hokfelt T, Goldstein M. High levels of neuropeptide Y in peripheral noradrenergic neurons in various mammals including man. Neurosci Lett 1983; 42:167–172.
98. Ribeiro Da Silva A, Kenigsberg RL, Cuello AC. Light and electron microscopic distribution of nerve growth factor receptor-like immunoreactivity in the skin of the rat lower lip. Neuroscience 1991; 43:631–646.
99. Lewin GR, Barde YA. Physiology of the neurotrophins. Annu Rev Neurosci 1996; 19:289–317.
100. Levi Montalcini R, Angeletti PU. Nerve growth factor. Physiol Rev 1968; 48:534–569.
101. Johnson EM Jr, Gorin PD, Brandeis LD, Pearson J. Dorsal root ganglion neurons are destroyed by exposure in utero to maternal antibody to nerve growth factor. Science 1980; 210:916–918.
102. Schallreuter KU, Lemke R, Brandt O, Schwartz R, Westhofen M, Montz R, Berger J. Vitiligo
and other diseases: coexistence or true association? Hamburg study on 321 patients. Dermatology 1994; 188:269–275.
103. Hegedus L, Heidenheim M, Gervil M, Hjalgrim H, Hoier-Madsen M. High frequency of thyroid dysfunction in patients with vitiligo
. Acta Derm Venereol 1994; 74:120–123.
104. Betterle C, Caretto A, De Zio A, Pedini B, Veller-Fornasa C, Cecchetto A, et al.. Incidence and significance of organ-specific autoimmune
disorders (clinical, latent or only autoantibodies) in patients with vitiligo
. Dermatologica 1985; 171:419–423.
105. Kao CH, Yu HS. Comparison of the effect of 8-methoxypsoralen (8-MOP) plus UVA (PUVA) on human melanocytes in vitiligo
vulgaris and in vitro. J Invest Dermatol 1992; 98:734–740.
106. Duthie MS, Kimber I, Norval M. The effects of ultraviolet radiation on the human immune system. Br J Dermatol 1999; 140:995–1009.
107. Abdel Naser MB, Hann SK, Bystryn JC. Oral psoralen with UV-A therapy releases circulating growth factor(s) that stimulates cell proliferation. Arch Dermatol 1997; 133:1530–1533.
108. Fitzpatrick TB. Mechanisms of phototherapy of vitiligo
. Arch Dermatol 1997; 133:1591–1592.
109. Viac J, Goujon C, Misery L, Staniek V, Faure M, Schmitt D, Claudy A. Effect of UVB 311 nm irradiation on normal human skin. Photodermatol Photoimmunol Photomed 1997; 13:103–108.
110. Krutmann J, Morita A. Mechanisms of ultraviolet (UV) B and UVA phototherapy. J Investig Dermatol Symp Proc 1999; 4:70–72.
111. Halder RM, Young CM. New and emerging therapies for vitiligo
. Dermatol Clin 2000; 18:79–89.
112. Simsek H, Savas C, Akkiz H, Telatar H. Interferon-induced vitiligo
in a patient with chronic viral hepatitis C infection. Dermatology 1996; 193:65–66.
113. Nouri K, Busso M, Machler BC. Vitiligo
associated with alpha-interferon in a patient with chronic active hepatitis C. Cutis 1997; 60:289–290.
114. Neumeister P, Strunk D, Apfelbeck U, Sill H, Linkesch W. Adoptive transfer of vitiligo
after allogeneic bone marrow transplantation for non-Hodgkin’s lymphoma. Lancet 2000; 355:1334–1335.
115. Alajlan A, Alfadley A, Pedersen KT. Transfer of vitiligo
after allogeneic bone marrow transplantation. J Am Acad Dermatol 2002; 46:606–610.
116. Au WY, Yeung CK, Chan HH, Lie AK. Generalized vitiligo
after lymphocyte infusion for relapsed leukaemia. Br J Dermatol 2001; 145:1015–1017.
