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Immunity. Author manuscript; available in PMC 2017 October 18. Published in final edited form as: Immunity. 2016 October 18; 45(4): 903–916. doi:10.1016/j.immuni.2016.09.013.
Deficient activity of the nuclease MRE11A induces T cell aging and promotes arthritogenic effector functions in patients with rheumatoid arthritis Yinyin Li1, Yi Shen1, Philipp Hohensinner1,2, Jihang Ju1, Zhenke Wen1, Stuart B. Goodman3, Hui Zhang1, Jörg J. Goronzy1, and Cornelia M. Weyand1,4
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1Division
of Immunology and Rheumatology, Stanford University School of Medicine, Stanford, CA
94305 2Department
of Internal Medicine II/Cardiology, Medical University of Vienna, Vienna, Austria
3Department
of Orthopedic Surgery and Bioengineering, Stanford University School of Medicine, Stanford, CA 94305
Abstract
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Immune aging manifests with a combination of failing adaptive immunity and insufficiently restrained inflammation. In patients with rheumatoid arthritis (RA), T cell aging occurs prematurely, but the mechanisms involved and their contribution to tissue-destructive inflammation remain unclear. We found that RA CD4+ T cells showed signs of aging during their primary immune responses and differentiated into tissue-invasive, pro-inflammatory effector cells. RA T cells had low expression of the double-strand-break repair nuclease MRE11A, leading to telomeric damage, juxtacentromeric heterochromatin unraveling, and senescence marker upregulation. Inhibition of MRE11A activity in healthy T cells induced the aging phenotype, whereas MRE11A overexpression in RA T cells reversed it. In human-synovium chimeric mice, MRE11Alow T cells were tissue-invasive and pro-arthritogenic, and MRE11A reconstitution mitigated synovitis. Our findings link premature T cell aging and tissue-invasiveness to telomere deprotection and heterochromatin unpacking, identifying MRE11A as a therapeutic target to combat immune aging and suppress dysregulated tissue inflammation.
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Correspondence To: Cornelia M. Weyand, M.D., Ph.D., Division of Immunology and Rheumatology, Department of Medicine, Stanford University, CCSR Building Room 2225, MC-5166, 269 Campus Drive West, Stanford, CA 94305; Phone: (650) 723-9027, Fax: (650) 721-1251, [email protected]. 4Lead Contact Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. COMPETING FINANCIAL INTERESTS The authors declare that no conflict of interest exists. Additional experimental procedures are presented in Supplemental Materials. AUTHOR CONTRIBUTIONS YL, JJG and CMW designed the research and analyzed data. YL, YS, PH, JJ, ZW and HZ were responsible for the experimental work. SG recruited patients. CMW, YL and JJG wrote the manuscript.
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INTRODUCTION Preceded by a decade-long period of preclinical disease, rheumatoid arthritis (RA) manifests with a symmetrical polyarthritis causing irreversible cartilage and bone destruction and shortens life expectancy due to accelerated cardiovascular disease. Immune aging affects the general population after 50 years of age, but is accelerated in RA patients (Weyand et al., 2009), where it is already noticeable in antigen-unprimed naïve T cells (Koetz et al., 2000).
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Cells devote a significant proportion of their machinery to DNA surveillance and repair to prevent cellular aging or death associated with genome instability (Chow and Herrup, 2015). Predictable loss of telomeric sequences with each cell replication allows telomeres to serve as molecular clocks. By tallying the number of cell divisions, telomeres are believed to effectively force mutation-harboring cells into cell cycle arrest. Senescent T cells not only remain viable, but actively shape the tissue microenvironment by secreting cytokines and tissue remodeling factors (Weyand et al., 2014). However, despite several senescence features, aging human lymphocytes are not in replicative arrest (Yang et al., 2016) and continue to participate in clonal expansion, distinguishing lymphocyte aging from senescence (Akbar and Henson, 2011; Chou and Effros, 2013; Sharpless and Sherr, 2015). Reversibility of senescence in human end-differentiated effector T cells further supports the model that aging of lymphocytes reflects progressive differentiation more than true senescence (Di Mitri et al., 2011). Whether aging T cells acquire effector functions that mediate tissue inflammation is not understood.
