Homeostatic proliferation leads to telomere attrition and increased PD-1 expression after autologous hematopoietic SCT for systemic sclerosis

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In the months that follow autologous hematopoietic stem cell transplantation (AHSCT), lymphopenia drives homeostatic proliferation, leading to oligoclonal expansion of residual cells. Here we evaluated how replicative senescent and exhausted cells associated with clinical outcomes of 25 systemic sclerosis (SSc) patients who underwent AHSCT. Patients were clinically monitored for skin (modified Rodnan’s skin score, mRSS) and internal organ involvement and had blood samples collected before and semiannually, until 3 years post-AHSCT, for quantification of telomere length, CD8+CD28 and PD-1+ cells, and serum cytokines. Patients were retrospectively classified as responders (n = 19) and non-responders (n = 6), according to clinical outcomes. At 6 months post-AHSCT, mRSS decreased (P < 0.001) and the pulmonary function stabilized, when compared with pre-transplant measures. In parallel, inflammatory cytokine (IL-6 and IL-1β) levels and telomere lengths decreased, whereas PD-1 expression on T-cells and the number of CD8+CD28 cells expressing CD57 and FoxP3 increased. After AHSCT, responder patients presented higher PD-1 expression on T- (P < 0.05) and B- (P < 0.01) cells, and lower TGF-β, IL-6, G-CSF (P < 0.01), and IL-1β, IL-17A, MIP-1α, and IL-12 (P < 0.05) levels than non-responders. Homeostatic proliferation after AHSCT results in transient telomere attrition and increased numbers of senescent and exhausted cells. High PD-1 expression is associated with better clinical outcomes after AHSCT.


Systemic sclerosis (SSc) is a chronic autoimmune disease characterized by microvascular damage, inflammation, and progressive fibrosis of the skin and internal organs [1]. Autologous hematopoietic stem cell transplantation (AHSCT) has been used to treat refractory severe SSc patients, aiming to renew the immune system, thereby inducing prolonged clinical response [2,3,4]. The immunological resetting that follows AHSCT is driven by thymic function rebound, leading to increased output of newly generated autotolerant clonotypes and expansion of regulatory T-cells [5,6,7]. Broadened T-cell receptor (TCR) diversity is achieved from 1 to 2 years after transplantation and correlates with thymic reactivation, leading to long-term remissions [5, 6, 8,9,10,11].

AHSCT includes high-dose immunosuppression that immediately abrogates autoimmunity. Then, at the early periods that comprehend weeks to months after transplantation, thymic-independent homeostatic proliferation mechanisms are triggered, in response to profound lymphopenia [5, 10]. Infused hematopoietic stem cells (HSCs) and residual cells that survived immunoablation or that were reinfused within the graft undergo repeated cell divisions to reconstitute a new immune system. Indeed, in multiple sclerosis (MS) patients, homeostatic proliferation resulted in replicative senescence and clonal exhaustion during the early immune reconstitution periods after AHSCT, as respectively shown by increased counts of CD57-expressing CD8+CD28 [5, 7, 12, 13] and of programmed death-1 positive (PD-1+) exhausted T-cells [12, 13]. In MS patients, high PD-1 expression on T- and B-cells shortly after transplantation has been associated with long-term disease control [13]. In this context, high rates of homeostatic proliferation constitute a barrier to transplantation tolerance [14], once residual cells undergoing intense homeostatic expansion may ultimately impair the establishment of autotolerance after AHSCT [15].

Telomere attrition has been described in inflammatory, malignant, and degenerative disorders, as a marker of biological aging [16, 17]. In autologous and allogeneic stem cell transplantation for hematological indications, oligoclonal T-cell expansions lead to clonal senescence with accelerated telomere shortening early after transplantation [18,19,20]. Shorter telomere length before transplantation is associated with higher transplant-related mortality [21]. In the setting of AHSCT for autoimmune diseases, the influence of proliferative exhaustion, clonal senescence, and telomere attrition on transplant outcomes has not been reported. Here we evaluated how telomere length, frequency of senescent and exhausted cells, and cytokine profiles associate with clinical outcomes of SSc patients treated with AHSCT.