117. Bouloc A, Grange F, Delfau Larue MH, Dieng MT, Tortel MC, Avril MF, et al.. Leucoderma associated with flares of erythrodermic cutaneous T-cell lymphomas: four cases. The French Study Group of Cutaneous Lymphomas. Br J Dermatol 2000; 143:832–836.
118. Harning R, Cui J, Bystryn JC. Relation between the incidence and level of pigment cell antibodies and disease activity in vitiligo
. J Invest Dermatol 1991; 97:1078–1080.
119. Bystryn JC. Serum antibodies in vitiligo
patients. Clin Dermatol 1989; 7:136–145.
120. Naughton GK, Eisinger M, Bystryn JC. Detection of antibodies to melanocytes in vitiligo
by specific immunoprecipitation. J Invest Dermatol 1983; 81:540–542.
121. Hertz KC, Gazze LA, Kirkpatrick CH, Katz SI. Autoimmune vitiligo
. Detection of antibodies to melanin-producing cells. N Engl J Med 1977; 297:634–637.
122. Naughton GK, Reggiardo D, Bystryn JC. Correlation between vitiligo
antibodies and extent of depigmentation in vitiligo
. J Am Acad Dermatol 1986; 155 I978–981.
123. Yu HS, Kao CH, Yu CL. Coexistence and relationship of antikeratinocyte and antimelanocyte antibodies in patients with non-segmental-type vitiligo
. J Invest Dermatol 1993; 100:823–828.
124. Xie P, Geoghegan WD, Jordan RE. Vitiligo
autoantibodies. Studies of subclass distribution and complement activation. J Invest Dermatol 1991; 96:627.
125. Uda H, Takei M, Mishima Y. Immunopathology of vitiligo
vulgaris, Sutton’s leukoderma and melanoma-associated vitiligo
in relation to steroid effects. II. The IgG and C3 deposits in the skin. J Cutan Pathol 1984; 11:114–124.
126. Aronson PJ, Hashimoto K. Association of IgA anti-melanoma antibodies in the sera of vitiligo
patients with active disease. J Invest Dermatol 1987; 88:475.
127. Hann SK, Park YK, Chung KY, Kim YI, Im S, Won JH. Peripheral blood lymphocyte imbalance in Koreans with active vitiligo
. Int J Dermatol 1993; 32:286–289.
128. Cui J, Harning R, Henn M, Bystryn JC. Identification of pigment cell antigens defined by vitiligo
antibodies. J Invest Dermatol 1992; 98:162–165.
129. Song Y-H, Connor E, Li Y, Zorovich B, Balducci P, Maclaren N. The role of tyrosinase in autoimmune vitiligo
. Lancet 1994; 344:1049–1052.
130. Baharav E, Merimski O, Shoenfeld Y, Zigelman R, Gilbrud B, Yecheskel G, et al.. Tyrosinase as an autoantigen in patients with vitiligo
. Clin Exp Immunol 1996; 105:84–88.
131. Okamoto T, Irie RF, Fujii S, Huang SKS, Nizze AJ, Morton DL, Hoon DSB. Anti-tyrosinase-related protein-2 immune response in vitiligo
patients and melanoma patients receiving active-specific immunotherapy. J Invest Dermatol 1998; 111:1034–1039.
132. Kemp EH, Waterman EA, Gawkrodger DJ, Watson PF, Weetman AP. Autoantibodies to tyrosinase-related protein-1 detected in the sera of vitiligo
patients using a quantitative radiobinding assay. Br J Dermatol 1998; 139:798–805.
133. Kemp EH, Gawkrodger DJ, Watson PF, Weetman AP. Autoantibodies to human melanocyte-specific protein Pmel17 in the sera of vitiligo
patients: a sensitive and quantitative radioimmunoassay (RIA). Clin Exp Immunol 1998; 114:333–338.