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Abnormalities in the DNA damage sensing and repair machinery of RA T cells have raised the question of whether such defects are mechanistically linked to T cell aging and to arthritogenic effector functions (Shao et al., 2009; Shao et al., 2010). The MRN complex, composed of Meiotic Recombination 11 Homolog A (MRE11A), RAD50 and Nijmegen Breakage Syndrome 1 (NBS1), senses DNA double-strand breaks to amplify DNA repair (Lamarche et al., 2010). The core component of the complex, MRE11A, has doublestranded (ds)DNA exonuclease and single-stranded (ss)DNA endonuclease activity in both homologous recombination and nonhomologous end-joining (Xie et al., 2009). MRE11A is recruited to healthy telomeres, where its function is not understood. In S. cerevisiae, RAD50 or MRE11A loss shortens telomeres and abolishes S phase telomerase recruitment (Takata et al., 2005). In high eukaryotes, MRE11A specifically interacts with the shelterin protein TRF2 (Diotti and Loayza, 2011). Conversely, MRN complex hypomorphism in mouse embryonic fibroblasts does not affect telomere length (Attwooll et al., 2009) and reduces telomere dysfunction-induced foci (TIF), revealing the complexity of MRE11A function. Which role MRE11A plays in healthy and stressed human cells, particularly in long-lived lymphocytes, is unknown. Here, we report that in RA T cells, age-related telomeric defects manifested not only as shortening, but also as damage accumulation. Such T cells unfolded higher-order satellite heterochromatin, expressed increased levels of the cell cycle regulators p16 and p21 but lacked p53. Aged RA T cells were low-expressers for the DNA repair nuclease MRE11A and their chromosomal ends were MRE11Alow. In healthy T cells, decreasing MRE11A mRNA by treatment with short interfering RNAs (siRNAs) or pharmacologic inhibition of
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MRE11A’s nucleolytic activity promptly induced telomeric damage and upregulated the senescence markers p16, p21, and CD57, concomitant with unraveling of pericentromeric satellite DNA. Spontaneous or induced deficiency of MRE11A’s nucleolytic function had a profound impact on T cell behavior and rendered T cells tissue-invasive and pro-arthrogenic, whereas reconstitution of MRE11A protein in patient-derived T cells protected synovial tissue from inflammatory attack. These data provide mechanistic evidence for a role of the MRE11A nuclease in not only regulating aging but also differentiation of T cells into tissueinjurious effector cells.
RESULTS Telomeres in RA T cells are not only shortened, but damaged
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Telomeric sequences in RA T cells are shortened relative to T cells from age-matched healthy individuals, and this has been attributed to increased proliferative pressure in an inflammatory environment (Koetz et al., 2000). However, T cell turnover measured by the proliferation marker Ki-67 correlates inversely with telomeric erosion (Schonland et al., 2003), suggesting alternative mechanisms underlying telomeric loss. Telo-FISH staining in metaphase nuclei confirmed that the vast majority of RA naïve CD4+ T cells had lowintensity telomeric signals (Figure 1A and 1B). Compared to healthy individuals, RA patients lacked high-brightness nuclei and almost all of their cells had a weak telomere signal (Figure 1C).
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The structural intactness of telomeric caps was examined through FISH-probe hybridization patterns in metaphase-arrested nuclei, revealing four structural patterns: telomeric apposition (long chromosomal arms opposed), telomeric fragmentation (signal doubling), telomeric loss (signal-free end) and telomeric fusion (physical linkage of two ends of the same or neighboring chromosomes) (Figure 1D). The majority of healthy metaphase T cell nuclei had structurally intact telomeres (56.5%). Thirty-one percent of nuclei displayed signal doublets and 12.5% displayed telomeric apposition. In contrast, RA T cell nuclei consistently contained damaged telomeres. End signal doubling, indicative of telomeric fragility, occurred in 41.9% of cells and 24.1% had opposed telomeres. Albeit encountered infrequently, RA samples showed evidence for severe structural damage, including fusions (2.8%) and total signal loss (1.6%) (Figure 1E).
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To understand the relationship between proliferation-induced telomeric shortening and TIF formation, we monitored TIF evolution in polyclonally expanding T cell populations by quantifying the recruitment of the DNA damage protein P53-Binding Protein 1 (53PB1) to telomeric ends marked through the shelterin protein TRF2. Naïve and memory CD4+ T cells from the same donor were placed under proliferative stress (Figure S1A) by repetitive polyclonal stimulation. Enforced proliferation markedly shortened telomeres in the naïve population, but lengths were relatively stable in memory counterparts, despite similar proliferation rates in both T cell subpopulations (Figure S1A, S1B, S1D). Naïve CD4+ T cells lost >1000 telomeric nucleotides over 3 weeks, but memory CD4+ T cell chromosomes were shortened by 300 base pairs (bp). Parallel monitoring of TIF by quantifying 53BP1 and TRF2 colocalization demonstrated rare foci in naïve CD4+ T cells, even after >1000 bp loss. In contrast, TIF appeared as early as day 7 in memory CD4+ T cells (Figure S1C). Overall, Immunity. Author manuscript; available in PMC 2017 October 18.
