The frequency of CD127low expressing CD4+CD25high T regulatory cells is inversely correlated with human T lymphotrophic virus type-1 (HTLV-1) proviral load in HTLV-1-infection and HTLV-1-associated myelopathy/tropical spastic paraparesis
- Jakob Michaëlsson†1, 5,
- Hugo Marcelo R Barbosa†2,
- Kimberley A Jordan1,
- Joan M Chapman1,
- Milena KC Brunialti2,
- Walter Kleine Neto3,
- Youko Nukui3,
- Ester C Sabino3,
- Marco Antonio Chieia2,
- Acary Souza Bulle Oliveira2,
- Douglas F Nixon1 and
- Esper G Kallas2, 4Email author
© Michaëlsson et al; licensee BioMed Central Ltd. 2008
Received: 25 June 2008
Accepted: 29 July 2008
Published: 29 July 2008
CD4+CD25high regulatory T (TReg) cells modulate antigen-specific T cell responses, and can suppress anti-viral immunity. In HTLV-1 infection, a selective decrease in the function of TReg cell mediated HTLV-1-tax inhibition of FOXP3 expression has been described. The purpose of this study was to assess the frequency and phenotype of TReg cells in HTLV-1 asymptomatic carriers and in HTLV-1-associated neurological disease (HAM/TSP) patients, and to correlate with measures of T cell activation.
We were able to confirm that HTLV-I drives activation, spontaneous IFNγ production, and proliferation of CD4+ T cells. We also observed a significantly lower proportion of CTLA-4+ TReg cells (CD4+CD25high T cells) in subjects with HAM/TSP patients compared to healthy controls. Ki-67 expression was negatively correlated to the frequency of CTLA-4+ TReg cells in HAM/TSP only, although Ki-67 expression was inversely correlated with the percentage of CD127low TReg cells in healthy control subjects. Finally, the proportion of CD127low TReg cells correlated inversely with HTLV-1 proviral load.
Taken together, the results suggest that TReg cells may be subverted in HAM/TSP patients, which could explain the marked cellular activation, spontaneous cytokine production, and proliferation of CD4+ T cells, in particular those expressing the CD25highCD127low phenotype. TReg cells represent a potential target for therapeutic intervention for patients with HTLV-1-related neurological diseases.
Between 10 and 20 million people are infected with HTLV-1 worldwide . Although most subjects are clinically asymptomatic during their lifetime, a proportion (5 to 10%) develop adult T cell leukemia/lymphoma (ATLL) or HTLV-1 associated myelopathy/tropical spastic paraparesis (HAM/TSP) . Epidemiological surveys have identified regions in the world where prevalence rates are considerably higher, including Japan, the Caribbean, South America, Africa, Melanesia and the Middle East [1, 3]. It has been estimated that the prevalence of HTLV-1 infection in South America ranges from 2 to 5% , with an estimated 1–2 million infected people in Brazil . The prevalence in blood donors ranges from 0.17 to 1.8% in different areas of the country [6, 7], with a 0.3% seroprevalence in the city of Sao Paulo blood donors .
HTVL-1 is a retrovirus encoding the group specific antigen (gag), protease (pro), polymerase (pol), and envelope (env) genes. Six proteins are encoded by the pX region of the genome, including the Tax protein, which is critical to viral replication and induction of cellular activation and transformation, increasing the expression and production of cytokines and receptors involved in T cell growth and transformation, such as IL-15 [9, 10] and IL-2 [11–13]. Tax also has the ability of interfering in the expression of several transcription factors and proto-oncogenes, as well as in the nucleic acid repair and apoptosis [14–17]. These effects combined seem to play a key role in the potential of HTLV-1 to induce cellular transformation and, consequently, trigger the development of ATLL.
It has been previously demonstrated that HTLV-1 proviral load is one of the key factors in the pathogenesis of HAM/TSP [18, 19], although host genetic factors are also independently associated with the development of the diseases, e.g. certain HLA [20, 21] and non-HLA [22, 23] genes. These invoke the hypothesis that both viral and genetic host factors are implicated in the pathogenesis of HAM/TSP.
The CD8+ T cell response to HTLV-1 can be readily detected [24–31], commonly directed against the HTLV-1-tax protein. The contribution of the CD8+ T cell response might be particularly important for viral control in HTLV-1 infection, since infected lymphocytes produce virtually no cell-free infectious HTLV-1 particles. However, it is noteworthy that the magnitude of the HTLV-1-specific T cell response is associated with higher proviral loads, highlighting the fact that T cells frequencies are determined by proviral load, as well as being a determinant of proviral load. CD4+ T cells are the main target for HTLV-1 infection, which induces CD4+ T cell activation, including proliferation and IFNγ production. The HTLV-1-specific CD4+ T cell response is directed mainly against Env, the HTLV-1 envelope surface .
