1. Introduction
Diabetic retinopathy (DR) is one of the most prevalent diabetic complications and a leading contributor to vision loss and blindness in the adult working population [
1,
2]. The detailed pathogenesis of DR is unclear, but it is accepted that inflammation is a key contributory factor to its multifactorial pathology [
3,
4,
5]. This is supported by overwhelming evidence wherein infiltrating leukocytes, including peripheral blood mononuclear cells (PBMCs) and the cytokines which they release, are involved in DR pathological changes including increased leukostasis, capillary degeneration, neurodegeneration, vascular permeability and atypical immune responses [
3,
4,
5,
6,
7,
8,
9].
Cytokines are a class of proteins produced by different types of cells—mostly leukocytes either constitutively or after activation; they function as molecular mediators of the innate and adaptive immunities, typically serving as intermediaries within and between these subsystems. The families of cytokines include chemokines which induce chemotaxis; interleukins (including most lymphokines) which perform various functions including maturation and proliferation of immune cells; interferons which function mainly to resolve pathogenic presence; as well as the tumor necrosis factor (TNF-α) family which regulates immune cell functions [
10,
11]. Various cytokines have been implicated in DR pathophysiology, as high concentrations of leukocyte-derived cytokines and growth factors including IL-1β, TNF-α, IL-1, IL-2, IL-6 and IL-8 have been reported in the vitreous humor, serum and plasma of DR patients [
12,
13,
14,
15,
16]. In diabetes, a hyperglycemic microenvironment inundated with dysregulated cytokine secretion and expression is a recipe for low-grade cellular activation and inflammation (parainflammation) that may persist to prompt the onset of diabetic complications, including DR and its progression [
17,
18].
We previously reported that DR in type-1-diabetes (T1D) is associated with increased innate and reduced adaptive cellular immunity [
19]. Despite such altered immunophenotype, we lack detailed information on how PBMC-cytokine production is altered in people with T1D at different stages of DR (early and advanced). In this follow-up study, we investigated whether cytokine production and secretion profiles of innate (CD11b
+/myeloid cells) and adaptive (CD3
+) peripheral blood immune cells are dysregulated in people with T1D with DR. Furthermore, we investigated whether PBMC cytokine secretion and expression is altered due to treatment in patients with advanced DR (proliferative diabetic retinopathy (PDR)). Uncovering this information is critical, not only to improve our understanding of DR pathophysiology, but also to design more effective therapies capable of normalizing the altered immune response underpinning DR. Furthermore, an association between diabetes’ duration and DR initiation and progression has been established [
20,
21,
22,
23,
24], but the potential relationship between diabetes’ standing and specific PBMC-expressed, PBMC-secreted and/or plasma-expressed cytokine biomarkers have not been investigated in people with T1D with DR.
Thus, herein, we determine cytokine secretion and expression profiles of different subsets of PBMCs under normal and stimulatory culture conditions as well as the plasma cytokine profiles in patients with T1D and early and advanced DR and sought to understand whether these relate to diabetes’ duration. The stimulatory conditions included (1) high glucose (HG) to mimic hyperglycemia, (2) mannitol to mimic hypertonicity and serve as osmotic control for HG, (3) phorbol myristate acetate (PMA) and Ionomycin to mimic inflammatory conditions and (4) lipopolysaccharide (LPS) to mimic the endotoxin-induced inflammatory environment.
2. Materials and Methods
2.1. Study Participants
The study was approved by the Office for Research Ethics Committees Northern Ireland (ORECNI, Ref: 14/NI/0084) and was conducted in accordance with the Declaration of Helsinki. All participants were informed about the study and provided written informed consent prior to their enrolment. Forty-one adults with T1D (Age ≥ 18) were recruited into three groups: 13 T1D with mild non-proliferative DR (mNPDR), 14 T1D with active proliferative DR (aPDR) naïve to treatment and 14 T1D with PDR that had received laser PRP and were clinically stable and quiescent afterwards (inactive PDR; iPDR). Thirteen age- and gender-matched healthy controls were also recruited. The diagnosis of mNPDR, aPDR, iPDR in T1D and of no DR in healthy controls was made based on clinical history and clinical examination, including fundus photography. Medical and family history, current medications, body mass index (BMI) and systolic and diastolic blood pressure were obtained. Exclusion criteria included history of cardiac disease or malignancy within the past 5 years; history of inflammatory diseases within the past 2 months; and other retinal disorders besides DR, active autoimmune disease and history or current use of immunosuppressive medications or steroids. Pregnant females, people with kidney failure and incapacity to undertake eye imaging due to any reason were also excluded from the study.