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53BP1 and TRF2 colocalization and erosion of telomeric sequences were unrelated (Figure 1F). Together, these data documented that telomeres in RA T cells are shortened, but more importantly, are sites of DNA damage. Premature aging of naïve CD4+ T cells in RA patients is related to telomeric damage
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The presence of circulating and synovial CD4+CD28null T cells in RA patients provides unequivocal evidence for in vivo T cell aging. Such end-differentiated T cells express CD57 and natural killer (NK) cell receptors (Warrington et al., 2001). To better define the aging process in naïve CD4+CD45RA+ T cells, we assessed the aging-associated cell cycle inhibitors p16, p21, and p53 (Sharpless and Sherr, 2015). Compared to healthy controls, RA T cells expressed higher levels of CDKN2A and CDKN1A mRNA and p16 protein (Figure 2A–B), while TP53 expression was reduced, suggesting a p16-dependent and p53independent aging pathway (Figure 2A). The glucuronyltransferase gene family member CD57 is considered a senescence marker on T cells (Focosi et al., 2010). The frequency of CD4+CD45RA+CD57+ T cells was low in healthy individuals (7%), but more than doubled in RA patients (20%) (Figure 2C).
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DNA condensation into senescent-associated heterochromatin foci is a hallmark of cellular senescence, but early stages of cellular aging can be associated with the unravelling of constitutive heterochromatin into visibly extended structures (Swanson et. al., 2013). To identify heterochromatin changes, we analyzed CD4+CD45RA+ T cell populations for evidence of unraveling of pericentromeric and centromeric satellite DNA. Hybridization of a probe to satellite-II (sat-II) DNA in stimulated healthy CD4+CD45RA+ T cells yielded compact, round and centric satellite signals (Figure 2D). Conversely, RA T cells had distended pericentromeric and centromeric satellites, usually affecting multiple sites within a nucleus. Eighty percent of RA cells had clearly unraveled satellites. In contrast, 80% of control cells had tightly packaged condensed satellites (Figure 2E), demonstrating that pericentromeric and centromeric satellite heterochromatin in RA T cells undergoes decondensation, consistent with entry into early senescence. To investigate whether aging-associated T cell phenotypes are mechanistically linked to telomeric damage, we used siRNAs to decrease telomere-protecting shelterin TERF2 mRNA. Transfecting TERF2 siRNAs or a TERF2 mutant plasmid reduced TERF2 mRNA (Figure S3A); induced robust TIF formation (Figure S3B) and doubled CDKN2A and CDKN1A mRNA transcripts (Figure 2F, Figure S3C). TERF2 knockdown was sufficient to induce p16 and CD57 expression (Figure 2G–H), recapitulating conditions in RA patients. T cells from RA patients are MRE11A deficient
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The presence of telomeric damage foci and satellite DNA distention in RA T cells raised the question whether the DNA repair machinery was fully intact. We examined RA T cells for the expression of key molecules involved in the DNA double strand break repair, including Ataxia Telangiectasia and Rad3-Related Protein (ATR), Ataxia Telangiectasia Mutated (ATM), MRE11A, NBS1, RAD50 and DNA-dependent protein kinase catalytic subunit (DNA-PKcs) by flow cytometry. Intracellular staining of peripheral blood mononuclear cells (PBMCs) demonstrated reduced MRE11A protein expression in naïve and memory CD4+ T
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cells from RA patients (Figure 3A–B). In both T cell subpopulations, MRE11A protein expression declined in an age-dependent fashion in RA patients as well as healthy donors. However, MRE11A protein was consistently lower in RA patients’ cells; reduced by 40– 50% in the naïve population and 35% in the memory population. CD4+ T cells of patients with the inflammatory polyarthritis, PsA, were indistinguishable from healthy T cells (Figure S4A). In RA patients, but not in PsA patients, CD8+ T cells, CD14+ monocytes and CD19+ B cells were also MRE11Alow expressors (Figure S4B, S4C, S4D). MRE11A lowexpression was most pronounced in T cells from untreated RA patients, eliminating immunosuppressive therapy as the causative factor (Figure 3C–D). Total nuclear and telomere-bound MRE11A was analyzed by dual-color immunostaining with anti-MRE11A and anti-TRF2. Staining intensity for MRE11A protein was markedly reduced in RA T cell nuclei, in which the signal for MRE11A and TRF2 colocalization was especially low, suggesting that MRE11Alow RA T cells have telomeric tips almost depleted of the nuclease (Figure 3E–F). Thus, aging is associated with progressive decline in the expression of the double strand break repair nuclease MRE11A, a process which is notably accelerated in RA T cells. Diminished MRE11A activity causes telomeric damage
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The exonuclease and endonuclease MRE11A partners with the ATPase RAD50 and NBS1 to form the MRN complex, a DNA break sensor critically involved in multiple repair pathways (Lamarche et al., 2010). siRNA-mediated knockdown of each of the three MRN components, MRE11A, RAD50 and NBS1, produced partial depletion of both transcripts and protein (Figure S5A–S5B) and induced widespread DNA damage (Figure 4A). The amount of unrepaired DNA breaks was quantified by measuring the intensity of 53BP1binding to damaged chromatin sites, where this checkpoint protein forms foci and regulates end-joining-mediated repair processes. MRE11A loss was most devastating for DNA integrity; 53BP1 was recruited to numerous DNA break sites in almost every single nucleus. In contrast, NBS1 knockdown left some nuclei unaffected. Partial loss of RAD50 was tolerated the best, producing the lowest 53BP1 signal. 