Introduction

Dopamine (DA) is a key neurotransmitter in the brain, where it is involved in a wide variety of central nervous system (CNS) functions including motivation, cognition, movement and reward. DA is however also increasingly regarded as a crucial transmitter in the neuroimmune network, contributing to the CNS-immune system interplay as well as in the communication among immune cells (Basu and Dasgupta 2000; Sarkar et al. 2010; Levite 2012). DA can affect most if not all human immune cells, including T and B cells, dendritic cells, macrophages, microglia, neutrophils and NK cells, and it has been suggested that DA exerts mainly stimulatory effects on resting immune cells and inhibitory effects on activated immune cells (Sarkar et al. 2010; Levite 2012). Immune cells themselves produce DA, which may act as autocrine/paracrine mediator on immune cells as well as on neighbouring cells (Bergquist and Silberring 1998; Cosentino et al. 1999; Marino et al. 1999; Cosentino et al. 2000; Cosentino et al. 2002a and b, Cosentino et al. 2005). Prominent examples include the inhibitory loop subserved by endogenous DA in human CD4+CD25high regulatory T lymphocytes, a specialized T cell subsets playing a key role in the control of immune homeostasis (Cosentino et al. 2007), and the influence exerted by dendritic cells-derived DA on the differentiation of naive CD4+ T cells (Nakano et al. 2009).

The involvement of DA-operated modulation of the immune response is well established in human disease such as multiple sclerosis (MS) (Zaffaroni et al. 2008; Cosentino et al. 2012; Cosentino and Marino 2013) and rheumatoid arthritis (RA) (Capellino et al. 2010; Nakano et al. 2011), and preliminary evidence has been recently published concerning their possible contribution to neuroinflammation and neurodegeneration in the 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP)-induced Parkinson’s disease (PD) in the mouse (González et al. 2013). Degeneration of CNS nigrostriatal pathway and the resulting deficit in brain DA is the most prominent feature of Parkinson’s disease (PD) (Dauer and Przedborski 2003) and substitution treatment with directly and indirectly acting dopaminergic agents is the mainstay of PD therapy. In recent years, the role of the adaptive immune system and the interactions between the peripheral immune system and neuroinflammation in PD have been the subject of intense investigation (Laurie et al. 2007; Reynolds et al. 2008; Stone et al. 2009; Saunders et al. 2012; Sanchez-Guajardo et al. 2013). Investigating the ability of DA to modulate the immune response, besides increasing basic knowledge of the neuroimmune network, will provide clues for the development of novel immunomodulating approaches for autoimmune disease such as MS and RA, and will likely clarify the relevance of any direct effects on peripheral immunity in the therapeutic activity of dopaminergic agents currently used in PD.

The effects of DA are mediated by receptors belonging to the 7-transmembrane, G-protein coupled receptors family. DA acts on 5 different dopaminergic receptors (DR) grouped into two families according to their pharmacological profile and main second messenger coupling: the D1-like (D1 and D5) which activate adenylate cyclase and the D2-like (D2, D3 and D4) which inhibit adenylate cyclase (Schetz 2009; Beaulieu and Gainetdinov 2011). The molecular and pharmacological heterogeneity of DR potentially represents an opportunity to develop targeted immunomodulating strategies. For example, DR resulting in the suppression of the regulatory function of CD4+CD25high T lymphocytes belong to the D1-like family (Cosentino et al. 2007), while those directly inducing T cell quiescence are D2-like DR D4 (Sarkar et al. 2006). More detailed information is however required about the patterns of DR expression on immune cells, with regard to differences among cell subsets and the relationship with the functional status of the cells. Unfortunately, only a few studies have been performed so far on human immune cells. According to McKenna et al. (2002), who investigated DR expression on leukocyte subpopulations using flow cytometry, T lymphocytes and monocytes had low expression of DR, neutrophils and eosinophils had moderate expression and B lymphocytes and NK cells had higher and more consistent expression. DR D3 and D5 were found in most individuals, DR D2 and D4 had more variable expression, and DR D1 was never found. In particular, in T lymphocytes DR were found in only 3–5 out of 19 individuals and their frequency was on average no more than 1–5 % (McKenna et al. 2002). Besser et al. (2005) reported the occurrence of DR D2 and D3 on 9–10 % of human T cells, and found no evidence of DR D1 expression, while Sarkar et al. (2006) documented the membrane expression of DR D4 by means of western blot analysis. Finally, Brito-Melo et al. (2012) analyzed the expression of DR D2 and D4 on CD4+ and CD8+ T lymphocytes: DR D2 were expressed on 2–10 % of CD4+ T cells and on 3–16 % of CD8+ T cells, while DR D4 were expressed on 2–16 % of CD4+ T cells and on 1–6 % of CD8+ T cells.

