T Cell–Independent Mechanisms Associated with Neutrophil Extracellular Trap Formation and Selective Autophagy in IL-17A–Mediated Epidermal Hyperplasia

Interleukin-23 and IL-17A have been implicated with epidermal hyperplasia. Although IL-23 partly regulates the expansion of IL-17A–producing conventional and nonconventional T cells (Th17 and γδT cells, respectively), IL-23 can induce epidermal hyperplasia independently of IL-17A via TNF- and IL-20R2–dependent mechanisms (1). Interestingly, the IL-23–IL-17 axis does not only regulate T cell differentiation but also neutrophil homeostasis and migration (2, 3). In agreement with these observations, we have also found that gene transfer of either IL-23 and IL-17, which is associated with epidermal hyperplasia, also expands neutrophil populations in vivo (4, 5). In keeping with a role of neutrophils in epidermal hyperplasia, others have shown that topical application of a Toll receptor agonist, imiquimod (IMQ), which stimulates neutrophil function, also provokes epidermal hyperplasia in mice (6, 7). Others demonstrated that, in the absence of IL-17RA signaling, IMQ-mediated psoriatic-like skin inflammation is partially reduced, indicating a strong association between activation of neutrophils via IL-17 and IMQ and skin inflammation (8).

Although the contribution of Th17 and γδT cells in epidermal hyperplasia is well documented (9–12), the contribution of activated neutrophils in the observed pathologies is largely unexplored. To investigate the role of activated neutrophils in epidermal hyperplasia, we performed IL-17 gene transfer to induce neutrophilia and activated neutrophils with IMQ in transgenic mice deficient of Th17 or γδT cells. Our studies uncover T cell–independent mechanisms and a direct role of neutrophils in disease initiation and progression.

A direct role for neutrophils in epidermal hyperplasia is supported by the distinct presence of neutrophil exudates (Munro’s microabscesses) in the stratum corneum of the epidermis in psoriasis patients (13, 14). This is also accompanied by an elevated expression of neutrophil biomarkers associated with neutrophil migration including CXCL1, CXCL8, leukotriene B4 (LTB4) and their receptors, CXCR2 and LTB4R1, and neutrophil-derived enzymes including myeloperoxidase (MPO), serpin peptidase inhibitor clade B member 1 (SERPINB1), and cathepsin G and neutrophil elastase (NE) in psoriasis patients (15–18). Lysosomal proteins, NE, MPO, and SERPINB1, support degradation and elimination of neutrophil phagocytic content through autophagy, a process that requires the assembly of an autophagosome regulated by Wdfy3, the master regulator of macroselective autophagy (19, 20). Wdfy3 contains a BEACH domain, originally named after the Chediak–Higashi syndrome, a disorder in humans that leads to neutropenia and defects in lysosomal trafficking, resulting in immunodeficiency (20, 21). In keeping with the putative role of Wdfy3 in neutrophil lysosomal trafficking and inflammation, a recent report demonstrated the requirement of autophagic pathways in neutrophil extracellular trap formation (NETosis) (22), a critical function of neutrophils associated with epidermal hyperplasia and with the secretion of IL-17A (23).

Because NE acts within phagolysosomes to digest phagocytized products, and Wdfy3 is required for the recognition and targeting of the various autophagic cargo for degradation, we primarily tested the hypothesis that NETosis and selective autophagy are critical for epidermal hyperplasia. We validated our observations using NE-deficient mice, which are defective in NE, a requirement for NETosis (24), and conditional Wdfy3-deficient mice that are required for selective autophagy. We further confirmed our in vivo observations with PMA in vitro studies using purified neutrophils that activate NADPH (NOX2)-mediated reactive oxygen species (ROS) production, required for both autophagy and NETosis (22, 25, 26).

Collectively, our data demonstrate that NETosis and selective autophagy may play a role in IL-17A–mediated epidermal hyperplasia and constitute a possible new mechanism, which is independent of IL-23R⁺ αβ and γδ T cells.