53BP1 and TRF2 colocalization studies confirmed the functional relevance of MRE11A at the telomere (Figure S5C). Overall, transfection of MRE11A-specific siRNA produced robust DNA damage, with almost 50% of the 53BP1 signal localizing to telomeric ends. Only one-third of the 53BP1 signal copositioned with the shelterin protein TRF2 in NBS1 or RAD50 insufficient cells (Figure 4B). Alternatively, the function of MRE11A was impaired by treating with the small molecule inhibitor Mirin, which inhibits MRE11A’s exonuclease activity while sparing the endonuclease activity (Dupre et al., 2008; Shibata et al., 2014). Inhibition of exonuclease activity in healthy naïve CD4+ T cells resulted in massive DNA damage, of which a substantial proportion localized to the telomere (Figure 4C and Figure S5D). These data confirmed that the nucleolytic activity of MRE11A is particularly important in protecting telomeric integrity. Impairing MRE11A activity induces the aging profile of RA T cells To investigate whether MRE11Alow T cells are prone to accelerate the aging process, we targeted MRE11A, RAD50 and NBS1 with interfering RNA technology. MRE11A knockdown potently induced CDKN2A, NBS1 knockdown resulted in a significant, but Immunity. Author manuscript; available in PMC 2017 October 18.
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small, increase in CDKN2A mRNA and RAD50 knockdown left CDKN2A expression unchanged. TP53 expression was unaffected by reducing MRE11A expression (Figure 4D). Treatment with the MRE11A inhibitor Mirin led to similar results (Figure 4E), indicating that MRE11A’s nucleolytic activity protected T cells from entering the aging program. Overall, the pattern of p16 induction paralleled the degree of telomeric damage induced by the genetic inhibition of the three molecules (Figure 2), with MRE11A trumping the two other components in TIF induction and p16 upregulation. Weaning of the inhibitory effects of the siRNAs and recovery of MRE11A expression were sufficient to reverse the upregulation of CDKN2A (encoding p16) and CDKN1A (encoding p21) (Figure S5E–S5F). CD57 expression in MRE11A siRNA-treated cells (Figure 4F) recapitulated the spontaneous aging profile in patient-derived T cells (Figure 2). Similarly, Mirin treatment more than doubled the frequency of CD57+CD4+ T cells (Figure 4G) and rapidly induced heterochromatin unpacking (Figure 4H). More than 80% of Mirin-treated cells had distended satellites, reproducing the constitutive satellite DNA unraveling in RA T cells (Figure 4I). These experiments mechanistically linked the nuclease MRE11A to the aging phenotype of T cells, including the upregulation of cell cycle regulators and the unraveling of satellite chromatin. Restoring MRE11A expression repairs telomeric damage and prevents T cell aging
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To examine whether MRE11A restoration is sufficient to revert the aging phenotype, an MRE11A construct was overexpressed in RA CD4+ naïve T cells (Figure 5A and 5B). Increasing the MRE11A protein concentration prevented telomeric 53BP1 recruitment; 53BP1 and TRF2 colocalization declined by 75% (Figure 5C and 5D), suggesting a direct role of MRE11A in telomere repair. Other components of the T cell aging program, such as p16 expression, were similarly affected (Figure 5E–5G). Sixty percent of control-transfected RA T cells were p16pos and frequencies were reduced to 20% after MRE11A overexpression (Figure 5H). Consistent with the data in Figure 4D, p53 expression appeared independent of MRE11A protein concentrations. These data confirmed that MRE11A has anti-aging function by securing telomeric intactness and controlling cell cycle regulators and that the aging signature of RA T cells is reversible. Inhibiting MRE11A’s nucleolytic activity renders T cells tissue-invasive and proinflammatory
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CD4+CD28null T cells participate in the inflammatory lesions in RA joints (Schmidt et al., 1996), but whether their accelerated aging is mechanistically connected to their proinflammatory functions is unknown. To establish tissue inflammation, T cells must migrate into the tissue site where they display proinflammatory effector functions, such as cytokine and chemokine release, activation of endothelial cells, stromal cells and innate immune cells. All phases of this process are captured in a humanized mouse model in which human synovial tissue is engrafted into immunodeficient mice and human CD45RO− PBMCs, either deriving from healthy individuals or from patients with RA, are adoptively transferred. Chimeric mice were reconstituted with either control or RA PBMCs and were treated with the MRE11A inhibitor Mirin or vehicle (Figure 6). Adoptively transferred PBMCs from
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healthy individuals formed sparse T cell infiltrates in the synovial grafts (Figure 6A). Inhibition of MRE11A’s nucleolytic activity enhanced T cell accumulation in the synovial tissue. MRE11Alow RA PBMCs spontaneously had a much higher degree of tissue invasiveness. Here, MRE11A inhibition marginally enhanced T cell recruitment into the synovium. Synovial T cell accumulation was similar in Mirin-treated control PBMCs and untreated RA PBMCs (Figure 6B–C). Inhibiting MRE11A activity resulted in upregulated expression of TNFSF11 (encoding RANKL), a molecule critically involved in rheumatoid bone destruction (Figure 6D). Also, the intensity of tissue inflammation was increased, as measured by the expression of the pro-inflammatory cytokines TNF, IL6 and IL1B (Figure 6E). Conversely, reducing MRE11A activity prompted increased tissue expression of the regulatory cytokines TGFB1 and IL10 (Figure 6F). Exposure of synovial tissue to the MRE11A inhibitor in the absence of transferred PBMCs had no impact on cytokine expression (Figure 6E–F).