2.2. Masking
Researchers who carried out laboratory experiments and analysis were masked to the origin of the clinical samples to be analyzed (i.e., whether they were coming from healthy volunteers or from patients with T1D and DR).
2.3. PBMC Isolation, Culture and Supernatant Collection
Venous blood (20 mL) was collected into tubes containing ethylenediaminetetraacetic acid (ETDA; BD Biosciences, Oxford, UK). Within 3 h of blood collection, PBMC were isolated by Ficoll-Paque (Histopaque; Sigma-Aldrich, Cambridge, UK) density gradient centrifugation. The PBMCs were cultured in RPMI 1640 medium containing 10% FCS and 1% penicillin-streptomycin under normoxia (21% oxygen); and incubated with either Escherichia coli lipopolysaccharide (LPS; 2.5 µg/mL), or PMA (100 ng/mL) + ionomycin (1 µg/mL), or 25 mM D-glucose, or 25 mM D-mannitol (all from Sigma–Aldrich(Cambridge, UK)) for 16 h. The supernatants were collected and stored at −80 °C until analysis.
2.4. Plasma Collection
After blood collection, the plasma was isolated by centrifugation of samples at 1200 rpm for 10 min at room temperature (RT). The plasma fraction was centrifuged again at 3000 rpm for 15 min at RT to pellet any residual cells and platelets. The samples were aliquoted and stored at −80 °C until analysis.
2.5. Protein Measurements in Plasma and PBMC Culture Supernatant
Soluble CD121a (sCD121a), IL-6, IL-8 and MCP-1 expression in plasma, and expression of cytokines: IL-1α, IL-1β, IL-6, IL-8, IL-10, IL-17A, IFN-γ, MCP-1 and TNF-α in culture supernatants of PBMCs with or without LPS stimulation were measured using Cytometric Bead Array Human Soluble Protein Detection Kit (CBA; BD Biosciences) per the manufacturer’s instructions. Briefly, capture beads coated with anti-cytokine antibodies were mixed and incubated with plasma or PBMC supernatant or standards in round bottom 96-well plates (Nunc; Thermo Scientific, Leicestershire, UK) for 1 h at RT in the dark. This was followed by an additional 2 h of incubation with phycoerythrin (PE)-conjugated anti-cytokine detection reagent at RT in the dark. The samples were washed, and the bead pellets were re-suspended in washing buffer. The resuspended samples were then run on a flow cytometer (FACS Canto II; BD Biosciences) that was equipped with BD Diva software. Collected data were subsequently analyzed using BD FCAP Array software version 3 (BD Biosciences). The total protein concentration per sample was measured using a Pierce BCA protein assay kit (Thermo Fisher Scientific, Waltham, MA, USA). The concentration of each cytokine was normalized to the total protein concentration in supernatant (pg/mg total protein).
2.6. PBMC Flow Cytometry: Surface Markers and Intracellular Cytokines
PBMCs cultured for 16h with different stimulants as described above were incubated alongside 1× monensin (Biolegend UK Ltd. London, UK) for 4 h. Monensin is a protein transport inhibitor that prevents cytokine secretion, thereby trapping them intracellularly. PBMCs were washed twice with FACS buffer (300× g, 4 °C for 7 min) and resuspended at 10 × 106 cells/mL; 20 µL (2 × 105 cells) of single cell suspension were allotted per FACS tube and incubated with 5 μL Human TruStain FcX (Fc Receptor blocking solution; BioLegend UK Ltd.) for 5 min at RT.
The cells were then incubated with cell surface fluorochrome-conjugated antibodies (See
Table 1) in a total volume of 100 μL FACS buffer for 30 min in the dark at 4 °C. Post-fluorochrome staining, cells were washed twice with FACS buffer and then fixed and permeabilized using the Foxp3 Transcription Factor Staining Buffer Set (eBioscience, San Diego, CA, USA) according to the manufacturer’s instructions. Samples were then incubated with 5 μL Human TruStain FcX (Fc Receptor blocking solution; BioLegend UK Ltd.) followed by incubation with either intracellular cytokine fluorochrome-conjugated antibodies or appropriate isotype controls (see
Table 1) in a total volume of 100 μL permeabilization buffer (eBioscience) for 40 min in the dark at 4 °C. Cells were washed and acquired on the FACSCanto II flow cytometer (BD Biosciences). Data analysis was performed using FlowJo software version 10.07 for Windows (Tree Star, Ashland, OR, USA).