134. Xie Z, Chen D, Jiao D, Bystryn JC. Vitiligo
antibodies are not directed to tyrosinase. Arch Dermatol 1999; 135:417–422.
135. Kemp EH, Gawkrodger DJ, Watson PF, Weetman AP. Immunoprecipitation of melanogenic enzyme autoantigens with vitiligo
sera: evidence for cross-reactive autoantibodies to tyrosinase and tyrosinase-related protein-2 (TRP-2). Clin Exp Immunol 1997; 109:495–500.
136. Kemp EH, Gavalas NG, Gawkrodger DJ, Weetman AP. Autoantibody responses to melanocytes in the depigmenting skin disease vitiligo
. Autoimmun Rev 2007; 6:138–142.
137. Helen Kemp E, Waterman EA, Hawes BE, O’Neill K, Gottumukkala RVSRK, Gawkrodger DJ. The melanin-concentrating hormone receptor 1, a novel target of autoantibody responses in vitiligo
. J Clin Invest 2002; 109:923–930.
138. Cui J, Arita Y, Bystryn JC. Cytolytic antibodies to melanocytes in vitiligo
. J Invest Dermatol 1993; 100:812–815.
139. Park YK, Kim NS, Hann SK, Im S. Identification of autoantibody to melanocytes and characterization of vitiligo
antigen in vitiligo
patients. J Dermatol Sci 1996; 11:111–120.
140. Moellman GE, Krass P, Halaban R, Kuklinska E, Lerner AB. On the subject of serum antibodies to melanocytes in vitiligo
. J Invest Dermatol 1985; 84:333.
141. Norris DA, Kissinger RM, Naughton GM, Bystryn JC. Evidence for immunologic mechanisms in human vitiligo
: patients’ sera induce damage to human melanocytes in vitro by complement-mediated damage and antibody-dependent cellular cytotoxicity. J Invest Dermatol 1988; 90:783–789.
142. Gilhar A, Zelickson B, Ulman Y, Etzioni A. In vivo destruction of melanocytes by the IgG fraction of serum from patients with vitiligo
. J Invest Dermatol 1995; 105:683–686.
143. Fishman P, Azizi E, Shoenfeld Y, Sredni B, Yecheskel G, Ferrone S, et al.. Vitiligo
autoantibodies are effective against melanoma. Cancer 1993; 72:2365–2369.
144. Gottumukkala RV, Gavalas NG, Akhtar S, Metcalfe RA, Gawkrodger DJ, Haycock JW, et al.. Function-blocking autoantibodies to the melanin-concentrating hormone receptor in vitiligo
patients. Lab Invest 2006; 86:781–789.
145. Wańkowicz-Kalińska A, van den Wijngaard RM, Tigges BJ, Westerhof W, Ogg GS, Cerundolo V, et al.. Immunopolarization of CD4+
T cells to type-1-like is associated with melanocyte loss in human vitiligo
. Lab Invest 2003; 83:683–695.
146. Abdel-Naser MB, Krüger-Krasagakes S, Krasagakis K, Gollnick H, Abdel Fattah A, Orfanos CE. Further evidence for involvement of both cell mediated and humoral immunity in generalized vitiligo
. Pigment Cell Res 1994; 7:1–8.
147. Al Badri AMT, Foulis AK, Todd PM, Garioch JJ, Gudgeon JE, Stewart DG, et al.. Abnormal expression of MHC class II and ICAM-1 by melanocytes in vitiligo
. J Pathol 1993; 169:203–206.
148. Okada T, Sakamoto T, Ishibashi T, Inomata H. Vitiligo
in Vogt–Koyanagi–Harada disease: immunohistological analysis of inﬂammatory site. Graefes Arch Clin Exp Ophthalmol 1996; 234:359–363.
149. Horn TD, Abanmi A. Analysis of the lymphocytic infiltrate in a case of vitiligo
. Am J Dermatopathol 1997; 19:400–402.