The aim of our study was to examine the expression of DR on circulating CD4+ T lymphocytes by means of a novel flow cytometric assay in whole blood. Phenotyping of DR on CD4+ T cells was performed by a 5-color flow cytometric analysis by use of a two-step protocol which allowed the identification of all the five DR on different cell subsets. In the present paper we report results obtained with human CD4+ naive T cells (CD3+CD4+CD45RA+CCR7+), central memory (TCM, CD3+CD4+CD45RA-CCR7+) and effector memory T cells (TEM, CD3+CD4+CD45RA-CCR7-). We concentrated on naive and memory CD4+ T cells since preliminary evidence supports a role of dopaminergic mechanisms in the differentiation of human naive CD4+ T cells (Nakano et al. 2009). Moreover, CD4+ effector/memory T cell phenotypes have been recently reported as associated with the profile of motor dysfunction in PD patients (Saunders et al. 2012) and we have recently obtained preliminary results showing reduced expression of DR in CD4+ T cells of PD patients (Kustrimovic et al. 2013). The assay was also used in cultured CD4+ T lymphocytes to investigate the expression of DR in viable and apoptotic cells and their changes occurring upon cell activation.

Material and Methods

Antibodies, Chemicals and Reagents

Rabbit polyclonal antibodies (ab) IgG anti-human DR D1 (cod. 324390), DR D3 (cod. 324402) and DR D5 (cod. 324408) were from Calbiochem-Inalco, Italy. Rabbit polyclonal ab anti-human DR D2 (cod. LS-C22924) and DR D4 (cod. LS-C22938) were from LifeSpan-Space Import Exp, Italy. Goat anti-rabbit IgG conjugated with phycoerythrin (PE) was obtained from R&D System, Space Import Exp, Italy. PerCPCy5.5-conjugated mouse IgG anti-human CD3 (CD3 PerCPCy5.5) (cod. 317336, clone OKT3; mouse IgG2a, k) and FITC-conjugated mouse IgG anti-human CD45RA (CD45RA FITC, cod. 304106, clone HI100; Mouse IgG2b, k) were obtained from Biolegend–Campoverde, Italy. APC-Cy7-conjugated mouse IgG anti-human CD4 (CD4 APCCy7, cod. 557871, clone RPA-T4; mouse IgG1, k), Alexa Fluor 647-conjugated rat IgG anti-human CCR7 (CD197) (CCR7 AF647, cod. 557736, clone 3D12; rat IgG2a, k), PE-Cy7-conjugated mouse IgG anti-human CD8 (CD8 PECy7, cod. 557746, clone RPA-T8; Mouse IgG1, k), APC conjugated mouse IgG anti human CD3 (CD3-APC, cod. 555342, clone HIT3a) and PE Cy7 conjugated mouse IgG anti-human CD25 (CD25 PECy7, cod. 557742, clone 2A3) were all from Becton Dickinson, Italy. Purified mouse ab anti-human CD3 (cod. 555330, clone UCHT1) and purified mouse ab anti-human CD28 (cod. 555726, clone CD28.2) were obtained from Becton Dickinson, Italy. Bovine serum albumin (BSA) and 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) were purchased from Sigma, Italy. RPMI 1640, heat-inactivated fetal bovine serum (FBS), glutamine, and penicillin/streptomycin were obtained from Euroclone, Italy. Ficoll-Paque Plus was from Pharmacia Biotech (Uppsala, Sweden).

Subjects

Peripheral venous blood samples were collected from healthy volunteers (n = 13; age [mean±SD]: 60.4 ± 13.2 years) between 8:00 a.m. and 10:00 a.m. and placed in universal tubes containing preservative-free heparin (215 IU/ml). Complete blood counts of the subjects included in the study were all within normal limits, as shown in Table 1.

Table 1 Complete blood count of the subjects included in the study
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The present study is part of a project aimed at assessing the pattern of expression and the functional role of DR on circulating lymphocytes in healthy subjects and in patients with Parkinson’s disease and the study protocol was approved by the Ethics Committee of the Ospedale di Circolo and Fondazione Macchi, Varese (I).