TReg cells are crucial for the control of autoimmune disease and maintenance of peripheral T cell tolerance (reviewed in Sakagushi et al. ). In addition, they can suppress pathogen-specific T cell responses, including response to viruses [34–37]. The mechanisms whereby TReg cells suppress T cell responses are not yet fully understood, but are likely to include both soluble factors, e.g. IL-10 and TGF-β, as well as cell-cell contact dependent mechanisms, e.g. through CTLA-4. CTLA-4 (CD152) is expressed by a large fraction of CD4+CD25+ T cells, and by a majority of CD4+CD25high T cells. CTLA-4 has also been shown to be one of mediators of TReg function [38, 39], and is considered a marker for TReg cells. In addition, it was recently demonstrated that TReg cells are characterized by low levels of the IL-7Rα (CD127low) [40–42], which together with CD25 help to distinguish TReg cells from activated normal CD4+ T cells in healthy individuals. FOXP3 is a key regulator of TReg cell function, but is not exclusive to TReg cells; it has been identified in human nonregulatory activated CD4+FoxP3+ T cells. Humans with mutations in FOXP3 present with a syndrome characterized by severe autoimmune and inflammatory disorders often early in life, denominated IPEX . Interestingly, it was recently shown that HTLV-1 tax can downregulate Foxp3 expression [43, 44].
We hypothesized that HTLV-1 compromises TReg cell function, resulting in higher T cell activation, which contributes to HAM/TSP development. We found a significantly higher frequency of CD4+Ki-67+ T cells and a lower proportion of CTLA-4+ TReg cells in subjects with HAM/TSP, compared to healthy controls. Moreover, we found an inverse correlation between HTLV-1 proviral load and frequency of CD127low/CTLA-4+TReg cells. Our data suggest a role for TReg cells in the pathogenesis of HAM/TSP, and reveal a potential new therapeutic target for patients with HAM/TSP.
Characteristics of study subjects.
Proviral Load (copies/1,000 cells)
CD4+ T cells/μl
CD8+ T cells/μl
CD3+ T cells/μl
CD4+ T cell activation and IFNγ production in healthy donors, HTLV-1 seropositive asymptomatics, and HAM/TSP patients
Decreased frequency of TReg cells in HAM/TSP patients
Decreased CTLA-4+ TReg cells correlate with increased CD4+ T cell proliferation in HAM/TSP patients
The frequency of CD4+CD25+CTLA-4+ and CD127low TReg cells was negatively correlated to HTLV-1 proviral load
Regulatory T cells are important for the maintenance of peripheral T cell tolerance to self antigens, and can also suppress T cell responses to tumors, parasites, viruses and bacteria. In this study we addressed the relationship between TReg cells, T cell activation, and HTLV-1 proviral load. Infection with HTLV-1 was associated with higher spontaneous IFNγ release by CD4+ T cells, but only in HAM/TSP there was a marked increase in T cell proliferation.
The HTLV-1 derived tax protein can downregulate expression of the FOXP3, which presence is associated with TReg cell function [43, 44]. Increased expression of tax can be expected in patients with HAM/TSP, who have higher proviral loads compared to asymptomatic carries. We observed a higher proportion of CD4+ IFNγ+ T cells in HTLV-1 infected subjects, which could also be indicative of a decreased TReg cell fraction. Interestingly, only the HAM/TSP patients presented with a higher cell proliferation, as measured by Ki-67 staining, which correlated markedly with HTLV-1 proviral load (data not shown). These observations suggest that HTLV-1 directly affects TReg cell number, and as proviral load increases, not only is the control of IFNγ lost, but controls on cell proliferation as well. Our data, together with the recent findings that HTLV-1 tax downregulates FOXP3 expression, indicate that TReg cell dysfunction can be a direct consequence of HTLV-1 infection.