Subjects and methods


We prospectively followed 25 patients with severe and progressive SSc, recruited from the Ribeirão Preto Medical School University Hospital (Ribeirão Preto, Brazil), who underwent AHSCT from 2010 to 2015. All patients met the 1980 American College of Rheumatology and/or the 2013 American College of Rheumatism/European League against Rheumatism classification criteria for SSc [22, 23]. Patient characteristics at enrolment are described in Table 1. The transplantation protocol, inclusion and exclusion criteria have been previously published [24]. Briefly, HSCs were mobilized with 2 g/m2 of cyclophosphamide plus granulocyte colony-stimulating factor (G-CSF) and subsequently collected by leukoapheresis. Then, patients were treated with 200 mg/kg intravenous cyclophosphamide plus 4.5 mg/kg rabbit anti-thymocyte globulin, followed by infusion of the non-manipulated autologous HSCs. Transplant protocols were approved by the institutional review boards (Ethics Committee of the University Hospital of the Ribeirão Preto Medical School and CONEP) and patients gave written consent.

Table 1 Patients characteristics before transplantation

Clinical follow-up

Patients were followed-up as previously described [8] during the first year after AHSCT and semiannually thereafter, until 36 months. The same observer assessed clinical response using repeated functional and physical examination of organ involvement. Clinical evaluations included assessment of the modified Rodnan’s skin score (mRSS), lung, kidney, gastrointestinal and heart function, as well as quantification of anti-topoisomerase I (anti-Scl-70) antibodies and C-reactive protein. Relapsing disease post-AHSCT was defined by one of the following criteria: increase of the mRSS by 25% from best improvement, or a decline in forced vital capacity (FVC) by 10%, renal crisis, start of total parenteral nutrition, or restarting of immune suppressive or modulating medication, as previously described [9, 24]. According to clinical response at 36-month follow-up after AHSCT, SSc patients were retrospectively classified as responders and non-responders. Responder patients presented improvement or stabilization of skin and lung involvement, in the absence of further immunosuppressive treatment. Conversely, patients that relapsed, according to the above-mentioned criteria, were considered non-responders.

Immune monitoring

Whole blood samples were collected for immune monitoring at baseline and every 6 months until 36 months after transplantation. EDTA-containing tubes were collected for fluorescence-activated cell sorting analysis. For sera isolation, the samples were spun after clot formation in a refrigerated centrifuge (3500 r.p.m., 15 min) and the serum was quickly frozen at − 80°C and stored until assayed. Peripheral blood mononuclear cells (PBMCs) were isolated from heparinized blood by Ficoll-Hypaque (Amersham-Pharmacia, Uppsala, Sweden) density-gradient separation. PBMCs were lysed and stored in TRIzol reagent (Invitrogen, Carlsbad, CA) at – 80 °C for DNA extraction following manufacturer’s protocol.

Flow cytometry analysis

Immunophenotyping was performed as previously published using predetermined optimal antibody concentrations and incubations [12, 13, 25]. Anti-human mAbs included the following: CD3 (UCHT1), CD4 (RPA-T4), CD8 (RPA-T8), CD19 (HIB19), CD28 (CD28.2), CD57 (NK-1), PD-1 (MIH4) (BD Pharmingen, San Diego, CA), and FoxP3 (PCH101) (eBioscience, San Diego, CA). The cells were acquired in FACSCalibur (BD Biosciences, San Jose, CA) cytometer and analyzed using Flow Jo (TreeStar, Ashland, OR) software. All analyses were performed on fresh blood. Blood cells were counted by the automated blood cell counter Mindray BC-2800 (Mindray Medical Instrumentation, China). Results are expressed as absolute cell numbers (cell/μL) according to the following formula: absolute lymphocytes (per μL) = number of total lymphocytes (per μL) × percentage of specific lymphocyte subpopulation obtained by flow cytometry (%)/100 [13].

Cytokine quantification

Platelet-derived growth factor (PDGF)-BB and monocyte chemoattractant protein (MCP)-3 serum levels were measured using specific enzyme-linked immunosorbent assays (eBioscience, Vienna, Austria), with respective detection limits of 4.6 pg/mL and 0.3 pg/mL. Interleukin (IL)-2, IL-6, IL-10, IL-12, IL-13, IL-17, IL-1β, transforming growth factor (TGF)-β1, MCP-1, MIP-1α, MIP-1β, G-CSF, and tumor necrosis factor-α (TNF-α) serum levels were determined by cytometric bead array (BD Biosciences), under respective sensitivities of 11.2, 1.6, 0.13, 0.6, 0.6, 2.9, 2.3, 14.9, 1.3, 0.2, 0.8, 1.6 and 1.2 pg/mL.