2.7. Flow Cytometry Gating Strategies
Flow cytometry confirmed the depletion of granulocytes in isolated PBMCs (
Figure 1a, right panel). Lymphocytes and myeloid-derived cells were identified by their surface expression of CD3 or CD11b (
Figure 1b) as well as using an FSC vs. SSC dot-plot to gate on PBMC myeloid cells (
Figure 1a, right panel). To analyze the expression of intracellular cytokine expression, gates of intracellular cytokines (e.g., IL-6, IL-17A, IL-10 and IFN-γ) were set on total live cells (
Figure 1c–f), as well as on CD11b
+ (
Figure 1g), PBMC myeloid cells (
Figure 1h–j) and CD3
+ (
Figure 1k–m) based on the appropriate isotype controls.
2.8. Statistical Analyses
Statistical calculations were performed with GraphPad Prism 6 (GraphPad, San Diego, CA, USA). Normality of continuous variables was determined using D’Agostino–Pearson omnibus normality test. Comparisons between two normally distributed groups were carried out using independent samples Student’s t-test while the Mann–Whitney test was used when normality was not confirmed. Linear regression analysis was used to investigate potential associations between T1D duration and PBMC cytokine expression. Data were presented as mean ± standard error of the mean (SEM) in figures and mean ± standard deviation (SD) in tables. p values < 0.05 were considered statistically significant.
Figure 1.
Gating strategies used in flow cytometry analysis to identify PBMC subsets and quantitate cytokine expression. (a,b) Representative data showing PBMC isolation and subsets: (a) Forward scatter (FSC−A) vs. side-scatter (SSC−A) plot of whole blood before (Left panel) and after PBMC isolation (Right panel), note absence of polymorphonuclear granulocytes. (b) CD3 vs. CD11b showing CD3+ and CD11b+ PBMCs. (c–f) Representative data used to quantitate overall cytokine expression via cytokine vs. SSC−A plots to indicate the PBMC expression of: (c) IL-6; (d) IL-17A; (e) IL-10 and (f) IFN-γ. (g–i) Representative data used to quantitate cytokine expression in the CD11b+ and myeloid cell subsets of PBMCs via cytokine vs. CD11b+ or SSC−A plots: (g) IL-6; (h) IL-17A (Left panel shows myeloid cell IL-17A expression in contour display while right panel shows IL-17A expression without isotype control overlay in zebra display) and (i) IL-10. (j–m) Representative data used to quantitate cytokine expression in the CD3+ subsets of PBMCs via cytokine vs. CD3 plots: (j) IL-6; (k) IL-17A; (l) IL-10 and (m) IFN-γ. Populations in blue color have been stained with isotype controls while populations in red color have been stained with dye-conjugated antibodies. The gate frequencies displayed on plots are the percentages of events within each gate.
Figure 1.
Gating strategies used in flow cytometry analysis to identify PBMC subsets and quantitate cytokine expression. (a,b) Representative data showing PBMC isolation and subsets: (a) Forward scatter (FSC−A) vs. side-scatter (SSC−A) plot of whole blood before (Left panel) and after PBMC isolation (Right panel), note absence of polymorphonuclear granulocytes. (b) CD3 vs. CD11b showing CD3+ and CD11b+ PBMCs. (c–f) Representative data used to quantitate overall cytokine expression via cytokine vs. SSC−A plots to indicate the PBMC expression of: (c) IL-6; (d) IL-17A; (e) IL-10 and (f) IFN-γ. (g–i) Representative data used to quantitate cytokine expression in the CD11b+ and myeloid cell subsets of PBMCs via cytokine vs. CD11b+ or SSC−A plots: (g) IL-6; (h) IL-17A (Left panel shows myeloid cell IL-17A expression in contour display while right panel shows IL-17A expression without isotype control overlay in zebra display) and (i) IL-10. (j–m) Representative data used to quantitate cytokine expression in the CD3+ subsets of PBMCs via cytokine vs. CD3 plots: (j) IL-6; (k) IL-17A; (l) IL-10 and (m) IFN-γ. Populations in blue color have been stained with isotype controls while populations in red color have been stained with dye-conjugated antibodies. The gate frequencies displayed on plots are the percentages of events within each gate.
