Possible involvement of NETosis in inflammatory processes in the eye: Evidence from a small cohort of patients

Animals

Forty 8-week-old pigmented male C57BL/6J mice in two sequential experiments were used in this study. Animals were obtained (Harlan Biotech Israel Ltd., Jerusalem, Israel) and handled according to the recommendations of the ARVO Statement for the Use of Animals in Ophthalmic and Visual Research and the hospital’s Institutional Animal Care and Use Committee.

Induction of ocular inflammation

Animals were anesthetized with intraperitoneal (IP) injection of ketamine 40 mg/kg and xylazine 10 mg/kg, and the pupils were dilated with topical administration of 0.8% tropicamide eyedrops. Intravitreal injections were performed under an operating microscope (Zeiss Opmi 6S Microscope; Carl Zeiss Microscopy GmbH, Oberkochen, Germany). A microsyringe (33-gauge; Hamilton syringe, Energy Way, Reno, NV) was placed intravitreally in the retrolental space of the eye, and 1 μl per mouse of interleukin-8 (IL-8; 100 ng/µl), (R&D Systems, Minneapolis, MN) or tumor necrosis factor alpha (TNF-α; R&D Systems; 50 ng/µl) was injected. The control group was injected with sterile saline in a similar manner.

Histology

At each time point (n=5/group), animals were euthanized with carbon dioxide, and the eyes were enucleated. The eyes were fixed in 4% paraformaldehyde (PFA) in PBS (Dissolve 1 tablet in 200 ml water to obtain a 137 mM NaCl, 2.7 mM KCl and 10 mM phosphate buffer solution, pH 7.4 at 25 °C) for 1 h, dehydrated by increasing the sucrose gradient (5%, 10%, 20%, and 30%, 30 min to 1 h each), and cryopreserved in optimum cutting temperature (OCT; Thermo Fisher Scientific, Grand Island, NY). Serial 10-µm cryosections parallel to the optic nerve axis were obtained and stained with hematoxylin and eosin (H&E; American MasterTech Scientific, Lodi, CA). For histopathologic evaluation, the anterior and posterior chambers were examined under light microscopy (Olympus Optical Co., Tokyo, Japan).

Antibodies

The primary antibodies used in this study were rabbit polyclonal NE (Abcam, Cambridge, MA, 1:100), fluorescein isothiocyanate (FITC)-conjugated mouse monoclonal to MPO (Abcam, 1:200), rabbit polyclonal citrulline histone 3 (H3Cit; Abcam, 1:100), rat monoclonal CD11b (Abcam, 1:100), and rat monoclonal Ly6G (Abcam, 1:100). Secondary antibodies against the primary antibodies were donkey anti-rabbit AlexaFluor 568 (Invitrogen, Grand Island NY, NY, 1:500), donkey anti-rabbit 647 (Abcam, 1:200), donkey anti-rat 488 (Abcam, 1:500), and donkey anti-rat AlexaFluor 568 (Abcam, 1:500), respectively. Matched isotype controls were used as internal controls.

Immunofluorescence

A multistep staining protocol was designed based on the characteristics of the primary antibodies used. Briefly, the slides were hydrated, permeabilized with 0.5% TritonX-100 in PBS for 10 min, followed by blocking with either 10% goat or donkey antiserum in PBS (depending on the secondary antibody used) for 1 h. Thereafter, the slides were washed several times and incubated with the primary antibody in blocking solution for 1 h at room temperature. Tissue sections were then rinsed several times with PBS and incubated for 1 h at room temperature with a secondary antibody against the respective primary antibody. A similar procedure was followed for all antibodies in a sequence procedure. Then the slides were rinsed, and the cell nuclei were counterstained with 4',6-diamidino-2-phenylindole (DAPI; Invitrogen, 2 drops/ml according to the manufacturer’s protocol), mounted with mounting medium (ProLong^TM Gold Antifade mountant, Invitrogen, Grand Island, NY), and images were captured digitally using the Axio Imager Apotome microscope (Zeiss, Göttingen, Germany). A minimum of 20 fields at 63X magnification per case was used to evaluate for the presence of neutrophils (CD11b). Similarly, the presence of NE/MPO/H3Cit triple staining in which nuclear morphology and NETs were determined.

DNase I treatment

Consecutive cryosections positive for NETs were hydrated, permeabilized, and either left untreated or subjected to digestion with DNase I (Abcam) by 20 U/ml for 30 min at 37 °C. Thereafter, the slides were washed and stained for H3Cit using the immunofluorescence protocol mentioned above.