Telomere length

Telomeres were measured by monochrome multiplex quantitative PCR on genomic DNA as previously described [26]. Briefly, the telomere primer pair TELG (5′-ACACTAAGGTTTGGGTTTGGGTTTGGGTTTGGGTTAGTGT-3′) and TELC (5′-TGTTAGGTATCCCTATCCCTATCCCTATCCCTATCCCTAACA-3′) (200 nM each), β-globin primer pair HBGU (5′-CGGCGGCGGGCGGCGCGGGCTGGGCGGCTTCATCCACGTTCACCTTG-3′) and HBGD (5′-GCCCGGCCCGCCGCGCCCGTCCCGCCGGAGGAGAAGTCTGCCGTT-3′) (300 nM each) were added to the master mix containing 1 × SyberGreen MM, 1 × QN ROX Ref dye to 0.2 ng of DNA in a final volume of 10 μL. Quantification was performed on ViiA7 Real-Time PCR System (Stage 1: 2 min at 95 °C; Stage 2: 2 cycles of 15 s at 94 °C, 15 s at 51 °C; and Stage 3: 32 cycles of 15 s at 94 °C, 10 s at 64 °C, 20 s at 74 °C, 20 s at 86 °C). After, the 74 °C and 86 °C reads were separately analyzed using LinRegPCR [27] and provided the Ct values for the telomere (T) and the single-copy β-globin gene template (S), respectively. Results were expressed as T/S ratio calculated using the reference DNA type XIII from human placenta (Sigma-Aldrich, St. Louis, MO).

Statistical analysis

Pre- and post-transplantation comparisons were performed by two-tailed Wilcoxon’s signed-rank test and values were described as mean ± SD. To compare responder vs. non-responder patients, the two-tailed Mann–Whitney U-test was used and results expressed as median ± interquartile range (IQR). Correlations were assessed by Spearman’s test. The area under the curve (AUC) of each lymphocyte subpopulation or marker was calculated by the trapezoid method [13]. Parameters were not adjusted for patient age. All figures and analyses were performed in GraphPad 7 (La Jolla, CA). Significance was set at P < 0.05.


Clinical response to AHSCT

In the 25 included SSc patients, there was significant decrease in the mRSS from 26.1 ± 8.1 before to 18.9 ± 8 at 6 months (P < 0.001), sustained until 36 months after AHSCT (15.9 ± 7.5, P < 0.001). Percentages of predicted FVC levels remained stable during the entire follow-up (63.2 ± 23.7% at baseline versus 58.2 ± 30.9% at 36 months after AHSCT, P > 0.05), whereas anti-Scl70 autoantibodies titers declined from 180.9 ± 41.9 U/mL before to 150.4 ± 63.2 U/mL at 6 months (P < 0.05), until 18 months post-transplant (127.7 ± 68.2 U/mL, P < 0.05). According to clinical response after AHSCT, 19 patients were classified as responders, as they remained stable or improved at clinical evaluations, and 6 non-responders, as they met at least one of the criteria for relapse after AHSCT and underwent subgroup analyses. Before transplantation, responder and non-responder groups of patients were not different for skin or organ-specific involvement (Table 1).

Homeostatic proliferation leads to telomere attrition and increased frequency of senescent cells early after transplantation

Before transplantation, patients presented negative correlations between age and T/S ratios (P = 0.013, rs = -0.59) and between age and frequency of CD8+CD28CD57+ senescent cells (P = 0.025, rs = -0.54), and positive correlation between T/S ratio and CD8+CD28CD57 + cells (P = 0.040, rs = 0.44, Supplementary Figure S1). There was no correlation between these parameters and clinical status of patients or organ-specific manifestations.

Relative T/S telomere length ratio significantly decreased from 2.97 ± 1.09 before to 2.07 ± 0.54 (P = 0.015, Fig. 1a) at 6 months after AHSCT, decreasing by 41% when compared with baseline (P = 0.015). No differences in telomere length were observed between responder and non-responder patient groups after AHSCT (Fig. 1b).