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To examine a possible link between tissue invasiveness, inflammatory capability and cellular aging, we quantified tissue expression of CDKN2A, CDKN1A and TP53 transcripts and measured p16 protein expression by dual-color immunohistochemistry (Figure 6G–I). Mirin treatment left the cell cycle regulator expression in noninflamed synovial cells unaffected, but promptly induced the CDKN2Ahigh, CDKN1Ahigh, TP53low profile in tissue-infiltrating cells (Figure 6G). Loss of MRE11A function rendered T cells and non-T cells p16-positive (Figure 6H–I). RA T cells formed dense synovial infiltrates, with a visible enrichment of CD3+ p16+ cells. Also, MRE11Alow RA T cells had a hypermigratory phenotype in Transwell assays performed in the absence of chemokines (Figure 6J). Hypermotility was shared by T cells from old individuals and T cells treated with the MRE11A inhibitor Mirin (Figure 6J). Overall, healthy T cells with inhibited MRE11A nuclease activity resembled RA T cells. These experiments assigned a mechanistic role to the nuclease MRE11A in several disease-relevant capabilities of RA T cells; specifically, their tissue-invasive and aggressive inflammatory behavior. Restoring MRE11A in RA cells prevents pro-arthritogenic effector functions
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To test whether restoring MRE11A expression was sufficient to correct the pathogenic behavior of RA CD4+ T cells, we overexpressed MRE11A in naïve and memory CD4+ T cells prior to adoptive transfer. Forced MRE11A overexpression was strongly antiinflammatory (Figure 7). Transfer of MRE11Ahigh RA T cells minimized tissue TRB transcript levels, diminished the density of the T cell infiltrate, reduced TNF, IL6, IL1B expression, increased TGFB1 and IL10 mRNA and prevented CDKN2A induction. Correcting MRE11A protein concentrations in the transferred T cells did not affect T cell lineage commitment, as indicated by the expression of the T cell transcription factors TBX21, GATA3, RORC and FOXP3 (Figure S6). These experiments confirmed that MRE11A is a critical regulator of pathogenic effector functions in tissue-residing RA T cells and that restoring MRE11A protein expression is sufficient to modify arthritogenic capabilities in naïve and memory CD4+ T cells.
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DISCUSSION
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RA is an HLA class II-associated autoimmune disease, in which arthritogenic T cells drive progressive inflammatory destruction of cartilage and bone. Patients with RA have a signature of premature immune aging, providing an excellent model system to study the interrelationship between failing T cell immunity and unremitting tissue inflammation. Here, we present data implicating defective DNA damage responses in accelerating T cell aging, which renders such T cells susceptible to differentiate into tissue-invasive, pro-athrogenic effector cells. Specifically, low expression of the double strand break repair nuclease MRE11A in RA T cells diminishes dsDNA exonucleolytic and ssDNA endonucleolytic activity, which appears critically important in maintaining intact telomeres, preventing entry into the aging program and averting tissue invasion of inflammation-biased T cells. These findings support the concept that telomeric erosion in MRE11Alow T cells is a consequence of insufficient repair and not proliferative stress and that telomeric damage is mechanistically linked to cellular behavior.