C57BL/6, Elane^−/−(NE)^−/−, Tcrδ^−/−, and LysM-Cre mice were purchased from Jackson Laboratories (Sacramento, CA). IL-23R^GFP reporter mice crossed on RAG1 background and the Wdfy3 floxed mice were previously described (27–29). All mice were used between 8 and 12 wk of age. The University of California at Davis Institutional Animal Care and Use Committee approved all animal protocols. Aldara cream (5% IMQ) was obtained from 3 M Pharmaceuticals. GFP and mouse IL-17A minicircle DNA constructs were produced as previously described (4). Serum IL-17A was assessed using an ELISA purchased from eBioscience (San Diego, CA). The CXCL1 ELISA kit was purchased from Sigma (St. Louis, MO). The caspase-3 assay was purchased from BioVision. For the NE assay, CD11b⁺Gr-1^hi or Ly6G⁺ sorted cells were assessed for the presence of NET-associated elastase according to the manufacturer’s instructions (Cayman Chemical, Ann Arbor, MI). In brief, PMA (20, 50 nM) was used to activate cells for NETosis for 1–4 h. DMSO was used as a control for the PMA, washing away unbound NE, followed by a digest of NET DNA by S7 nuclease. The supernatant is taken and later combined with an elastase substrate, and the absorbance is read at 405 nm. ROS was assessed using DHR123 (Life Technologies) before being analyzed by a microplate reader. Cells were incubated with diphenyleneiodonium (NADPH oxidase inhibitor) for 1 h (20 μM) before activating the cells for NETosis. All protocols were followed per manufacturer’s instructions.

CD11b⁺Gr-1^hi cells were sorted from the bone marrow of eight mice per group: WT, LysM^Cre, and Wdfy3-LysM^Cre. The transferred blots were blocked with phosphoblocking buffer (Millipore) for 1 h at room temperature. The Abs used were Wdfy3 at 1:1000 (Novus), β-actin at 1:1000 (Cell Signaling), and anti-rabbit IgG HRP at 1:1000 (Cell Signaling). Immunoreactive bands were analyzed by LiCOR using appropriate secondary Abs.

Eight micrograms of IL-17A of GFP control MC DNA was injected hydrodynamically via retro-orbital delivery. Two days after GFP or IL-17A gene transfer, mouse dorsal fur was shaved and treated with Veet hair removal cream, and 20 mg of IMQ was applied to the mouse dorsal skin and assessed the following day. In vivo and ex vivo imaging were performed using a Maestro 2 Cri imager.

Mouse skins were fixed in 10% formalin buffered in PBS and paraffin embedded for sectioning (6 μm). Tissue sections were stained with H&E Y (Sigma). Tissue sections were assessed using a FluoView FV1000 Confocal Microscope. A licensed dermatologist who was blinded to the experimental conditions assessed epidermal thickening. Epidermal thickness (in micrometers) was determined by measuring the interfollicular epidermal area including or excluding parakeratotic areas and corresponding length on H&E-stained longitudinal paraffin sections from mouse dorsal skin. Analysis and quantification were performed on the Olympus software and in Photoshop CS3 (Adobe).

Mice were sacrificed and bone marrow extracts, spleens, and skins were collected. Spleens were injected with collagenase D (Sigma) in media (anti-MEM + 5% FBS + penicillin/streptomycin). Total skin was minced using scissors and incubated with digestion buffer (media + 1 U/ml of Dispase) for 2 h at 37°C, followed by addition of collagenase D for 30 min. Digestion was stopped by adding 10 mM of EDTA. All tissues were passed through 70-μm cell strainers (BD). Cells were treated with ammonium chloride buffer (150 mM of NH₄Cl, 10 mM of KHCO₃, and 100 μM of EDTA) to lyse erythrocytes. Cells were pretreated with anti-CD16/32 mAb for 10 min (to block nonspecific binding; BD Biosciences). The cells were stained with anti-CD45 (30-F11, allophycocyanin), anti-CD11b (M1/70, PB), Annexin V (allophycocyanin), 7-aminoactinomycin D, anti–Gr-1 (RB6-8C5, PeCy7), anti-Ly6G (1A8, FITC, allophycocyanin, or Pacific Blue; 1A8), and anti-CXCR2 (TG11/CXCR2, Alexa 647), and isotype controls were all obtained from BioLegend. Anti–IL-17RA (PAJ-17R, PE) was purchased from eBioscience. AccuCheck counting beads (Life Technologies) were used to determine absolute cell number per square centimeter based on the manufacturer’s protocol. Cells were analyzed on a BD FACSAria flow cytometer (BD Biosciences). The data were analyzed using FlowJo software (Tree Star, Ashland, OR).