DR Staining in Whole Blood

Samples of 100 μL of whole blood were added with 3 mL of a lysis buffer containing (g/L) NH4Cl 8.248, KHCO3 1.0 and EDTA 0.0368 in order to remove the erythrocytes. Incubation was performed at room temperature (RT) for 5 min, during which samples were gently vortexed. Samples were then centrifuged at 600 g for 5 min at RT, supernatants were removed and cells were washed one time in 1 mL of PBS (pH 7.4) supplemented with 1 % BSA (PBS/BSA) and finally resuspended in 50 μL PBS/BSA. Total leukocytes were counted by means of a hemocytometer and cell viability, determined by the trypan blue exclusion test, was always >99 %.

From each subject 7 aliquots of 100 μL were prepared: 5 were used for DR staining, 1 was used as control for the secondary PE-goat anti-rabbit (PEGAR) ab, and 1 was used as negative control (no ab). The staining protocol consisted of two steps. During the first step each aliquot was stained for one of the five DR by an indirect labelling procedure (primary ab+secondary ab labelled with PE). During the second step all the aliquots were incubated with a cocktail of anti-human CD3, CD4, CD45RA and CCR7 ab for the identification of T lymphocytes, CD4+ T lymphocytes and the following CD4+ T lymphocyte subsets: naive (CD3+CD4+CD45RA+CCR7+), central memory (TCM, CD3+CD4+CD45RA-CCR7+), and effector memory (TEM, CD3+CD4+CD45RA-CCR7-).

In the first step, each aliquot was subjected to indirect immune fluorescence staining by using rabbit polyclonal IgG directed against each of the human D1-like (D1 and D5) and D2-like (D3, D4, and D5) DR. Each sample was added with the primary anti-DR antibody (final dilution 1:100) and incubated for 30 min on ice in the dark. Cells were then washed one time with PBS/BSA at 600 g for 5 min at RT, and finally resuspended in 200 μl of PBS/BSA added with 10 μl of PEGAR ab, incubated for 30 min on ice in the dark and finally washed (600 g for 5 min at RT) and resuspended in 50 μL PBS/BSA.

In the second step, a cocktail of the following ab was added to each aliquot, according to manufactures recommendations: pre-titrated PerCPCy5.5-conjugated mouse anti-human CD3, FITC-conjugated mouse anti-human CD45RA, APC-Cy7-conjugated mouse anti-human CD4, and AlexaFluor 647-conjugated mouse anti-human CCR7. Aliquots were incubated for 30 min in dark at RT, washed with 1 mL of PBS/BSA (600 g for 5 min at RT) and finally resuspended in at least 300 μL PBS and kept on ice until analysis.

Acquisition was then performed on a BD FACSCanto II flow cytometer (Becton Dickinson Italy, Milan, Italy) with BD FACSDiva software (version 6.1.3). Lymphocytes were identified by their classical forward scatter (FSC) and side scatter (SSC) signals and a minimum of 20.000 lymphocytes from each sample were collected in the gate. Data were analyzed with FlowJo software (version 8.3.2). The results were finally expressed as percentage of positive cells (%).

PBMC Isolation

PBMC were isolated from 20 mL whole blood samples by using Ficoll-Paque Plus density gradient centrifugation. Cells were resuspended and, if necessary, any residual contaminating erythrocytes were lysed by addition of 5 mL of lysis buffer, followed by incubation for 5 min, during which samples were gently vortexed, and centrifugation at 100 g for 10 min at RT.

Cells were washed twice in PBS by addition of 15 ml of PBS and centrifugation at 300 g and 10 min at RT, and resuspended at the final concentration of 10x106 cells in 10 ml of RPMI/10 % FBS for subsequent culture. Typical PBMC preparations contained at least 80 % lymphocytes, as assessed by flow cytometry. Cell viability, assessed by the trypan blue exclusion test, was always >99 %.

PBMC Culture and Staining for DR

PBMC were cultured in RPMI/10 % FBS for 48 h at 37 °C in a moist atmosphere of 5 % CO2. Anti-CD3/anti-CD28 ab (0.01–0.1 μg/mL) were added at the beginning of cell culture. Cells were finally collected, washed twice with 15 ml of PBS/BSA, centrifuged at 300 g for 10 min at RT and finally resuspended in at least 1 mL PBS/BSA for DR staining.