Study population

The study was approved by the Rabin Medical Center Institutional review board and adhered to the tenets of the Declaration of Helsinki and ARVO statement on human subjects and was approved by the Ethics Committee. The study was approved by the institutional review board; all patients provided written informed consent before the procedure. The study included 24 patients who underwent pars plana vitrectomy during 2013–2016. Patients were divided according to their pathology. Patients with PDR were divided into two groups: PDR1 (no need for repeated surgical intervention) and PDR2 (repeated vitreous bleeding or other complication). The control group (n=10) was composed of patients with macular hole (MH) or epiretinal membrane (ERM). Exclusion criteria were age less than 18 years, previous vitrectomy, or combined cataract and pars plana vitrectomy. All patients underwent ophthalmic examination, and the medical and ocular history was recorded.

Sample collection

Vitreous samples were obtained via standard pars plana vitrectomy. During acquisition, infusion was set to air to prevent dilution. Samples were aliquoted (100 µl/vial) and stored at −80 °C pending laboratory analysis. No more than one freeze–thaw cycle was allowed before analysis.

Analyzing levels of MPO, H3Cit-MPO complex, and NE-MPO complex in the vitreous

MPO levels in the vitreous samples 100 µl/well (the samples were diluted 1:200 with a dilution buffer, R&D Systems) were measured using the Human MPO DuoSet enzyme-linked immunosorbent assay (ELISA) kit, according to the manufacturer’s protocol (R&D Systems). MPO is present in extruded NET complexes mostly associated with H3Cit and NE. Such complexes were quantified with modified capture ELISA [11] In brief, a 96-well plate was precoated with human anti-H3Cit (Abcam, dilution 1:12,000) or human anti-NE antibodies (Abcam, dilution 1:12,000) overnight. The following day vitreous sample (diluted 1:200 with a dilution buffer, from the MPO DuoSet ELISA kit, R&D Systems) after which the MPO-associated complexes were detected using the human MPO ELISA kit, according to the manufacturer’s protocol (R&D Systems).

Statistical analysis

A descriptive statistic was calculated for each marker. MPO, H3CitH3, and NE-MPO levels were compared between groups using linear transformation and one-way ANOVA (SPSS software). Least significant difference post hoc analysis was conducted to evaluate the significance of differences between groups. A p value of less than 0.05 was considered statistically significant, and a p value of less than 0.01 was considered statistically highly significant.

Histopathology

Histopathological analyses were conducted to evaluate cellular infiltration upon stimulation. Figure 1 is a representative image of three consecutive experiments with five animals per group. The sham-injected eyes presented with clear anterior and posterior chambers devoid of infiltrating immune cells (Figure 1A,D). IL-8 (Figure 1B,E) and TNF-α (Figure 1C,F) induced massive cellular infiltration, ranging in size and shape, into the anterior and posterior chambers (vitreous body), respectively. Cell infiltration was seen in the anterior chamber (arrows) and cornea (arrowheads), as well as in the ciliary body and the iris (data not shown). In the posterior chamber, cellular infiltration was seen in the vitreous body (Figure 1E,F, arrows) along with retinal folding, disorganization of the sensory layers, as well as retinal gliosis (data not shown; Figure 1).

Cellular infiltration

As neutrophils are the first responders at sites of infection [2] and play a key role in NETosis [4], we evaluated eyes injected with IL-8 and TNF-α for their presence, using the CD11b marker. The sham-injected eyes were negative for CD11b+ cells in the anterior and posterior chambers (Figure 2A,D, respectively). Massive infiltration of CD11b+ cells was observed in the anterior chamber and the vitreous (the posterior chamber) of the eyes injected with IL-8 (Figure 2B,E) and TNF-α (Figure 2C,F), respectively. The presence of CD11b-positive cells is consistent with the histopathology results shown in Figure 1. Neutrophils were further validated with costaining for Ly6G and CD11b (Appendix 1) and defined by the classic polymorphonuclear granulocyte morphology (Appendix 2; Figure 2).

NET production by infiltrating neutrophils

We then asked the question whether the infiltrating neutrophils were associated with NETs formation. Eyes were examined for the presence of specific NET markers by triple-staining for NE (red), MPO (green), and H3Cit (purple) at different time points. NET complexes were found only at 24 h post-injection. Figure 3 and Figure 4 are representative images of the anterior and posterior chambers, respectively. The sham-injected eyes were neutrophil-free and negative for all three markers (Figure 3 and Figure 4A,D,G,J,M). Netting neutrophils were found in the anterior and posterior chambers of the eyes injected with IL-8 and TNF-α and were based on costaining of NE (red; Figure 3, Figure 4E,F; arrows), MPO (green; Figure 3, Figure 4H,I; arrows), and H3Cit (purple; Figure 3, Figure 4K,L, arrows). DAPI (blue; Figure 3, Figure 4N,O, arrows) shows the nuclear morphology. Both cytokines were accompanied by many NET loci, associated with very strong staining of citrullinated histones (purple). NETosis was particularly pronounced in areas of aggregated neutrophils and was manifested by the extracellular colocalization of all the three markers as shown in the merged images (Figure 3, Figure 4B,C). Isotype controls were used as internal controls (Appendix 3). No significant difference was found for the time period of NET clearance between the anterior and posterior chambers.