Histology sections (6 μm) of each paraffin block were stained and used for immunofluorescence microscopy. Sections were fixed in 4% paraformaldehyde and blocked for 1 h in blocking buffer (1% Triton X, 10% donkey serum in PBS or 2% BSA), then immunostained with DAPI and conjugated Ab against Ly6G (1A8, FITC), NE (Abcam), Wdfy3 (Novus), and donkey anti-goat Alexa 594 secondary and isotype controls (FITC, IgG2a; Alexa Fluor 594 IgG1). CD11b⁺Gr-1^hi or Ly6G⁺ cells were sorted from the spleen or bone marrow and incubated in RPMI 1640, 1% penicillin/streptomycin, 1% BSA, and allowed to adhere to 0.001% poly-l-lysine–coated slides (Thermo Fisher Scientific, Waltham, MA) for 1 h at 37°C. Neutrophils for NE staining were treated with DMSO or 50 nM PMA for 2 or 4 h. The slides were visualized using a confocal microscope (Nikon A1). Fluorescence quantification for NE was performed using ImageJ (National Institutes of Health, Bethesda, MD) to calculate the corrected total cell fluorescence: CTCF = integrated cell density − (area of selected cell × mean fluorescence of background).

Skin tissue was subject to RNA isolation using the RNeasy kit (Qiagen) including a DNase I digest step. Content and purity of RNA were controlled with a Nanodrop spectrophotometer (Thermo Fisher Scientific). cDNA synthesis was performed using the Omniscript reverse transcription kit (Life Technologies). Quantitative real-time PCR for the candidate genes was performed using SYBR green chemistry (Agilent), and values were normalized to Gapdh. The cycling threshold upper limit was fixed to 40 cycles. Cycling parameters were 95°C (20 s) and 60°C (45 s) in a Mx3005P qPCR machine (Stratagene).

Total RNA was isolated from mouse dorsal skins after GFP or IL-17A gene transfer from involved and noninvolved skin at day 3, pooled from six mice per group, using a RNeasy kit (Qiagen). Biotinylated cDNA was synthesized using the Ovation RNA Amplification System V2 (Nugen) and FL-Ovation cDNA Biotin Module V2 (Nugen), then hybridized with Mouse Genome 430 2.0 Arrays (Ilumina). Data are deposited in National Center for Biotechnology Information's Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE86999).

Neutrophils were depleted using two strategies. The first strategy used 500 μg of anti-Ly6G (1A8) Abs or isotype (IgG2b; LFT-2) as a control 2 d before gene transfer and upon gene transfer. Neutrophil depletion was confirmed on day 2 before gene transfer by staining isolated splenocytes with anti–Gr-1 and anti-CD11b Abs and analyzing the cells by flow cytometry. Second, neutrophil depletion was achieved by using cyclophosphamide as previously described (30). PBS injections served as controls for the cyclophosphamide injections. Neutrophil depletion was confirmed on day 6 before gene transfer by staining isolated splenocytes with anti–Gr-1 and anti-CD11b Abs and analyzing the cells by flow cytometry.