Fig. 1

Homeostatic proliferation early after AHSCT results in short telomere length and increased counts of CD57- and FoxP3-expressing CD8+CD28- senescent T-cells. a Mean ( ± SD) relative telomere length value as measured by multiplex quantitative RT-PCR analysis on genomic DNA before (0 months) and at semiannual time-points after AHSCT in 25 patients. Results are expressed as T/S ratio. *P < 0.05 comparing pre and post-transplant values (Wilcoxon’s). b Median ( ± IQR) relative telomere length value in responder (n = 19, blue) and non-responder (n = 6, red) patients. Mann–Whitney non-parametric test was used to compare the patient groups. c Correlation between the percentage of CD8+CD28 T-cells and T/S ratio values (Spearman’s) at 6 months post-transplant. d Percentage of CD8+CD28CD57+ (left) and CD8+CD28FoxP3+ (right) T-cells before and at 6 months after AHSCT. Wilcoxon non-parametric test was used to compare pre and post-transplant time points. Median ( ± IQR) e CD8+CD28CD57+ and f CD8+CD28FoxP3+ T-cell values in responder (n = 19, blue) and non-responder (n = 6, red) patients. Mann–Whitney non-parametric test was used to compare patient groups

At 6 months after transplantation, there was significant increase in CD8+CD28 cell percentages, from 13.14 ± 8.75% at pre-AHSCT to 38.14 ± 13.14% at 6 months (Supplementary Figure S2A, P < 0.001), negatively correlating with T/S ratios (rs = – 0.76, P = 0.036, Fig. 1c). At the same time point, expressions of the senescence marker CD57 [5, 7, 12, 13] and of the immunoregulatory transcription factor FoxP3 [28, 29] were increased on CD8+CD28 cells, from 31.24 ± 18.73% at baseline to 44.43 ± 22.27% at 6 months (P = 0.032) and 72.77 ± 26.42 to 83.83 ± 23.39 (P = 0.005), respectively (Supplementary Figure S2B). As result, CD8+CD28CD57+ and CD8+CD28FoxP3+ T-cell percentages (Fig. 1d) and absolute counts (Supplementary Figure S3A) increased, remaining higher (P < 0.05) than baseline values until 1 and 2 years after AHSCT, respectively (Supplementary Figure S4). CD57 expression by CD8+CD28 cells negatively correlated with telomere length (rs = – 0.34, P = 0.015). Frequencies (Fig. 1e, f) and absolute counts (Supplementary Figure S3B-C) of CD8+CD28CD57+ and CD8+CD28FoxP3+ T-cell subsets were not significantly different between responder and non-responder patients after AHSCT. However, CD8+CD28CD57+ cells tended to be more frequent in the responder patients (Fig. 1e, f).

Cytokine and C-reactive protein levels decreased after AHSCT

When compared with pre-transplant levels, concentrations of IL-6, IL-1β, and C-reactive protein were decreased at 6 months after AHSCT and returned to baseline values at 18 months until the end of follow-up, while the concentrations of all other cytokines did not change (Supplementary Table 1). When patient groups were analyzed separately, no differences were observed from 6 to 12 months, but responder patients presented lower levels of TGF-β, IL-6, G-CSF (P < 0.01), and of IL-1β, IL-17A, MIP-1α, and IL-12 (P < 0.05), at 24 months after AHSCT, than non-responders (Fig. 2).

Fig. 2

Reduction of pro-inflammatory cytokines associates with clinical responsiveness of SSc patients to AHSCT. IL-2, IL-6, IL-10, IL-12, IL-13, IL-17, IL-1β, TGF-β1, PDGF, MCP-1, MCP-3, MIP-1α, MIP-1β, G-CSF, TNF-α, and C-reactive protein serum levels were determined before (0 months) and at subsequent time points after AHSCT in 18 responder (blue) and 5 non-responder (red) patients. *P < 0.05, **P < 0.01, responders vs. non-responders (Mann–Whitney)

PD-1 is a marker of responsiveness to AHSCT

At 6 months post-transplant, SSc patients presented higher PD-1 expression on CD3+CD4+ (P = 0.005) as well as on CD3+CD8+ (P = 0.024) T-cells when compared with pre-transplant values, whereas no changes were observed in PD-1 expression on CD19+ B-cells (Supplementary Figure S5A and Fig. 3a). This increase was not sustained, returning to baseline levels at 1 year post-AHSCT (Supplementary Figure S5B). Similar profile was observed for PD-1+ T- and B-cell counts (Supplementary Figure S5C). At 6 months post-AHSCT, the group of responder patients presented higher PD-1 expression on both CD4+ (P < 0.05) and CD8+ (P < 0.05) T-cells, as well as on CD19+ (P < 0.01) B-cells when compared with non-responder patients (Fig. 3b). Throughout follow-up, responder patients presented higher AUC of CD4+PD-1+ T-cells (P = 0.047) and of CD19+PD-1+ B-cells (P = 0.015) when compared with non-responders (Fig. 3c). Responder patients presented higher CD3+CD4+PD-1+ T-cell and CD19+PD-1+ B-cell counts than non-responders at baseline. In addition, responder patients were the only group to experience increased counts of CD3+CD8+PD-1+ T-cells after AHSCT as compared with baseline (Supplementary Figure S6).