NET degradation with DNase I treatment

We confirmed the presence of extracellular DNA in the NETs using DNase I treatment followed by immunostaining for citrullinated histone. Figure 5A,B illustrate the decrease in the number of H3Cit loci for IL-8 and TNF-α, respectively. The sham-injected eyes were negative for H3Cit staining indicating the absence of the NETs ex vivo treatment with DNase exerted no effect in these samples (Appendix 4 and Appendix 5). IL-8 and TNF-α were associated with infiltrated cells surrounded by extruded DNA (indicated by the H3Cit-positive staining) in the anterior and posterior chambers (Figure 5A,B, black bars). DNase I treatment induced a significant degradation of DNA, leaving fade traces of H3Cit (Figure 5A,B, gray bars, p<0.05 and p<0.001, anterior and posterior chambers, respectively).

Evaluation of NETs complexes in vitreous samples of patients with PDR

We evaluated the generation of NET components, such as MPO, H3-CitH3, and NE-MPO complexes, in the vitreous of patients with PDR. The demographic and clinical characteristics of the patients with PDR are summarized in Table 1.

The patients were divided into two groups based on the severity of PDR [20]: The PDR1 group had no need for repeated surgical intervention, and the PDR2 group had repeated vitreous bleeding or other complication. The results are summarized in Figure 6. We found that patients with PDR2 exhibited significantly higher levels of MPO (Figure 6A; 173±230) compared to the patients with PDR1 (Figure 6A; 12±33, *p<0.05) or the controls (Figure 6A; 0, **p<0.01). The levels of H3Cit-MPO and NE-MPO complexes were also statistically significantly higher in the patients with PDR2 (Figure 6B; 776.0±1274, Figure 6C; 573±911, respectively) compared to those in the patients with PDR1 (Figure 6B,C; 0, *p<0.05) and the controls (Figure 6B,C; 0, *p<0.05).

Neutrophil validation

We evaluated the eyes injected with IL-8 and TNF-α for their presence, using the CD11b/Ly6G markers. The sham-injected eyes were negative for CD11b/Ly6G cells in both posterior chambers (Appendix 1). Massive infiltration of CD11b/Ly6G cells was observed in the posterior chamber of the eyes injected with IL-8 and TNF-α (Appendix 1). The presence of Ly6G (Appendix 1) and CD11b (Appendix 1) positive cells is consistent with the histopathology results shown in Figure 1. Neutrophils were validated as defined by the classic polymorphonuclear granulocyte morphology (Appendix 2, DAPI staining, higher resolution).

Validation of NET production: Isotype control analysis

As NET production is the scope of this paper and authentication of the results was highly valuable, staining using isotype controls was conducted throughout the study for each of the antibodies and multiple sites of the anterior and posterior chambers. As can be seen, the donkey anti-rabbit AlexaFluor 568 had no non-specific staining at either the anterior or posterior chambers (Appendix 3, respectively). Minimal autofluorescence was observed when the donkey anti-rat 488 at both the anterior and posterior chambers (Appendix 3, respectively). No staining was observed for the donkey anti-rabbit 647 isotype control (Appendix 3, anterior and posterior chambers, respectively). DAPI nuclear staining was used as an internal control for neutrophil validation and localization (Appendix 3, arrows).

NET degradation by DNase I treatment

The presence of extracellular DNA in the NETs was evaluated using DNase I treatment followed by immunostaining for citrullinated histone. Appendix 4 and Appendix 5 are representative images of the anterior and posterior chambers, respectively. Sham-injected eyes were negative for H3Cit staining, indicating the absence of NETs (Appendix 4 and Appendix 5). Ex vivo treatment with DNase exerted no effect in these samples. IL-8 and TNF-α were associated with infiltrated cells surrounded by extruded DNA, as indicated by positive staining for H3Cit (purple; Appendix 4 and Appendix 5, arrows). Upon DNase I treatment, the majority of the DNA was degraded, leaving fade traces of H3Cit (Appendix 4 and Appendix 5, arrows). Interestingly, in the IL-8 group, the ganglion cell layer was also positive for H3Cit (Appendix 5), which may be related to retinal gliosis (see also histological section, Figure 1).