Splenocytes or bone marrow aspirates from GFP or IL-17A MC-injected mice treated with IMQ on day 3 were sorted for CD11b⁺Gr-1^hi or CD11b⁺Gr-1^lo. Purity of the CD11b⁺Gr-1^hi cells ranged from 90 to 95%. The cells were labeled with VivoTrack 680 (Perkin Elmer) for 15 min at 37°C, washed three times with PBS, and injected i.v. into naive recipients (5 × 10⁶ cells per mouse), which were treated the same day with 20 mg of IMQ to shaved dorsal skin. Whole-body in vivo imaging was performed after 24 h using the Maestro 2 Cri imager. Optical imaging for VivoTrack 680 was performed using a red filter set (680–950 nm, long-pass filter). A camera was used to acquire captured images at constant exposure times.

Statistical analysis and graphical representations were done using Prism5 software (GraphPad Software). Significant differences in gene expression, imaging, flow cytometry, and epidermal thickening were assessed using a Mann–Whitney U test or Kruskal–Wallis test followed by post hoc Dunn’s test where appropriate.

To determine the role of IL-17A in vivo, we used a gene transfer approach as previously described with minor modifications (4). IL-17A was elevated in the serum within 24 h and further expanded the CD11b⁺Gr-1^hi population to be studied (Supplemental Fig. 1A–F). Phenotypic analysis of the expanded CD11b⁺Gr-1^hi population further confirmed the expression of MPO, as well as CXCR2⁺ and IL-17R⁺, similar to the CD11b⁺Gr-1^hi cells after GFP gene transfer (Supplemental Fig. 1E, 1F). Moreover, the expansion of the neutrophils in the periphery after IL-17A gene transfer correlated with an increase in the serum CXCR2 ligand, CXCL1 (Supplemental Fig. 1G). To determine the cellular and molecular mechanisms of epidermal hyperplasia, we induced the IL-17A–expanded neutrophil population in mouse dorsal skin with IMQ (a known activator of neutrophils) (Fig. 1A). We performed IMQ dose–response and time-course experiments after IL-17A gene transfer at low (0, 10, 20 mg) and high (62.5 mg) IMQ concentrations over 5 d to determine a suboptimal dose that could be exacerbated by IL-17A (20 mg) by histology and gene expression analysis (Supplemental Fig. 2A, 2B). We found that IL-17A gene transfer exacerbated IMQ-induced epidermal hyperplasia in WT mice on day 4 after gene transfer, compared with IMQ-induced GFP MC-injected controls (Fig. 1B, 1C). Histological analysis of dorsal skin from IL-17A MC-injected mice revealed a mixed cell infiltrate containing neutrophils in the upper epidermis (Munro’s microabscesses), along with epidermal hyperplasia (Fig. 1B, 1C). Immunofluorescence imaging of dorsal skin after IL-17A gene transfer confirmed that neutrophils accumulate in the skin (Fig. 1D; Supplemental Fig. 2C–E). Consistent with the histological phenotype, we found an increase in CD11b⁺Gr-1^hi neutrophils by flow cytometric analysis in the skin (Fig. 1E, 1F). The increase in CD11b⁺Gr-1^hi neutrophils correlated with an increase in genes associated with keratinocyte proliferation and skin inflammation including K16, S100a8, Serpinb1, Cxcl1, Cxcr2, and Ltb4r1 (Fig. 1G).

To explore whether the effects of IL-17A were independent of IL-23R⁺ T cells and γδT cells, we performed IL-17A gene transfer in Il23r^−/−Rag1^−/− and Tcrδ^−/− mice. Epidermal hyperplasia was reduced in mice deficient in Il23r^−/−Rag1^−/− and Tcrδ (Fig. 1H). In keeping with these observations, the IL-17A–associated elevation in CD11b⁺Gr-1^hi neutrophils in the skin persisted in Il23r^−/− and Tcrδ^−/− mice (Fig. 1I, 1J). These data further correlated with an elevation of keratinocyte and inflammatory markers in Il23r^−/−Rag1^−/− and Tcrδ^−/− mice (Fig. 1K), demonstrating that IL-17A can act independently of IL-23R and T cell mechanisms to induce skin inflammation.