Fig. 3

Higher PD-1 expression on T- and B-cells correlates with better clinical outcomes. a Mean ( ± SD) PD-1 expression before and at 6-months post-transplant on CD3+CD4+ T-cells, CD3+CD8+ T-cells and CD19+ B-cells immunophenotyped by flow cytometry (Wilcoxon’s). b Median ( ± IQR) PD-1 expression on T- and B-cells before and at 6 months post-transplant in responder and non-responder patients. *P < 0.05, **P < 0.01 comparing patient groups (Mann–Whitney) and comparing pre and post-transplant time points (Wilcoxon’s). c Median ( ± IQR) PD-1 expression before (0 months) and at subsequent time points in 20 responder (blue) and 6 non-responder (red) patients. *P < 0.05, responders vs. non-responders (Mann–Whitney)

There was no correlation between T/S ratio and PD-1 expression on CD4+ (rs = 0.19, P = 0.66), CD8+ (rs = 0.5, P = 0.21), and CD19+ (rs = 0.35, P = 0.44) cells. In addition, frequencies of PD-1hi T- and B-cells were not different before and after AHSCT, and were also not different between responder and non-responder patients (data not shown).


In autoimmune diseases, thymic rebound associates with positive clinical outcomes after AHSCT [5, 6, 8,9,10,11, 25]. However, the increase in thymic function is a delayed event after transplantation. In the first year that follows AHSCT, immunological tolerance is usually based on thymus-independent mechanisms, such as depletion of autoreactive cells [30,31,32] and lymphopenia-induced proliferation of regulatory T-cells, which have also been associated with better clinical outcomes [6, 12, 13, 25]. The rise of homeostatic-proliferating cells limits the space available for autoreactive cells that were not depleted or that were re-infused within the graft, thereby controlling the “immunological space” through expansion of senescent/exhausted cells, whereas newly generated cells with broader diversity and higher immunoregulatory functions are not yet being produced by the thymus [33, 34]. Considering the lack of studies evaluating early immune mechanisms after AHSCT for SSc patients, we raised the question whether telomere attrition and increase of senescent/exhausted cells could influence long-term clinical response to AHSCT.

Telomere dysfunction has been described in several bone marrow disorders, including aplastic anemia and hematopoietic malignancies [17]. SSc patients present longer telomere lengths [35] and reduced telomerase activity [36] when compared with healthy counterparts and to other autoimmune conditions, indicating possible role of a specific aberrant telomere biology on disease pathogenesis. HSCT settings provide an opportunity to evaluate telomere dynamics in a highly proliferative environment and to further evaluate how it associates with the process of immune reconstitution and with clinical outcomes. Several studies have shown that telomere attrition is limited to the first year after HSCT for hematological indications due to high rates of homeostatic proliferation [18,19,20]. Indeed, our SSc patients presented significant reduction of telomere length at 6 months post-AHSCT, as consequence of high rates of homeostatic proliferation, indicating that this may be a universal mechanism after HSCTs. The telomere dynamics is different between leucocytes and lymphocyte subpopulations. Although granulocytes, monocytes, B-cells, and naive CD4T-cells present lower telomere lengths until 1 year post-transplantation, memory CD4T-cells tend to present higher telomere length [37]. This indicates that transplant-induced replicative stress may be restricted to specific lymphocyte subpopulations. This is a subject for future analyses.

The observed negative correlation between telomere length and frequency of CD8+CD28 senescent cells indicates that telomere shortening early after AHSCT is probably related to cellular senescence in the lymphopenic environment [5, 12, 13]. In addition, these data corroborate previous findings of shorter telomere lengths in senescent CD8+CD28 cells [38, 39]. Here we show that both responder and non-responder groups of patients presented similar rates of telomere reduction during the entire follow-up, indicating that telomere attrition does not associate with clinical response of SSc patients after AHSCT. Important, results may have been biased by the heterogeneity of certain parameters and by the dependence of telomere length and frequency of senescent cells on patient age. Prospective studies considering these factors are needed to further define the role of these parameters on clinical response after AHSCT in autoimmune diseases.