In order to better understand the role of TReg cells in HTLV-I infection and disease, we used CTLA-4 and CD127 staining in CD4+CD25high cells as markers for TReg subsets. CD127 and CTLA-4 have been described as useful markers for TReg, and facilitate the identification of TReg cells, even without staining for FOXP3 . In this study, we found that an increased frequency of CD127lowCD4+CD25+ TReg in controls correlated negatively with CD4+ T cell proliferation (Ki-67), indicating that these cells indeed have a regulatory immunophenotype. In contrast, increased frequency of these cells correlated with increased CD4+ T cell proliferation in HTLV-1 infected individuals, suggesting that these cells are not regulatory T cells in these individuals. In addiction, the elevated frequency of CTLA-4+ TReg cells was negatively correlated to CD4+ T cell proliferation only in HAM/TSP patients, which suggest that it is a better immunophenotype of TReg cells in HAM/TSP patients, but more studies are necessary to confirm this.
We could detect a negative association between the frequency of CTLA-4+ or CD127low TReg cells and proviral load, extending recent findings of an association between FOXP3 expression and HTLV-1 infection . We speculate that therapeutic manipulation of regulatory T cells could positively impact disease pathogenesis. Two mechanisms might be involved, the first by suppressing the exuberant anti-HTLV-1 CD8+ T cell mediated immune response, and the second by suppression of CD4+ T cell proliferation, which can result in lower proviral load. However, stimulating an expansion of TReg cells could also provide additional targets for HTLV-1 replication, so such studies should proceed with great caution.
In this study, there are some limitations. The study was cross sectional, and with a limited number of patients in each group. We hope that future longitudinal studies can assess changes in TReg cells over time in HTLV-1 infected patients. We, and others, working in the regulatory T cells field, are limited by the lack of definitive phenotypic markers of TReg, and CD4+CD25+/high remains the standard identifiers. In this study we have added other markers, but at the time the study was conducted, the FOXP3 antibody, commonly used to detect a TReg cell population was not commercially available. However, this may have been a fortuitous event, as recent reports suggest that FOXP3 is also expressed on non regulatory T cells in humans [46, 47]. As we did not have access to tissue samples from these subjects, we cannot exclude redistribution of cells out of the peripheral blood into tissues, and the study of regulatory T cells at secondary lymphoid sites and within CSF will be of interest for a future study of HTLV-1 associated disease.
In conclusion, our data suggest a role of TReg cells in the pathogenesis of HAM/TSP. Further studies should help delineate the ability of expanded TReg cells to affect T cell proliferation in HTLV-1 patients and the potential development of therapeutic modulation of regulatory T cells in HTLV-1 patients.
In this study, we showed that HTLV-I drives activation, spontaneous IFNγ production, and proliferation of CD4+ T cells. HAM/TSP patients have a decreased frequency of TReg cells in peripheral blood, compared to healthy subjects, markedly in the CD4+CD25highCTLA+ phenotype. The proportion of CD127low TReg cells correlated inversely with HTLV-1 proviral load. These results suggest that TReg cells may be subverted in HAM/TSP patients, and contributes to the identification of novel therapeutic targets for patients with HTLV-1-related disease.
Three groups of volunteers were enrolled. The first consisted of seven HTLV-1-negative control volunteers; the second consisted of ten HTLV-1 seropositive volunteers without clinical and laboratory evidence of HTLV-1-associated disease, and the last group was composed of nine patients with the diagnosis of HTLV-1 associated myelopathy/tropical spastic paraparesis (HAM/TSP). After approval by the Institutional Review Board, written informed consent was obtained from all the participants according to the guidelines of Brazilian Ministry of Health. Samples were collected in EDTA-treated vacuum tubes, and PBMC were frozen into liquid nitrogen after separation using a ficoll gradient.
DNA extraction and determination of HTLV-1 proviral load
HTLV-1 proviral DNA was extracted from PBMCs using a commercial kit (Qiagen GmbH, Hilden Germany) following the manufacturer's instructions. The extracted DNA was used as a template to amplify a fragment of 158 bp from the viral tax region using previously published primers . The SYBR green real-time PCR assay was carried out in 25 μl PCR mixture containing 10× Tris (pH 8.3; Invitrogen, Brazil), 1.5 mM MgCl2, 0.2 μM of each primer, 0.2 mM of each dNTPs, SYBR Green (18.75 Units/r × n; Cambrex Bio Science, Rockland, ME) and 1 unit of platinum Taq polymerase (Invitrogen, Brazil). The amplification was performed in the Bio-Rad iCycler iQ system using an initial denaturation step at 95°C for 2 minutes, followed by 50 cycles of 95°C for 30 seconds, 57°C for 30 seconds and 72°C for 30 seconds. The human housekeeping β globin gene primers GH20 and PC04  were used as an internal control calibrator. For each run, standard curves for the value of HTLV-1 tax were generated from MT-2 cells of log10 dilutions (from 105 to 100 copy). The threshold cycle for each clinical sample was calculated by defining the point at which the fluorescence exceeded a threshold limit. Each sample was assayed in duplicate and the mean of the two values was considered as the copy number of the sample. The amount of HTLV-1 proviral load was calculated as follows: copy number of HTLV-1 (tax) per 1,000 cells = (copy number of HTLV-1 tax)/(copy number of β globin/2) × 1000 cells. The method could detect 1 copy per 103 PBMCs cells.