150. Honda Y, Okubo Y, Koga M. Relationship between levels of soluble interleukin-2 receptors and the types and activity of vitiligo
. J Dermatol 1997; 24:561–563.
151. Yeo UC, Yang YS, Park KB, Sung HT, Jung SY, Lee ES, Shin MH. Serum concentration of the soluble interleukin-2 receptor in vitiligo
patients. J Dermatol Sci 1999; 19:182–188.
152. Caixia T, Hongwen F, Xiran L. Levels of soluble interleukin-2 receptor in the sera and skin tissue fluids of patients with vitiligo
. J Dermatol Sci 1999; 21:59–62.
153. Rosenberg SA. Cancer vaccines based on the identification of genes encoding cancer regression antigens. Immunol Today 1997; 18:175–182.
154. Shi YL, Li K, Hamzavi I, Lim HW, Zhou L, Mi QS. Elevated circulating soluble interleukin-2 receptor in patients with non-segmental vitiligo
in North American. J Dermatol Sci 2013; 71:212–214.
155. Harris JE, Harris TH, Weninger W, Wherry EJ, Hunter CA, Turka LA. A mouse model of vitiligo
with focused epidermal depigmentation requires IFN-γ for autoreactive CD8+
T-cell accumulation in the skin. J Invest Dermatol 2012; 132:1869–1876.
156. Ogg GS, Dunbar PR, Romero P, Chen JL, Cerundolo V. High frequency of skin-homing melanocyte-specific cytotoxic T lymphocytes in autoimmune vitiligo
. J Exp Med 1998; 188:1203–1208.
157. Lang KS, Caroli CC, Muhm A, Wernet D, Moris A, Schittek B, et al.. HLA-A2 restricted, melanocyte-specific CD8+
T lymphocytes detected in vitiligo
patients are related to disease activity and are predominantly directed against MelanA/MART1. J Invest Dermatol 2001; 116:891–897.
158. Van Den Wijngaard R, Wankowicz-Kalinska A, Le Poole C, Tigges B, Westerhof W, Das P. Local immune response in skin of generalized vitiligo
patients: destruction of melanocytes is associated with the prominent presence of CLA+
T cells at the perilesional site. Lab Invest 2000; 80:1299–1309.
159. Van den Wijngaard R, Wankowicz-Kalinska A, Pals S, Weening J, Das P. Autoimmune
melanocyte destruction in vitiligo
. Lab Invest 2001; 81:1061–1067.
160. Das PK, Van Den Wijngaard RMJGJ, Wankowicz-Kalinska A, Le Poole IC. A symbiotic concept of autoimmunity and tumour immunity: lessons from vitiligo
. Trends Immunol 2001; 22:130–136.
161. Savill J. Recognition and phagocytosis of cells undergoing apoptosis. Br Med Bull 1997; 53:491–508.
162. Dimmeler S, Zeiher AM. Nitric oxide and apoptosis: another paradigm for the double-edged role of nitric oxide. Nitric Oxide 1997; 1:275–281.
163. Gogas H, Ioannovich J, Dafni U, Stavropoulou Giokas C, Frangia K, Tsoutsos D, et al.. Prognostic significance of autoimmunity during treatment of melanoma with interferon. N Engl J Med 2006; 354:709–718.
164. Becker JC, Guldberg P, Zeuthen J, Bröcker EB, Straten PT. Accumulation of identical T cells in melanoma and vitiligo
-like leukoderma. J Invest Dermatol 1999; 113:1033–1038.
165. Jäger E, Maeurer M, Höhn H, Karbach J, Jäger D, Zidianakis Z, et al.. Clonal expansion of Melan A-specific cytotoxic T lymphocytes in a melanoma patient responding to continued immunization with melanoma-associated peptides. Int J Cancer 2000; 86:538–547.