CD8+CD28 T-cells bear immunoregulatory properties [40], such as down-modulation of cytotoxic activity [41] and production of inhibitory soluble factors [42], and their numbers are usually found decreased in autoimmune diseases [43]. In type 1 diabetes (T1D) patients, increased CD8+CD28 cell frequencies after AHSCT may have contributed to hinder the autoimmune destruction of insulin-producing cells and to improve glycemic control [25]. Here we observed that responder SSc patients presented a trend toward an increase of these cell counts after AHSCT, indicating possible roles of homeostatic proliferation and immunological attrition. Furthermore, increased FoxP3 and CD57 expressions on CD8+CD28 T-cells early post-transplantation indicates the presence of senescent T-cells with immunoregulatory properties, which may contribute to control SSc activity.

Replicative senescence leads to clonal exhaustion and to cells expressing high levels of PD-1 [44, 45]. In transplantation, PD-1 is required for self-tolerance during the establishment of immune homeostasis [46]. We have previously reported in MS patients that high PD-1 expression on CD8+ T-cells and on CD19+ B-cells after AHSCT correlates with better clinical outcomes [13]. In the present study, higher PD-1 expression by CD4+ and CD8+ T-cells was observed only in the responder patients after AHSCT, indicating that in SSc PD-1 expression may be a marker of positive clinical response after AHSCT.

In the context of AHSCT, high PD-1 levels are expressed by cells after several rounds of proliferation [12, 33], as a possible mechanism to avoid development of secondary autoimmune diseases in lymphopenic environments [47]. PD-1 expression does not correlate with telomere length and increased PD-1 expression may be related to normalized expression of the PDCD1 gene [12]. In addition, PD-1-expressing cells are more susceptible to apoptosis. In fact, elevated frequencies of Fas+CD4+/CD8+ T-cells, as well as increased expression of apoptosis-related molecules, were demonstrated in early phases of immune reconstitution in MS and T1D patients after AHSCT, also indicating increased susceptibility to apoptosis [5, 48, 49]. Deeper phenotypic evaluation of PD-1+ T-cells after AHSCT is still required to assess if these cells are CD45RO+ or terminally differentiated effector memory cells re-expressing CD45RA (TEMRA) [50].

AHSCT is followed by profound lymphopenia. Although thymic rebound is not yet installed, homeostatic mechanisms are essential to restore the immunological response against pathogens. Our results demonstrate that telomere attrition, as result of replicative senescence, is not related to clinical responsiveness of SSc patients to AHSCT. However, we show increased expression of PD-1 in responder patients, which may be a general mechanism to keep potentially autoreactive CD8+ T-cell clones under control after AHSCT. In accordance with a report on MS patients [13], PD-1 expression appears to be a reliable immune marker of clinical response in SSc patients after AHSCT.


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This work was supported by the financial research agencies CNPq, INSERM, and FAPESP (Scholarship numbers: 2013/18678-3, 2014/20922-2; Center for Cell-Based Therapy, CEPID-FAPESP, grant number 2013/08135-2).

Author contributions

MCO and KCRM are the principal investigators and take primary responsibility for the paper. LCMA, KCRM, AT, and MCO designed the study. LCMA, JRL-J, CD, IF, and EC performed the experiments. LCMA, JRL-J, DAM, BPS, and HM-T collected the data and performed data analysis. DTC provided essential funding to the development of this work. LCMA, KCRM, AT, EC, and MCO wrote the final report. All authors contributed to the editing of the final report. All authors agree on all of the content of the submitted manuscript.

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Correspondence to Maria Carolina Oliveira.

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The authors declare that they have no conflict of interest.

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These authors share senior authorship: Kelen C. R. Malmegrim, Maria Carolina Oliveira.

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Arruda, L.C.M., Lima-Júnior, J.R., Clave, E. et al. Homeostatic proliferation leads to telomere attrition and increased PD-1 expression after autologous hematopoietic SCT for systemic sclerosis. Bone Marrow Transplant 53, 1319–1327 (2018) doi:10.1038/s41409-018-0162-0

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