From each sample, aliquots of 100 μL were prepared: 5 were used for staining of each DR and one was used as negative control. DR staining was performed as previously detailed for the first step of DR staining in whole blood. The second step consisted in incubation with mouse anti-human CD3-APC and mouse anti-humanCD4-APC-Cy7, added according to the manufacturer’s recommendations for 30 min at RT in dark. After incubation and washing with PBS/BSA at 1,400 g and 5 min at RT, cells were resuspended in 300 μl of PBS/BSA and kept in ice until flow cytometry analysis.

In each experiment, a sample of 1 × 106 PBMC was labelled with mouse ab anti-human CD3-APC, mouse ab anti-human CD4-APC-Cy7 and mouse ab anti-human CD25-PECy7, and CD25 expression was assessed. Acquisition was performed as previously described for whole blood.

Staining of DR on Viable and Apoptotic Cells

In some experiments, the expression of DR on viable and apoptotic CD3+CD4+ T lymphocytes was also assessed. To this end, PBMC were cultured alone and with anti-CD3/anti-CD28 ab 0.1 μg/mL and thereafter stained for DR as above described. In the second step pellets were also added with 5 μL of 7-aminoactinomycin (7-AAD) and incubated for 5 min on ice in the dark. After incubation, 150 μl of PBS/BSA was added and then samples were immediately analysed. Viable, early apoptotic (EA) and late apoptotic (LA) CD3+CD4+ T cells were identified by 3-color flow cytometric analysis according to an established method (Lecoeur et al. 1997).

Statistical Analysis

Data are shown as means±standard error of the mean (SEM) unless otherwise stated. Statistical significance of the differences among values was assessed by repeated measures ANOVA followed by Bonferroni’s post test or by Friedman’s post test, for normal and non-normal distributed data respectively, or by two-tailed Student’s t test, as appropriate. Correlations between DR expression and CD25 expression were assessed by Spearman correlation analysis. Calculations were performed using commercial software (GraphPad Prism version 5.00 for Windows, GraphPad Software, San Diego California USA, www.graphpad.com).

Results

Expression of DR in CD4+ T Lymphocytes in Whole Blood

CD3+CD4+ T lymphocytes expressing DR were identified by means of the following gating strategy: first, lymphocytes were gated on the scatter dot plot in the typical region of low side scatter (SSC) and forward scatter (FSC), CD3+CD4+ double positive cells were then identified among gated lymphocytes, and DR+cells were finally enumerated (Fig. 1, panel a).

Fig. 1
figure 1

Expression of DR on CD3+CD4+ T cells. Panel a: Gating strategy used to identify DR+CD3+CD4+ T lymphocytes. Total lymphocytes were identified on a biparametric dot plot SSC vs FSC (left, region 1 [R1]), to exclude monocytes, granulocytes and debris. From the cells in R1, CD3+CD4+ T cells were selected by using the CD3 vs CD4 dot plot (middle, R2). Finally, DR expression was assessed by a single-parameter histogram (right), where the dark line represents DR+ cells and the light line represents the negative control (cells stained with only the secondary PEGAR ab). Panel b: Comparison of the expression of D1-like and D2-like DR on CD3+CD4+ T cells. * = P < 0.001 vs DR D3 and P < 0.05 vs DR D4; # = P < 0.001 vs DR D2, P < 0.01 vs DR D3, P < 0.05 vs DR D4; § = P < 0.05 vs DR D2

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CD3+CD4+ T lymphocytes expressed all the five DR (Fig. 1, panel b), which were identified in cells from all the 15 subjects included in the study, however with remarkable interindividual variability: the D1-like DR D1 were expressed by 2.5–22.8 % of CD3+CD4+ T cells and D5 by 2.0–20.9 %, while the D2-like DR D2 by 1.1–7.9 %, D3 by 1.9–15.0 %, and D4 by 0.8–17.0 %. D1-like DR D1 and D5 frequencies were significantly higher than those of the D2-like DR D3 and D4. DR D5 frequency was also significantly higher than that of DR D2. Among D1-like receptors, DR D1 and D5 frequencies were not significantly different, while among D2-like receptors, the frequency of DR D4 was significantly higher than that of DR D2 (Fig. 1, panel b).