4. Discussion
Our results suggest that T1D DR PBMCs, particularly CD11b myeloid cells, are proinflammatory, and the increased IL-6, IL-10 and IL-17A derived from them may contribute to the retinal pathology in DR. A decreased level of PBMC-derived IFN-γ was detected, which may contribute to the diminished adaptive immunity in T1D DR patients [
19]. Our observations are consistent with those of our previous study [
19] where we found the proinflammatory immunophenotype in T1D DR patients is the result of an enhanced innate response and an impaired adaptive response. Our data also suggest that the IL-1/CD121a pathway may contribute to the early stages of DR.
Regarding cytokine secretion from PBMC, a higher level of IL-10 appears to be related to T1D DR pathology and may be involved in the early stages of DR as it was increased predominantly in mNPDR but less so in aPDR. Higher levels of IL-1α and IL-6 were also observed in mNPDR compared to healthy controls, suggesting that they may contribute to early T1D DR. The likely sources of the increase in secreted IL-6 and IL-10 related to early T1D DR are myeloid PBMCs. In the advanced stage of DR, the secretion of IL-6 and IL-10 by PBMCs was decreased, suggesting that this reduced secretion may be involved with the active proliferation of vessels in DR.
Concerning intracellular cytokine expression profiles in T1D DR, the increase in IL-17A
+ PBMCs, especially CD11b
+IL-17A
+ cells and decrease in CD3
+IFN-γ
+ cells appear to be critical factors. Decreased IFN-γ and IL-6 expression by CD3 T cells is likely to be involved in the early stages of T1D DR. Meanwhile, higher intracellular IL-6 expression by CD11b
+/myeloid cells and CD3
+ T cells is related to the advanced stages of T1D DR. These data have value in helping us to understand how the immune system may contribute to DR, especially at the early and advanced stages of DR. Activated immune cells may participate in retinal vascular/neuronal degeneration by (1) releasing inflammatory cytokines into the plasma. Consequently, these cytokines circulate throughout the body and increase to a threshold where they damage endothelial cells; (2) releasing inflammatory cytokines which directly affect other cells that are in close contact with them. For example, leukostasis of CD11b
+ cells [
25] may release IL-17A to adjacent endothelial cells leading to BRB damage. In this case, locally produced IL-17A is sufficient to damage endothelial cells even though systemic levels of IL-17A may remain unchanged; (3) releasing “protective” or anti-inflammatory cytokines as part of a compensatory immune mechanism that may lead to repair. Consequently, this anti-inflammatory mechanism becomes dysfunctional, leading to an imbalance between pro-inflammatory and anti-inflammatory immune mechanisms that may allow DR to progress to an advanced stage.
IL-17A is a proinflammatory cytokine involved in the augmentation of immune response by prompting increased production of other proinflammatory cytokines including IL-6 and IL-1β, thereby creating a connection between T-cells’ activation and inflammation, implicating it in various autoimmune diseases [
26,
27]. IL-17A can be produced by Th17 T cells [
26], γδ T cells and NK cells [
28,
29]. Our data suggest that the increase in IL-17A
+ PBMCs may be involved in T1D DR, which is in line with findings by Honkanen et al. [
30], wherein the authors reported increased IL-17 expression and secretion in T1D PBMCs. Systemic levels of IL-17A are significantly elevated in the serum of patients with T2D DR [
31] and in T1D DR [
32] as well as in PBMCs of T2D patients with no DR [
8] compared to controls. We recently reported that IL-17A can directly damage the BRB via the JAK1 signaling pathway [
33]. Data from the current study and the literature suggest that IL-17A may be a critical player in sustaining DR pathology.
In DR, the plasma, serum [
12,
14], vitreous [
15,
16] and PBMCs of DR patients express higher concentrations of IL-6 [
34]. We found IL-6-producing CD11b
+ cells were significantly increased in aPDR and the level was positively correlated with T1D duration. Diabetes’ duration is a clinical risk factor for DR and our data suggest that T1D duration has a positive correlation with PBMC-derived IL-6 which may promote DR pathology. The increased IL-6 intracellular expression was paired, however, with a decreased IL-6 secretion in aPDR. This suggests PBMC IL-6 accumulation and defective IL-6 trafficking/signaling in the aPDR stage. It has been suggested that the IL-6 secretion bottleneck is mostly due to defective IL-6 transcription and translation [
35]; however, the IL-6 trafficking process is not fully understood, and it is unclear whether the increase in IL-6 synthesis with decrease in IL-6 trafficking into the microenvironment is due to deficiencies in the post-Golgi trafficking pathways that mediate IL-6 release or insufficient external cellular stimulation to prompt cytokine release. Remarkably, IL-6 is an important activator of signal transducer and activator of transcription 3 (STAT3) [
36,
37]. We have previously shown [
38] that in T1D DR, there are higher levels of activated STAT3 in circulating leukocytes of patients with mNPDR but not in aPDR, which may be explained by current findings.