166. Rivoltini L, Radrizzani M, Accornero P, Squarcina P, Chiodoni C, Mazzocchi A, et al.. Human melanoma-reactive CD4+
CTL clones resist Fas ligand-induced apoptosis and use Fas/Fas ligand-independent mechanisms for tumor killing. J Immunol 1998; 161:1220–1230.
167. Kao CH, Yu HS. Depletion and repopulation of Langerhans cells in nonsegmental type vitiligo
. J Dermatol 1990; 17:287–296.
168. Jimbow K, Quevedo WC Jr, Fitzpatrick TB, Szabo G. Some aspects of melanin biology: 1950–1975. J Invest Dermatol 1976; 67:72–89.
169. Moretti S, Spallanzani A, Amato L, Hautmann G, Gallerani I, Fabiani M, Fabbri P. New insights into the pathogenesis
: imbalance of epidermal cytokines at sites of lesions. Pigment Cell Res 2002; 15:87–92.
170. Imokawa G, Yada Y, Kimura M, Morisaki N. Granulocyte/macrophage colony-stimulating factor is an intrinsic keratinocyte-derived growth factor for human melanocytes in UVA-induced melanosis. Biochem J 1996; 313:625–631.
171. Puri N, Van Der Weel MB, De Wit FS, Asghar SS, Das PK, Ramaiah A, Westerhof W. Basic fibroblast growth factor promotes melanin synthesis by melanocytes. Arch Dermatol Res 1996; 288:633–635.
172. Grichnik JM, Burch JA, Burchette J, Shea CR. The SCF/KIT pathway plays a critical role in the control of normal human melanocyte homeostasis. J Invest Dermatol 1998; 111:233–238.
173. Swope VB, Abdel Malek Z, Kassem LM, Nordlund JJ. Interleukins 1α and 6 and tumor necrosis factor-α are paracrine inhibitors of human melanocyte proliferation and melanogenesis. J Invest Dermatol 1991; 96:180–185.
174. Martínez-Esparza M, Jiménez-Cervantes C, Beermann F, Aparicio P, Lozano JA, García-Borrón JC. Transforming growth factor-β1 inhibits basal melanogenesis in B16/F10 mouse melanoma cells by increasing the rate of degradation of tyrosinase and tyrosinase-related protein-1. J Biol Chem 1997; 272:3967–3972.
175. Santos Rosa M, Bienvenu J, Whiher JBurtis CA, Ashwood ER. Cytokines. Tietz textbook of clinical chemistry 1999. Philadelphia: Saunders; 587–592.
176. Grimes PE, Soriano T, Dytoc MT. Topical tacrolimus for repigmentation of vitiligo
. J Am Acad Dermatol 2002; 47:789–791.
177. Tanghetti EA. Tacrolimus ointment 0.1% produces repigmentation in patients with vitiligo
: results of a prospective patient series. Cutis 2003; 71:158–162.
178. Lepe V, Moncada B, Castanedo-Cazares JP, Torres-Alvarez MB, Ortiz CA, Torres-Rubalcava AB. A double-blind randomized trial of 0.1% tacrolimus vs 0.05% clobetasol for the treatment of childhood vitiligo
. Arch Dermatol 2003; 139:581–585.
179. Castanedo Cazares JP, Lepe V, Moncada B. Repigmentation of chronic vitiligo
lesions by following tacrolimus plus ultraviolet-B-narrow-band. Photodermatol Photoimmunol Photomed 2003; 19:35–36.
180. Grimes PE, Morris R, Avaniss Aghajani E, Soriano T, Meraz M, Metzger A. Topical tacrolimus therapy for vitiligo
: therapeutic responses and skin messenger RNA expression of proinflammatory cytokines. J Am Acad Dermatol 2004; 51:52–61.
181. Latha B, Babu M. The involvement of free radicals in burn injury: a review. Burns 2001; 27:309–317.
182. Yildirim M, Baysal V, Inaloz HS, Can M. The role of oxidants and antioxidants in generalized vitiligo
at tissue level. J Eur Acad Dermatol Venereol 2004; 18:683–686.