Expression of DR in CD4+ T Naive, TCM and TEM Lymphocytes

To identify CD4+ T cell subsets, the gating strategy for the selection of CD3+CD4+ T cells was followed by an additional step, by use of a biparametric dot plot CD45RA vs CCR7 which allowed the identification of T naive (CD3+CD4+CD45RA+CCR7+), TCM (CD3+CD4+CD45RA-CCR7+), and TEM (CD3+CD4+CD45RA-CCR7-) (Fig. 2, panels a–c ). The frequencies of the various subsets in the subjects included in the study were: T naive, 43.4 ± 2.7 %; TCM, 28.7 ± 1.9 %; TEM, 24.8 ± 1.6 %, respectively.

Fig. 2
figure 2

Expression of DR on CD4+ naïve/memory T cell subsets. Panels a–d: Gating strategy used to identify CD4+ T naive, TCM and TEM cells. Total lymphocytes were identified on a biparametric dot plot SSC vs FSC (R1) (a). CD3+CD4+ T cells in R1 were selected by using the CD3 vs CD4 dot plot (R2) (b). CD4+ T cell subsets in R2 were identified by a biparametric dot plot CD45RA vs CCR7 (c). DR expression was assessed by a single-parameter histogram, where dark lines represent DR+ cells and light lines represent negative controls (cells stained with only secondary PEGAR ab) (d). Panels show the analysis of DR D5 in a representative sample. Panels e–f: comparison of the expression of individual DR among naïve T cells, TCM and TEM cells (e) and among DR subtypes in T naïve, TCM and TEM cells (f). * = P < 0.05; ** = P < 0.01

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DR+ cells were finally enumerated in T naive, TCM, and TEM cells (Fig. 2, panel d), where all the five DR were expressed, however to a significantly different extent (Fig. 2, panels e–f). In T naive cells, DR D1 expression was significantly higher than in TCM, while DR D2 and D4 expression was significantly lower than in TCM and in TEM (need to see graphs and significances again) . Although the expression of all DR in TEM was slightly higher than in TCM (in particular for the D1-like DR D1 and D5 and for the D2-like D2), the differences never reached the statistical significance (Fig. 2, panel e).

In T naive cells, the D1-like DR D1 and D5 were significantly more expressed than the D2-like DR D2 D3 and D4, while in TCM and in TEM DR D5 was significantly more expressed than DR D3 (significances on graphs). D2-like DR expression did not significantly differ in neither T naive, nor TCM or TEM (Fig. 2, panel f).

DR Expression in Cultured CD4+ T Lymphocytes

In comparison to CD4+ T cells in whole blood, in CD4+ T cells cultured for 48 h the frequency of DR D5 was significantly lower (7.7 ± 0.8 % vs 13.1 ± 1.2 %, P < 0.01) and the frequency of DR D2 was significantly higher (12.5 ± 1.0 % vs 3.1 ± 0.5 %, P < 0.01). No significant differences were observed in the frequencies of DR D1 (10.5 ± 1.1 % vs 11.6 ± 1.4 %, P = 0.523), DR D3 (6.5 ± 0.6 % vs 6.6 ± 1.0 %, P = 0.909), and DR D4 (10.8 ± 1.0 % vs 8.1 ± 1.6 %, P = 0.137).

Stimulation with anti-CD3/anti-CD28 ab (0.01–0.1 μg/mL) resulted in a significant increase of CD25 expression on CD3+CD4+ T cells, from 26.2 ± 3.4 % in resting cells to 78.7 ± 6.3 % with 0.01 μg/mL and 67.1 ± 1.2 % with 0.1 μg/mL (in both cases, P < 0.01 vs resting cells). Expression of DR however significantly increased only after stimulation with anti-CD3/anti-CD28 ab 0.1 μg/mL (Fig. 3, panel a). All the five DR significantly increased on average by 55.3 %–97.4 % over resting values.

Fig. 3
figure 3

Expression of DR on CD4+ T cells cultured alone and in the presence of soluble antiCD3/anti-CD28 ab. Panel a: Expression of DR in resting cells and in the presence of antiCD3/antiCD28 ab at the concentration of 0.01 or 0.1 μg/ml. * = P < 0.01 vs resting cells. Panel b–c: Expression of DR on viable, early apoptotic (EA) and late apoptotic (LA) CD4+ T cells cultured in resting conditions (open columns) and in the presence of soluble antiCD3/anti-CD28 ab 0.1 μg/ml (dashed columns). * = P < 0.01 vs resting; # = P < 0.01 vs viable cells; § = P < 0.01 vs EA cells

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There was a significant correlation between CD25 expression and DR D5 expression in stimulated cells (r = 0.762, P = 0.037), but not in resting cells. Expression of CD25 did not show any other correlations with DR, either in resting or in stimulated cells.