IL-10 plays an immunosuppressive role by inhibiting IL12 and TNF-α production [
39,
40] and is mainly produced by Th2 cells [
41]. Although IL-10 is considered to be anti-inflammatory, especially on monocytes due to its suppression of production and secretion of proinflammatory cytokines such as IL-12, IL-8 and TNF-α [
42], IL-10 can induce various signaling that initiate or sustain inflammation [
36,
37]. In agreement with our studies, reports have shown that IL-10 production is elevated in T1D patients compared to controls [
32,
43] as well as in T1D diabetic nephropathy. Increased secretion of IL-10 by PBMCs in the early stages of DR may be a compensatory immune mechanism that confers protection on the retina. However, our data suggest that after an extended period of increased anti-inflammatory activity of IL-10, as DR advances, there is a reduction in PBMC-secreted IL-10 which may contribute to neovascular processes in aPDR. The exact cause for reduced PBMC secretion of IL-10 in advanced DR is unclear, but it is likely that the PBMCs that release IL-10 may become impaired, allowing the advancement of DR.
IFN-γ is mainly produced by Th1 cells after their differentiation in response to pathogenic infections, driving cell-mediated immunity and stimulating B-cells for opsonizing antibody production [
44]. Irregular IFN-γ expression is linked with autoinflammatory and autoimmune diseases [
45]. The diminished IFN-γ observed in DR patients especially in mNPDR patients aligns with Foss et al. [
43] and may shed more light on diminished adaptive immunity, susceptibility to infections and non-resolution of low-grade inflammation in diabetic patients [
19,
43,
46,
47].
We found increased levels of secreted IL-1α in supernatant of mNPDR PBMCs compared to controls. We also found that sCD121a or sIL-1R1, an inflammatory receptor to IL-1α, IL-1β and IL-1Ra [
48] is increased in the plasma of T1D DR, and in mNPDR compared to controls. The binding actions of CD121a account for IL-1 mediated inflammation and pathology [
48,
49]; loss of IL-1 signaling in CD121a
−/
− mice attenuates the formation of acellular capillaries in diabetic retina [
50] and infiltration of pro-inflammatory leukocytes into ischemic tissue [
51]. Our results suggest that increased IL-1α secretion and plasma levels of sCD121a may be involved in T1D DR by activating IL-1-mediated inflammation. sCD121a may be released as a protective response during the early stages of DR to counteract the noxious effects of IL-1α, and IL-1/CD121a signaling may partially explain early retinal changes including capillary acellularity in DR.
The strengths of this study include its prospective study design, the detailed phenotyping of patients and, importantly, the masking of laboratory researchers to the precedence of the blood samples (i.e., whether they had been obtained from people with DR or from healthy controls). The additional strengths of the study include (1) the use of different stimuli to induce cytokine production in PBMC and (2) the detection of intracellular cytokines. This study design allowed us to understand how the immune cells would respond under disease conditions and the key cellular sources of disease-causing cytokines (e.g., increased IL-6 and IL-17 production by CD11b cells and impaired IFN-γ production by CD3 cells in DR). Limitations include the cross-sectional study design, small patient numbers in each of the DR subgroups and the recruitment of all patients from a single site (Belfast, Northern Ireland). Additionally, it is difficult to determine what size of change in the different data presented may be mechanistically important and clinically relevant, independently of whether statistically significant. Furthermore, we detected differences in values among the different individuals studied. This interindividual variability, however, is not surprising since it is expected that pathogenic pathways may not affect all individuals in the same manner. Even within “homogeneous” groups of people with DR based on ETDRS severity, for example, many patients will reach the moderate non-proliferative DR stage but only few will develop DMO or PDR. Furthermore, some patients may develop DMO and never progress to PDR and vice versa; some will suffer both, others neither. Thus, is it likely that the differential weight of pathogenic pathways among people with T1D DR may explain subtle subject-to-subject variability in phenotype and differences in onset, progression and even complications of DR.