PBMCs were thawed and stained with directly conjugated antibodies. Three different panels of antibodies were used to evaluate the expression of proteins associated with TReg cells and T cell activation. All antibodies were from BD Biosciences, unless otherwise noted. All panels contained PerCP-conjugated anti-CD4 and allophycocyanin-conjugated anti-CD25, and in addition contained (1) FITC-conjugated anti-GITR and PE-conjugated anti-CD127 (Beckman Coulter, Miami, FL), (2) FITC-conjugated anti-CD45RA and PE-conjugated anti-HLA-DR and (3) FITC-conjugated anti-Ki-67 and PE-conjugated anti-CD152 (CTLA-4). Cells stained with PerCP-conjugated anti-CD4 alone and allophycocyanin-conjugated CD25 alone were used to establish positive gates for FITC- and PE-conjugated antibodies. For panel 1 and 2, cells were stained with all antibodies in PBS supplemented with 0.5% bovine serum albumin (BSA) and 2 mM EDTA (FACS buffer), followed by two washes in FACS buffer and fixation in 1% paraformaldehyde (PFA). For panel 3, cells were first stained with PerCP-conjugated anti-CD4 and allophycocyanin-conjugated anti-CD25, followed by two washes in FACS buffer and fixation in 1% PFA. The cells were subsequently washed twice with PBS containing 0.1% saponin (perm buffer), prior to staining with PE-conjugated anti-CD152 and FITC-conjugated anti-Ki-67 diluted in perm buffer. All samples were analyzed on a FACSCalibur flow cytometer (Becton Dickinson) equipped with a 488 nm argon and a 633 nm red-diode lasers for four color detection. Acquisition and analyses were performed using CellQuest software (Becton Dickinson). Fluorescence voltages and compensation values were determined using unstained cells and cells single-stained with each of the fluorochrome-conjugated antibodies, respectively. The gating strategy used was to gate on lymphocytes using a forward scatter versus side scatter gate, followed by gating on CD4+ cells. The gate for CD4+CD25+ cells was set using cells cells stained with the PerCP-conjugated anti-CD4 antibody alone. Positive gates for the FITC- and PE-conjugated antibodies were set using cells stained with only PerCP-conjugated anti-CD4 and APC-conjugated anti-CD25 antibodies.
Cytokine flow cytometry
PBMCs were thawed and cultured for 24 hours in 96-well U-bottom plates at a concentration of 4 × 105 cells/well. Brefeldin A (BFA) was added at a concentration of 5 μg/ml for the last 5 hours of the culture. After culture, cells were harvested, stained with PE-conjugated anti-CD4, fixed in 4% PFA for 20 min, prior to being washed twice with perm buffer. The cells were subsequently stained with PerCP-conjugated anti-CD3 and allophycocyanin-conjugated anti-IFNγ, washed twice in perm buffer and resuspended in FACS buffer, prior to being analyzed on a FACSCalibur. All antibodies were from BD Biosciences.
Data sets were compiled and analyzed in Statistica, release 6.0 (Statsoft, Tulsa, OK) and Prism, release 4.0 (GraphPad Software, San Diego, CA). Groups comparisons were performed using non-parametric Kruskal Wallis ANOVA by ranks test; associations between variables were evaluated by Spearman rank order correlation's test. Critical p values were considered statistically significant if below 0.05.
We would like to thank Marli Campos, Rosemeire Gabriel, and Perpétua for the assistance in blood collection, and Dr. Lishomwa Ndhlovu for helpful scientific discussions. We are also grateful to Karina I. Carvalho, Helena Tomiyama, and Raphaella Goulart for the continuous laboratory support. This work was supported by the Fundação de Amparo a Pesquisa do Estado de São Paulo (FAPESP), grant #04/10918-6, NIH grants (AI41531, AI52731, AI060379, AI064520), and The Fogarty International Center, grant #D43 TW00003. Dr. Barbosa and Ms. Brunialti were supported by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES), Brazilian Ministry of Education.
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