After stimulation with anti-CD3/anti-CD28 ab 0.1 μg/mL, the percentage of viable cells significantly increased (76.1 ± 2.7 % vs 85.6 ± 2.8 %, P = 0.027), while that of EA and LA cells did not change (EA: 11.6 ± 2.6 % vs 10.4 ± 2.4 %, P = 0.742; LA: 0.8 ± 0.3 % vs 1.7 ± 0.5 %, P = 0.117). The frequency of all DR was significantly higher in EA and LA cells in comparison to viable cells, ad in particular the frequency of D1-like DR D1 and D5 was also significantly higher in LA cells in comparison to EA cells, however stimulation with anti-CD3/anti-CD28 ab significantly increased the expression of all DR on viable cells, without affecting DR expression on EA or LA cells (Fig. 3, panel b and c).

Discussion

The presence of DR on human lymphocytes has been a controversial issue up to less than two decades ago, mainly due to the inherent limitations of radioligand binding techniques (Vile and Strange 1995). The use of DR subtype-specific antibodies was first attempted in immunocytochemical procedures on human peripheral blood lymphocytes, leading to the identification of the D2-like DR D3 and D4 but not of DR D2 (Amenta et al. 1999), and of the D1-like DR D5 but not of DR D1 (Ricci et al. 1999). Human lymphocytes however are a highly heterogeneous population composed of several subsets of cells, each with similar morphology but profoundly different molecular phenotype and functions. As a consequence, very high numbers of cells must be examined, and their phenotype determined, to obtain meaningful results. In this regard, flow cytometry is a powerful technique providing rapid analysis of multiple characteristics of single cells. Flow cytometry allows the phenotyping of any particles flowing in a stream of fluid, by use of both light scattering at different angles (which identifies differences in size and internal complexity), and light emitted from fluorescently labelled antibodies (which can be used to identify both cell surface and cytoplasmic antigens). Flow cytometry has been used for the first time to identify DR on human lymphocytes by McKenna et al. (2002), who documented DR expression on human T- and B-lymphocytes, monocytes, neutrophils, eosinophils and NK cells. Subsequently, by use of flow cytometry Besser et al. (2005) identified D2-like DR on human T cells, and Brito-Melo et al. (2012) analyzed DR D2 and D4 on CD4+ and CD8+ T lymphocytes.

In the present study, we used flow cytometry to characterize the expression of all the five DR on human circulating CD4+ T cells and to assess differences in DR expression in the various populations of CD4+ memory T lymphocytes, which represent distinct subsets each with peculiar homing capacity and effector functions (Sallusto et al. 1999; Sallusto et al. 2004). According to our results, CD4+ T cells always expressed all the five DR: D1-like DR were identified on average on 11.6–13.1 % of CD4+ T cells, while D2-like DR were less expressed, being present on only 3.1–8.1 % of the cells. Remarkably, the expression of all DR showed high interindividual variability, ranging from 1–2 % up to 8–17 % for D2-like DR and from 1–2 % up to 21–23 % for D1-like DR. These findings differ from those reported by McKenna et al. (2002), who found DR in T lymphocytes in only 3–5 out of 19 individuals and with expression frequencies in the average range 1–5 %. Brito-Melo et al. (2012) however reported DR D2 on 2–10 % and DR D4 on 2–16 % of CD4+ T cells. McKenna et al. (2002) performed their assays on cells obtained by density gradient centrifugation and investigated DR in the whole CD3+ T lymphocyte population. On the contrary, Brito-Melo et al. (2012) stained DR in whole blood and performed erythrocyte lysis only thereafter, identifying DR in CD4+ (and CD8+) T cells, which is the same procedure which we adopted in our study. In addition, Brito-Melo et al. (2012) analyzed at least 10,000 cells per sample, which is about the same order of magnitude which we used (20,000 cells per samples). No information on the number of cells analyzed per sample is provided by McKenna et al. (2002).