To demonstrate whether the CD11b⁺Gr-1^hi cell population was responsible for inducing epidermal hyperplasia, we performed a neutrophil depletion (Supplemental Fig. 3). Anti-Ly6G significantly reduced the expansion of CD11b⁺Gr-1^hi cells in the spleen and skin after IL-17A gene transfer compared with isotype and vehicle controls (Supplemental Fig. 3). The reduction of neutrophils corresponded with a reduction in epidermal thickening, epidermal hyperplasia, reduced mixed cell infiltrates, and a lack of Munro’s microabscesses (Fig. 2A, 2B). Moreover, the reduction in neutrophils was associated with a significantly diminished induction of K16, S100a8, Serpinb1, Cxcl1, Cxcr2, and Ltb4r1 in the skin upon IL-17A gene transfer (Fig. 2C). These data indicate that neutrophil depletion suppresses IL-17A–mediated epidermal hyperplasia in vivo.

We next performed an adoptive transfer of CD11b⁺Gr-1^hi cells into naive recipients in the presence of IMQ to evaluate the pathogenicity of neutrophils in the absence of systemic IL-17A expression. We found that CD11b⁺Gr-1^hi, but not CD11b⁺Gr-1^lo, from IL-17A gene transfer mice localized to the skin after application of IMQ as detected by VivoTrack 680 (31) labeling (Fig. 2D, 2E). Histological analysis of mouse dorsal skin showed increased epidermal thickening and evidence of epidermal hyperplasia along with Munro’s microabscesses (Fig. 2E, 2F). Moreover, gene expression of K16, S100a8, Cxcl1, and Ltb4r1 were elevated upon adoptive transfer with IL-17A MC CD11b⁺Gr-1^hi cells compared with the adoptive transfer of IL-17A MC CD11b⁺Gr-1^lo or GFP MC CD11b⁺Gr-1^hi cells (Fig. 2G). Notably, epidermal hyperplasia was not observed in the control experiments involving adoptive transfer of IL-17A MC CD11b⁺Gr-1^lo or GFP MC CD11b⁺Gr-1^hi, in the presence of IMQ.

We next performed in vitro studies on isolated neutrophils to determine whether neutrophil function (phagocytosis, neutrophil activation, or NETosis) is critical to confer pathogenicity in our IL-17A gene transfer model. Neutrophils isolated from IL-17A gene transfer mice treated with IMQ showed no detectable difference in neutrophil phagocytosis (Fig. 3A) or neutrophil activation (Fig. 3B) compared with GFP gene transfer controls. However, immunofluorescence staining of sorted CD11b⁺Gr-1^hi cells from IL-17A gene transfer mice treated with IMQ revealed a marked increase in the expression of NE compared with controls (Fig. 3C), which also correlated with an increase in NE elastase in the presence of PMA (Fig. 3D). These findings were also confirmed in vivo by immunofluorescence imaging of dorsal skins after IL-17A gene transfer and IMQ, which revealed increased NE protein expression compared with controls (Fig. 3E, 3F). To further elucidate the role of NE, we performed IL-17A gene transfer in combination with IMQ in Elane-deficient mice (herein referred to as NE^−/− mice) (Fig. 3G). Histological analysis of dorsal skins from IL-17A MC-injected NE^−/− mice exhibited reduced epidermal thickening and reduced epidermal hyperplasia and Munro’s microabscesses compared with WT controls (Fig. 3H, 3I). The reduction in skin pathology was consistent with a reduction in gene expression of K16 (Fig. 3J). The expansion of neutrophils in the spleen (Fig. 3K, 3L) and the migration of neutrophils to the skin after IL-17A gene transfer and IMQ were not inhibited in NE^−/− mice (Fig. 3M, 3N). These data demonstrate that NE plays a critical role in the pathogenesis of IL-17A–mediated skin pathology.