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In this study, we defined the T cell aging program in RA, which is distinct from cellular senescence (Chou and Effros, 2013). How T cells age and whether immune cells follow a “universal” aging program is insufficiently understood, but RA T cells and T cells from aged humans lack the irreversible cell cycle arrest that is considered a cardinal feature of senescence (Goronzy and Weyand, 2013; Sharpless and Sherr, 2015). In contrast, cell cycle passage of RA T cells is shortened and the entire population of patient-derived CD4 T cells is hyperproliferative (Fujii et al., 2009). Synovial T cells have all converted into effector memory T cells, but naïve CD4+ T cells in RA already display abnormalities, e.g., due to increased apoptotic susceptibility clonal expansion is relatively futile (Schonland et al., 2003). Long before differentiating into effector memory cells, RA T cells fail to repair DNA damage (Shao et al., 2009), are energy deprived (Yang et al., 2013) and chronically activate the DNA-PKcs-JNK pathway (Shao et al., 2010). The downregulation of MRE11A was a global phenomenon in RA T cells affecting the entire population and was not selective for a subpopulation. Flow cytometric quantification of MRE11A protein produced unimodal distribution patterns in both CD45RA+ and CD45RA− populations, indicating a broadly distributed defect amongst T cells. MRE11A overexpression rescued the pathogenic effector functions of both, naïve CD45RA+ and memory CD45RO+ CD4+ T cells, when adoptively transferred into chimeric mice. Experiments with isolated CD45RA+CD4+ T cells allowed focusing on early phases in the life cycle of T cells, as they begin to make the transition into effector and memory T cells, that eventually mediate the pathogenic functions in the tissue environment. The human CD8+, but not the CD4+ T cell compartment contains small populations of T cells that increase in frequency with age and that can reacquire a naïve-like phenotype after in vivo stimulation while maintaining the ability to rapidly respond to restimulation (Fuertes Marraco et al., 2015; Pulko et al., 2016). In healthy humans, such CD8 naïve-like T cells with effector function have relatively long telomeres, longer than memory T cells, indicating that they do not have a defect in telomeric maintenance. Cellular senescence, genomic instability, epigenetic alterations, stem cell exhaustion, loss of proteostasis, mitochondrial dysfunction and deregulated nutrient sensing are all critical components in organismal aging (Lepez-Otin et al., 2013). Current data defined the T cell
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aging signature as p16high, p21high and p53low. p16INK4A accumulation is characteristic of aging in other lymphoid and nonlymphoid cells (Chen et al., 2011; Cosgrove et al., 2014; Janzen et al., 2006; Liu et al., 2011; Molofsky et al., 2006; Signer et al., 2008; Sousa-Victor et al., 2014). More difficult to understand is the p21-gain and p53-loss combination, as DNA-damage-induced ATM-p53-p21 signaling typically drives growth arrest of senescent T cells (Bartkova et al., 2006; Fumagalli et al., 2014; Nakamura et al., 2008; Rodier et al., 2011). However, in cancer cells p21 expression can occur in the absence of p53 function (Jeong et al., 2010; Macleod et al., 1995). In RA T cells, p53 is consistently low (Maas et al., 2005; Shao et al., 2009), associating aging with a p53-independent, p16-dependent mechanism. Naïve RA CD4+ T cells include a considerable population of CD57+ cells. How CD57 alters the behavior of old versus young CD4+ T cells is unknown, but it could possibly affect tissue homing.
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Epigenetic changes to chromatin are essential to cellular aging. Senescence-associated distension of satellites (SADS) formation is a unifying process in most models of early senescence, believed to signify the loss of higher-order chromatin packaging (Swanson et al., 2013). The packaging of pericentromeric and centromeric satellite DNA, especially satII DNA, in RA T cells was visibly altered. Whether distensions in sat-II DNA are associated with transcription at these loci will need examination. So far, pericentromeres are considered transcriptionally inert chromatin conformations, but recent evidence emphasizes active and regulated transcription, especially in the context of development, tumorigenesis, senescence and cellular stress (Dejardin, 2015; Saksouk et al., 2015). MRE11A inhibition induced pericentromere unraveling, linking satellite packaging with DNA repair and emphasizing the nuclease’s role in protecting the epigenetic landscape. The MRN complex is a multifunctional enzyme conglomerate known to activate cell cycle checkpoints when sensing DNA breakage (Stracker and Petrini, 2011). In addition to a critical role in ATM function and double strand break repair, MRN is also implicated in suppression of lymphomagenesis (Balestrini et al., 2015) and regulating metastatic breast cancers (Gupta et al., 2013). In humans, MRE11A mutations cause an ataxia-telangiectasia-like disease, with considerable variability in clinical phenotypes (Taylor et al., 2015). In primary human T cells, MRE11A appears to have primarily a protective function in telomere maintenance. Reducing MRE11A abundance by 50% rapidly induced the formation of telomeric damage foci, long before such T cells shortened telomeric length. MRE11A deficiency was more effective in inducing telomeric damage foci than the knockdown of RAD50 or NBS1. Vice versa, replenishing MRE11A protein by forced overexpression promptly reversed telomeric damage. These data suggest MRE11A’s participation in telomere repair and protection in human T cells.