183. Ross ACShils ME, Olson JA, Shike M, Ross AC. Vitamin A. Modern nutrition in health and disease 1999: 9th ed.. Baltimore: Lippincott Williams & Wilkins; 305–327.
184. Ines D, Sonia B, Riadh BM, Amel EG, Slaheddine M, Hamida T, et al.. A comparative study of oxidant-antioxidant status in stable and active vitiligo
patients. Arch Dermatol Res 2006; 298:147–152.
185. Yildirim M, Baysal V, Inaloz HS, Can M. The role of oxidants and antioxidants in generalized vitiligo
in tissue. J Eur Acad Dermatol Venereol 2004; 18:683–686.
186. Khan R, Satyam A, Gupta S, Sharma VK, Sharma A. Circulatory levels of antioxidants and lipid peroxidation in Indian patients with generalized and localized vitiligo
. Arch Dermatol Res 2009; 301:731–737.
187. Casp CB, She JX, McCormack WT. Genetic association of the catalase gene (CAT) with vitiligo
susceptibility. Pigment Cell Res 2002; 15:62–66.
188. Dell’Anna ML, Maresca V, Briganti S, Camera E, Falchi M, Picardo M. Mitochondrial impairment in peripheral blood mononuclear cells during the active phase of vitiligo
. J Invest Dermatol 2001; 117:908–913.
189. Nohl H, Hegner D. Do mitochondria produce oxygen radicals in vivo? Eur J Biochem 1978; 82:563–567.
190. Chance B, Sies H, Boveris A. Hydroperoxide metabolism in mammalian organs. Physiol Rev 1979; 59:527–605.
191. Heales SJR, Bolaños JP, Stewart VC, Brookes PS, Land JM, Clark JB. Nitric oxide, mitochondria and neurological disease. Biochim Biophys Acta 1999; 1410:215–228.
192. Schallreuter KU, Wood JM, Pittelkow MR, Gütlich M, Lemke KR, Rödl W, et al.. Regulation of melanin biosynthesis in the human epidermis by tetrahydrobiopterin. Science 1994; 263:1444–1446.
193. Dell’Anna ML, Urbanelli S, Mastrofrancesco A, Camera E, Iacovelli P, Leone G, et al.. Alterations of mitochondria in peripheral blood mononuclear cells of vitiligo
patients. Pigment Cell Res 2003; 16:553–559.
194. Dell’Anna ML, Ottaviani M, Albanesi V, Vidolin AP, Leone G, Ferraro C, et al.. Membrane lipid alterations as a possible basis for melanocyte degeneration in vitiligo
. J Invest Dermatol 2007; 127:1226–1233.
195. Schallreuter KU, Gibbons NCJ, Zothner C, Elwary SM, Rokos H, Wood JM. Butyrylcholinesterase is present in the human epidermis and is regulated by H2
: more evidence for oxidative stress in vitiligo
. Biochem Biophys Res Commun 2006; 349:931–938.
196. Gohil VM, Hayes P, Matsuyama S, Schägger H, Schlame M, Greenberg ML. Cardiolipin biosynthesis and mitochondrial respiratory chain function are interdependent. J Biol Chem 2004; 279:42612–42618.
197. Haines TH, Dencher NA. Cardiolipin: a proton trap for oxidative phosphorylation. FEBS Lett 2002; 528:35–39.
198. Xu Y, Sutachan JJ, Plesken H, Kelley RI, Schlame M. Characterization of lymphoblast mitochondria from patients with Barth syndrome. Lab Invest 2005; 85:823–830.
199. Schallreuter KU, Moore J, Wood JM, Beazley WD, Peters EMJ, Marles LK, et al.. Epidermal H2
accumulation alters tetrahydrobiopterin (6BH4
) recycling in vitiligo
: Identification of a general mechanism in regulation of all 6BH4
-dependent processes? J Invest Dermatol 2001; 116:167–174.