Because recent evidence has shown that autophagy is critical for NETosis (22), we next sought to investigate whether deletion of the master regulator of selective autophagy, Wdfy3, would affect NETosis. Wdfy3 protein and gene expression were found to be elevated in the inflamed skin (Supplemental Fig. 4A–C). Wdfy3^flox/flox mice were crossed with LysM^Cre to selectively ablate Wdfy3 in myeloid cells to generate Wdfy3^flox/flox-LysM^Cre/+ mice (herein referred to as Wdfy3-LysM^Cre). The deletion efficiency of the LysM^Cre was confirmed in CD11b⁺Gr-1^hi cells, and there was no difference in the total cell number of CD11b⁺Gr-1^hi, CD11b⁺, and CD11c⁺ myeloid cells or any impairment in IL-17A–mediated neutrophil expansion, circulation, and survival, as well as neutrophil priming, activation, or phagocytosis (Supplemental Fig. 4D–H and data not shown). To assess the role of Wdfy3-LysM^Cre sorted neutrophils in NETosis, we stimulated the cells with PMA. In vitro immunofluorescence analysis of Wdfy3-LysM^Cre neutrophils showed that a greater proportion of nuclei were distinct demarcated multilobar rings with uniform DNA staining and had a significant reduction of diffuse nuclei undergoing NETosis, which commonly lacks structure and has a heterogeneous pattern consistent with breakdown of the nuclear envelope, chromatin decondensation, and intracellular chromatin dispersion (Fig. 4A, 4B). Wdfy3-LysM^Cre neutrophils also showed a marked reduction in NET elastase (Fig. 4C) and reduced production of ROS compared with PMA-treated WT or NE^−/−controls (Fig. 4D).

To confirm the requirement of Wdfy3-deficient neutrophils in epidermal hyperplasia, we performed IL-17A gene transfer in Wdfy3-LysM^Cre mice (Fig. 5A). Histological analysis of dorsal skins from IL-17A MC-injected Wdfy3-LysM^Cre mice at day 4 revealed reduced epidermal thickening and absence of epidermal hyperplasia and Munro’s microabscesses in contrast with WT controls (Fig. 5B, 5C). Consistent with the histology findings, we observed a reduction in gene expression of K16, S100a8, Cxcl1, Cxcr2, and Ltb4r1 in IL-17A MC-injected Wdfy3-LysM^Cre dorsal skins compared with IL-17A MC-injected WT mice (Fig. 5D). Adoptive transfer of sorted CD11b⁺Gr-1^hi cells from Wdfy3-LysM^Cre mice into WT mice failed to induce skin pathology (Fig. 5E–G), and this was consistent with a reduction in gene expression of K16, S100a8, Cxcl1, and Ltb4r1 (Fig. 5H). Taken together, these data demonstrate a novel role for Wdfy3 in mediating neutrophil function via NE in skin inflammation.

In our IL-17A gene transfer experiments with γδT cells and Il23r^−/−Rag1^−/− transgenic mice, we clearly demonstrate IL-23R⁺ and T cell–independent mechanisms of IL-17A–induced skin pathology. The induction of epidermal hyperplasia was found to be dependent on CD11b⁺Gr-1^hi neutrophils because neutrophil depletion was able to prevent skin pathology. Our data are in agreement with similar studies, whereby conditional overexpression of IL-17A in keratinocytes caused severe psoriasis-like skin inflammation in mice that was reduced by neutrophil depletion (32). Using the same conditional overexpression of IL-17A in keratinocytes, other studies have also supported the role of IL-17A in neutrophil-associated Munro’s microabscesses through IL-6 (33). These data are also in agreement with our previous findings where gene transfer of IL-17 also elevated serum IL-6 in vivo and was associated with skin inflammation and neutrophil exudates (4). Although it is agreed that there is prominent formation of neutrophil-rich Munro’s microabscesses (a clinical marker with the highest histological occurrence for the diagnosis of psoriasis) (34), little is known about the neutrophil cellular and molecular mechanisms that confer pathogenicity.