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MRE11A is an abundant protein in human T cells. A 50% reduction, however, had profound consequences, revealed by the reversal of several phenotypes after MRE11A overexpression. Although overexpression prevented TIF formation and suppressed p16, p21 and CD57 expression, the mechanistic connection between telomeric damage and T cell aging awaits further clarification. Pharmacological inhibition of MRE11A’s nucleolytic activity recapitulated all abnormalities of aged T cells, placing insufficient exonuclease activity at the pinnacle of the T cell aging program. Systemic inflammation was insufficient to explain
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MRE11A downregulation. In PsA, an aggressive multi-organ inflammatory syndrome, MRE11A expression was normal. Untreated RA patients had the lowest MRE11A concentrations; immunosuppressive therapy improved but did not normalize the nuclease’s expression, implicating mechanisms other than systemic inflammation in MRE11A suppression.
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In previous reports, the failure to degrade cytoplasmic DNA induced inflammatory responses in bone marrow macrophages to eventually induce polyarthritis (Kawane et al., 2006). Current data link DNA damage in T cells, not macrophages, to functional abnormalities driving synovitis. Mechanistically, MRE11A’s nuclease activity appears to be related to in vivo T cell trafficking and T cell control of the synovial microenvironment. Human synovium and human T cells in chimeric mice enabled us to investigate how insufficient MRE11A activity regulates nonlymphoid tissue infiltration and inflammation. RA T cells have been reported to be spontaneously hypermigratory (Hwang et al., 2015) and we found that nuclease inhibition rendered them hyper-motile and tissue-invasive. Once in the tissue, MRE11Alow T cells supported p16 and p21 expression, left p53 unaffected and greatly increased the abundance of classical pro-inflammatory cytokines. MRE11A restoration did not change T cell lineage commitment but appeared to regenerate an antiinflammatory tissue environment, suggestive for a role of T cells in regulating joint-residing cells, such as synovial fibroblasts.
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The current study has implication for the understanding of the immune aging process in humans and could change the approach towards treating senescence-associated tissue inflammation. Experimental data focused attention on DNA damage sensing and repair as a key event in immune aging, particularly in RA. Impaired DNA damage responses affected T cells in their naïve state and fundamentally change their differentiation program, biasing them towards tissue-invasive and pro-inflammatory behavior. Inflammation per se, as in PsA patients, was insufficient to repress MRE11A expression and the associated T cell aging profile. MRE11A insufficiency appeared involved in multiple aspects of T cell aging, from unraveling of heterochromatin, to telomeric damage, changed expression of cell cycle regulators and altered tissue homing. Restoring MRE11A expression was sufficient to reverse aging features of RA T cells, identifying DNA damage repair, and particularly MRE11A, as a promising therapeutic target to treat immune-aging related disease.
EXPERIMENTAL PROCEDURES Patients and control individuals
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RA patients recruited into the study fulfilled the 2010 diagnostic criteria for RA and were positive for anti-CCP antibodies and for rheumatoid factor (Table S1). Healthy controls were matched for age and gender. The following were exclusion criteria for their enrollment: personal or family history of autoimmune disease, malignancy, chronic infection or another inflammatory syndrome. The Institutional Review Board approved the study and written informed consent was obtained from all participants.
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T cell purification and culture
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Peripheral blood mononuclear cells (PBMCs) were separated from whole blood using Lymphocyte Separation Medium (Corning). Naïve CD4+ T cells were isolated from PBMCs using a human naïve CD4+ T cell isolation kit according to the manufacturer’s instructions (STEMCELL Technologies). This purification procedure minimizes the contamination of CD4+CD45RA+ cells with end-differentiated CD4+CD28− T cells, which accounted for 0.4% of cells in both patients and controls (Figure S2). Naïve CD4+ T cells were cultivated in RPMI 1640 medium (Fisher Scientific) supplemented with 10% FBS (Atlanta Biologicals) and Pen/Strep (Fisher Scientific). Cells were stimulated for 72 hrs using antiCD3/CD28 beads (Life Technologies) at a ratio of 1 bead per 2 cells. Telomere FISH
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Telomere FISH was performed using a PNA probe (Panagene, Daejeon, Korea) as described (Batista et al., 2011; Gu et al., 2012). Proliferating T cells were treated overnight with 0.2 µg/ml colcemid (Life Technologies), swollen in KCl buffer (12.3 mM HEPES, 0.53 mM EGTA, 64.4 mM KCl), fixed in methanol/acetic acid (3:1) and dropped onto glass slides. Metaphase spreads were rehydrated in PBS, fixed in 4% formaldehyde and dehydrated in an alcohol series. Slides were incubated with hybridization mixture (70% formamide, 10 mM NaHPO4, pH 7.4, 10 mM NaCl, 20 mM Tris buffer, pH 7.5), placed on a heating block at 80°C for 5 min to denature chromosomal DNA and incubated for 30 min to 2 hrs at room temperature with the PNA probe (0.05 µg/ml). After washing, slides were mounted with ProLong Gold Antifade with DAPI (Life Technologies) and analyzed with a Leica microscope (Leica, Heidelberg, Germany). To visualize MRE11A binding, metaphase spreads were first incubated with anti-MRE11A (Cell Signaling Technology, 4847P; overnight, 4°C) and secondary Alexa Fluor® 488 goat anti-rabbit IgG (H+L) antibody (Life Technologies, A-11008) for 1 hr. After washing, slides were dehydrated in an alcohol series, hybridization mixture was applied and slides were placed on a heat block (80°C) for 5 min followed by 1 hr incubation as above. In situ DNA hybridization for SADS For DNA hybridizations, cells were denatured by incubating at 70°C for 2 min in 70% formamide. Preparations were immediately dehydrated through cold 70%, 95%, and 100% ETOH for 5 min each and then air-dried. Cells were hybridized with directly labeled oligonucleotides and heated at 70°C to 80°C for 10 min. Cells were incubated at 37°C in a humidified chamber overnight (Lawrence et al., 1988). Oligo sequences were sat-II: Cy3 direct label 5′-ATTCCATTCAGATTCCATTCGATC-3′.