200. Davis MD, Ribeiro P, Tipper J, Kaufman S. ‘7-Tetrahydrobiopterin,’ a naturally occurring analogue of tetrahydrobiopterin, is a cofactor for and a potential inhibitor of the aromatic amino acid hydroxylases. Proc Natl Acad Sci USA 1992; 89:10109–10113.
201. Kowlessur D, Citron BA, Kaufman S. Recombinant human phenylalanine hydroxylase: novel regulatory and structural properties. Arch Biochem Biophys 1996; 333:85–95.
202. Hasse S, Gibbons NCJ, Rokos H, Marles LK, Schallreuter KU. Perturbed 6-tetrahydrobiopterin recycling via decreased dihydropteridine reductase in vitiligo
: more evidence for H2
stress. J Invest Dermatol 2004; 122:307–313.
203. Schallreuter KU, Wood JM, Ziegler I, Lemke KR, Pittelkow MR, Lindsey NJ, Gutlich M. Defective tetrahydrobiopterin and catecholamine biosynthesis in the depigmentation disorder vitiligo
. Biochim Biophys Acta 1994; 1226:181–192.
204. Elwary SM, Headley K, Schallreuter KU. Calcium homeostasis influences epidermal sweating in patients with vitiligo
. Br J Dermatol 1997; 137:81–85.
205. Iyengar B. Modulation of melanocytic activity by acetylcholine. Acta Anat (Basel) 1989; 136:139–141.
206. Rakonczay Z, Brimijoin S. Biochemistry and pathophysiology of the molecular forms of cholinesterases. Subcell Biochem 1988; 12:335–378.
207. Schallreuter KU, Elwary SMA, Gibbons NCJ, Rokos H, Wood JM. Activation/deactivation of acetylcholinesterase by H2
: more evidence for oxidative stress in vitiligo
. Biochem Biophys Res Commun 2004; 315:502–508.
208. Shalbaf M, Gibbons NC, Wood JM, Maitland DJ, Rokos H, Elwary SM, et al.. Presence of epidermal allantoin further supports oxidative stress in vitiligo
. Exp Dermatol 2008; 17:761–770.
209. Darier J.Vitiligo
. In: La pratiquedermatologique.
4 Paris: Masson; 1904. pp. 846–858.
210. Gauthier Y, Andre MC, Taïeb A. A critical appraisal of vitiligo
etiologic theories. Is melanocyte loss a melanocytorrhagy
? Pigment Cell Res 2003; 16:322–332.
211. Mottaz JH, Thorne EG, Zelickson AS. Response of the epidermal melanocyte to minor trauma. Arch Dermatol 1971; 104:611–618.
212. Warfvinge K, Agdell J, Andersson L, Andersson A. Attachment and detachment of human epidermal melanocytes. Acta Derm Venereol 1990; 70:189–193.
213. Hunter JAA, McVittie E, Comaish JS. Light and electron microscopic studies of physical injury to the skin. II. Friction. Br J Dermatol 1974; 90:491–499.
214. Le Poole IC, Van Den Wijngaard RMJGJ, Westerhof W, Das PK. Tenascin is overexpressed in vitiligo
lesional skin and inhibits melanocyte adhesion. Br J Dermatol 1997; 137:171–178.
215. Jimbow K, Chen H, Park JS, Thomas PD. Increased sensitivity of melanocytes to oxidative stress and abnormal expression of tyrosinase-related protein in vitiligo
. Br J Dermatol 2001; 144:55–65.
216. Schallreuter KU, Wood JM, Lemke KR, Levenig C. Treatment of vitiligo
with a topical application of pseudocatalase and calcium in combination with short-term UVB exposure: a case study on 33 patients. Dermatology 1995; 190:223–229.