Recent reports have highlighted novel roles for the function of neutrophils in destructive diseases that are dependent on NETosis, a unique form of cell death that is characterized by the release of decondensed chromatin and granular contents to the extracellular space (35). Indeed, it has been reported that NETosis is prominent in necrotic areas and the prevention of NET release eliminates tissue damage in liver tissue (36). Specifically, inhibition of NE, an enzyme implicated in the initial decondensation of DNA and the proteolytic degradation of the nuclear envelope, diminished damage to the liver (36). In keeping with these observations, other groups have also shown that NE is critical in the formation of NETs, and mouse peritoneal neutrophils derived from NE-deficient mice lack nuclear decondensation activity in vitro and fail to form NETs (24). We also found that neutrophils isolated from IL-17A gene transfer mice in the presence of IMQ had elevated NE expression, and in vivo this expression was localized within the inflamed skin. Furthermore, because NE^−/− mice were protected from IL-17A–mediated epidermal hyperplasia, our data demonstrate NE to be critical for disease pathogenesis.

In keeping with these observations, other reports have also shown that neutrophils isolated from blood of psoriatic patients show frequent NET formation, and NET-associated enzymes are further localized in psoriatic lesions (23). Furthermore, it was recently shown that NETosis requires autophagy and the generation of superoxide (22). To investigate the role of autophagy in this process, we blocked the selective macroautophagy pathway by ablating Wdfy3, an adaptor protein responsible for the degradation of aggregated proteins in macroautophagy (19). Interestingly, Wdfy3 contains a BEACH domain, which has been associated with neutropenia and defects in lysosomal trafficking, in Chediak–Higashi syndrome (20, 21).

Wdfy3, the master regulator of selective macroautophagy, similarly to NE, was also highly expressed in neutrophils in vitro, and in vivo this expression was localized within the inflamed skin. Our cre-lox system approach with a LysM promoter to selectively ablate Wdfy3 in myeloid cells also resulted in protection from disease. Although LysM is not specific to neutrophils and is found in various myeloid cells, our adaptive transfer of CD11b⁺Gr-1^hi cells from IL-17A gene transfer mice and the fact that IL-17A gene transfer did not induce any other myeloid population (data not shown) confirmed the specificity. Moreover, there were no detectable differences in the percent of apoptotic or necrotic neutrophils in the absence of Wdfy3, nor an apparent difference in the priming, activation, or phagocytic capacity of the Wdfy3-deficient neutrophils compared with controls, suggesting that no other neutrophil function was impaired in the Wdfy3-deficient neutrophils. We further confirmed our observations using PMA-stimulated neutrophils as previously described by Remijsen et al. (22). Our data showed that Wdfy3-deficient neutrophils exhibit a marked reduction in intracellular ROS compared with controls, suggesting that Wdfy3 may attenuate NETosis via modulating ROS.

Although our data identify NETosis as a major player in the pathogenesis of epidermal hyperplasia, other cellular mechanisms may also contribute. For instance, on a cellular level, nucleic acids can induce myeloid cell activation to stimulate the secretion of various proinflammatory cytokines to recruit inflammatory cells such as neutrophils, which in the presence of stimulatory cytokines and danger signals promote the cycle of inflammation within the skin (37). Because NE deficiency in mice can lead to increased susceptibility to infections (38), it is possible that Wdfy3 may be a more suitable target with a higher therapeutic potential.

Collectively, these results identify Wdfy3 as a novel regulator of neutrophil function, which may be exploited to debilitate neutrophil function as a therapeutic approach in cutaneous disorders such as psoriasis.

We thank Hyun-Seock Shin and Jack Davis for technical assistance with the preparation of MC DNA and histological sections, respectively. We thank Dr. Blythe P. Durbin Johnson and Dr. Matt Lee Settles at the Bioinformatics Core in the Genome Center at the University of California at Davis for assistance with DNA microarray analysis.

The authors have no financial conflicts of interest.