The relative prevalence of D1-like over D2-like DR in CD4+ T cells may imply that the effects of DA on these cells are mediated mainly through D1-like DR. Most effects of DA on T cells however have been reported to be dependent upon D2-like DR activation, including β1 integrin-dependent adhesion to fibronectin (Levite et al. 2001), increased production of TNF-α and IL-10 (Besser et al. 2005), and inhibition of the proliferation of activated T cells (Sarkar et al. 2006), although at least production of TNF-α and IL-10 was shown to be affected also by D1-like DR (Besser et al. 2005). All those studies however were performed on whole population of T cells, therefore the relative contribution to the observed responses of CD4+ and CD8+ T cells remains to be established. Moreover, the few studies performed on CD4+ T cells suggest that the direct effects of DA on these cells are likely mediated mainly by D1-like DR. Indeed, we previously reported that DA-mediated functional inhibition of human CD4+CD25high T regulatory lymphocytes (including IL-10 and TGF-β production and suppression of CD4+CD25- T effector cell proliferation) is mediated by D1-like DR possibly of the D5 subtype (Cosentino et al. 2007), and more recently Nakano et al. (2009) showed that DA released by dendritic cells upon interaction with naive CD4+ T cells acts on D1-like DR to shift T-cell differentiation towards the Th2 lineage, thus resulting in increased GATA-3 expression and IL-4 and IL-5 production.

To our best knowledge, this is the first study extensively reporting the expression of all the five DR on human CD4+ naive and memory T lymphocyte subsets. Memory is indeed a peculiar feature of the acquired immune system, and in particular in the T cell compartment presentation of novel antigens by dendritic cells to naive T cells triggers a primary immune response leading to the clonal expansion of both “effector” cells, which immediately fight the foreign pathogen, and “memory” cells, which may persist for the whole life and subserve later recall responses in secondary lymphoid organs. According to the established model (Lanzavecchia and Sallusto 2000; Sallusto et al. 2004), TCM mediate reactive memory, by homing to T cell areas of secondary lymphoid organs and readily proliferating and differentiating to effector cells upon antigenic stimulation, while TEM afford protective memory, by migrating to inflamed peripheral tissues and displaying immediate effector function. According to our results, human CD4+ naive T cells, TCM, and TEM cells express all the five DR, however with distinct expression patterns: naive T cells express more D1-like than D2-like DR, which on the contrary are increased in TCM and TEM cells. It was previously suggested that D1-like DR could be more expressed than D2-like DR in human CD4+ naive T cells (Nakano et al. 2008). The present study now provides definite evidence for the prevalence of D1-like over D2-like DR in CD4+ naive T cells, additionally showing, in CD4+ TCM and TEM cells, upregulation of D2-like DR (in particular DR D2 and D4 in both TCM and TEM) and downregulation of D1-like DR (in particular DR D1 in TCM). Preferential expression of D1-like DR in naive T cells is in agreement with the proposed role of DA during dendritic cell-induced naive T cell stimulation (Nakano et al. 2009), while increased D2-like DR on TCM and TEM cells suggests a prominent role of such receptors in differentiated memory cells. Stimulation of D2-like DR on human T cells has been suggested to induce increased integrin-dependent trafficking and extravasation of T cells in periphery (Levite et al. 2001). Interestingly, naive T cells migrate almost exclusively through lymphoid tissue by means of L-selectin-dependent mechanisms, while memory T cells, particularly TEM, migrate mainly to peripheral tissues, to skin and to mucosal surfaces, to provide protection at sites vulnerable to pathogens, and their migration depends primarily on the involvement of β-integrins (Sallusto et al. 2004). It could be therefore speculated that D2-like DR expression in TCM and TEM cells contributes to the peculiar functions of these cells and warrants additional studies to assess the potential of D2-like DR as targets for the pharmacological manipulation of memory functions in CD4+ T cells.

The occurrence of DR on naive and memory CD4+ T cell subsets is a remarkable finding also in view of emerging evidence about the relationship between peripheral immune cells and neuroinflammation and neurodegeneration in PD (Laurie et al. 2007; Reynolds et al. 2008; Stone et al. 2009; Saunders et al. 2012; Sanchez-Guajardo et al. 2013). In peripheral blood of PD patients, decreased numbers of total lymphocytes, decreased CD4+ T cells, and reduced naïve and increased memory CD4+ T cells have been consistently reported (Fiszer et al. 1994; Bas et al. 2001; Calopa et al. 2010; Saunders et al. 2012), and recently such alterations have been associated with motor function scores and disease severity, supporting a role of chronic immune activation in PD progression (Saunders et al. 2012). DR expression on naive and memory CD4+ T cells now implies the possibility that dopaminergic agents, which are first line treatments in PD, may act not only by replacing DA deficiency in the CNS but also by directly affecting peripheral immune responses associated with disease progression. We recently reported preliminary evidence regarding the involvement of DR expressed on circulating lymphocytes in PD by showing that CD4+ T cells obtained from patients consistently show reduced levels of all DR in comparison to cells obtained from healthy controls (Kustrimovic et al. 2013). Such results were obtained in a small group of subjects and are now being replicated in a larger cohort. In any case, the possibility that the immune effects of dopaminergic agents contribute to their overall therapeutic efficacy in PD deserves careful consideration.

In the present study we also examined DR expression in human cultured CD4+ T cells with particular regard to the effects of stimulation with anti-CD3/anti-CD28 ab, and three main findings arise from this set of experiments. The first one is that, in comparison to CD4+ T cells in whole blood, in CD4+ T cells in culture there is reduced expression of DR D5 (on average, from about 13 to 6 %) and increased expression of DR D2 (on average, from 3 % to more than 10 %). To what extent such modifications may impact on functional responses cannot be established on the basis of our data, however future studies should take into account the possibility that in vitro conditions affect the results of experiments aimed at investigating the relative contribution of different DR subtypes to the functional responses induced by DA in human CD4+ T cells. The second finding is that stimulation with anti-CD3/anti-CD28 ab increased the expression of all DR, by 71–84 % in the case of D1-like DR and by 55–97 %. in the case of D2-like DR. Remarkably, this increase was seen only with high concentrations of anti-CD3/anti-CD28 ab, while increased expression of CD25, i.e. the α-chain of the IL-2 receptor (which is critical for IL-2-sustained proliferation of T cells), occurred already at low concentrations of anti-CD3/anti-CD28 ab, possibly implying that DR and CD25 expression on CD4+ T cells differ in their sensitivity to stimulation. Interestingly, it has been reported that different conditions of antigen persistence and antigen load may lead to distinct phenotypic patterns of CD4+ memory T cell responses (Harari et al. 2004). Investigating the relationship between conditions of antigenic stimulation and DR expression on CD4+ T cells will contribute to clarify the contribution of DR to the functional modulation of the various memory T cell subsets. Finally, and possibily most interestingly, the frequency of DR was higher in apoptotic cells in comparison to viable cells, and stimulation increased all DR on viable cells, without affecting their expression on apoptotic cells. We previously reported that in human PBMC DA decreases oxidative metabolism and apoptosis, possibly through D1-like DR-dependent mechanisms (Cosentino et al. 2004). The present findings further support the involvement of dopaminergic pathways in the regulation of viability and apoptosis in human lymphocytes. In addition, stimulation-induced upregulation of DR selectively occurring in viable cells suggests the preferential contribution of DA in the regulation of lymphocyte functions in activated cells. As a whole, it can be temptatively proposed that in resting cells dopaminergic pathways participate mainly in apoptotic processes (as suggested by the high proportion of apoptotic cells expressing DR), while their functional relevance increases in activated cells (in line with stimulation-induced upregulation of DR in viable cells). Results further emphasize the need to investigate carefully the possible immune effects of dopaminergic agents already in clinical use, since any positive findings would readily provide the proof-of-concept to assess their therapeutic potential as novel immunomodulating agents.

In conclusion, evidence has been provided that human CD4+ T lymphocytes express all the five subtypes of DR, however with distinct patterns in the different subsets. While D1-like DR are preferentially expressed in naive T cells, D2-like DR expression is higher in TCM and in TEM cells, possibly in agreement with available evidence showing an involvement of D1-like DR in dendritic cell-induced differentiation of naive T cells (Nakano et al. 2009) and of D2-like DR in integrin-dependent trafficking of T cells to peripheral tissues (Levite et al. 2001), which is a peculiar function of memory T cells (in particular of TEM). Moreover, high expression of DR in apoptotic cells and stimulation-induced DR upregulation in cultured CD4+ T cells suggests the involvement of DR in the apoptotic process and further supports the involvement of DR in the functional regulation of activated cells, requiring further investigations to assess the role of DR subtypes in the modulation of specific responses. Several issues nonetheless still await clarification. For instance, it was not possible to establish whether different DR subtypes are coexpressed on the same cells, and there is therefore considerable room for improvement also from the technical point of view. In any case, convincing evidence has been increasingly provided in recent years concerning the functional relevance of dopaminergic pathways in human T cells. Unravelling their complexity will clarify the relevance of any direct effects on peripheral immunity in the therapeutic activity of dopaminergic agents currently used in PD, eventually paving the way to the repurposing of these agents as novel immunomodulating drugs.