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Confocal microscopy Cells were fixed in 4% paraformaldehyde, permeabilized with 0.05% Triton X-100 and incubated with primary antibodies to 53BP1 (Cell Signaling Technology, 4937S) and TRF2 (Abcam, ab13579) at 4°C for overnight. Incubation with secondary antibodies was performed at room temperature for 1 hr using Alexa Fluor 488 labeled goat anti-rabbit (Life Technologies, A-11008) and Alexa Fluor 546 labeled goat anti-mouse antibodies (Life
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Technologies, A-21123). The images were analyzed using the LSM710 microscope system with the ZEN 2010 software (Carl Zeiss) and a 63× oil immersion objective (Carl Zeiss). Telomere length measurement Genomic DNA was extracted directly from CD4+ T cells using a Mini Genomic DNA Kit (IBI Scientific) according to manufacturer’s instructions. Telomere length was determined as described (Cawthon, 2002). The primer sequences were as follows: tel 1, 5′-GGTTTTTGAGGGTGAGGGTGAGGGTGAGGGTGAGGGT-3′; tel 2, 5′-TCCCGACTATCCCTATCCCTATCCCTATCCCTATCCCTA-3′; 36B4u, 5′-CAGCAAGTGGGAAGGTGTAATCC-3′; 36B4d, 5′-CCCATTCTATCATCAACGGGTACAA-3′
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Synovitis induction in chimeric mice
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NOD.Cg-PrkdcscidIl2rgtm1Wjl/SzJ (NSG) female mice (10 to 14 weeks old) (The Jackson Laboratory) were used as previously described (Seyler et al., 2005). Pieces of human synovial tissue were placed into a subcutaneous pocket on the upper dorsal midline. In this model, complete engraftment is reached within 1 week. Following engraftment, mice were injected intravenously with 10 million cells of CD45RO− PBMCs. Chimeric mice from the same litter and carrying the same synovial tissue were randomly assigned to one of two treatment arms: (A) vehicle (DMSO) control and (B) treatment with Mirin, 1 mg/kg/day. All treatments were delivered by daily intraperitoneal injection over a period of 9 days. For the MRE11A overexpression experiments, mice from the same litter were randomly assigned to two treatment arms and engrafted with aliquots of the same synovial tissue. CD45RO− PBMCs were prepared from RA patients and transfected with either a control plasmid (10 million cells transferred into group A chimeras) or a MRE11A overexpression plasmid (10 million cells transferred into group B chimeras). Transfected cells were rested for 24 hrs before the adoptive transfer. At the completion of the experiments, mice were sacrificed and synovial tissues were harvested and embedded in OCT (Tissue-Tek; Sakura Finetek) for histological studies or were shock frozen in liquid nitrogen for RNA extraction. Immunohistochemical staining
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Frozen tissue blocks were cut at 5 µm thickness and picked up on X-tra® Slides (Leica Biosystems). After air-drying for 30 min at room temperature, sections were immediately fixed with acetone at −20°C for 20 min. Slides were rehydrated in 1× PBS for 10 min. To quench endogenous peroxidase activity, sections were incubated with peroxidase reagent (3% H2O2 in 1× PBS) for 15 min and gently washed twice in 1× PBS for 5 min. Slides incubated with a primary antibody cocktail overnight at 2°C to 8°C. The cocktail contained both monoclonal mouse anti-human CD3, (Clone F7.2.38; 1:100, Dako) and anti-p16 ARC antibody (EP1551Y; 1:100, Abcam, ab51243). Upon finishing and rinsing, the secondary antibody cocktail [peroxidase anti-rabbit IgG (H+L) (Vector Laboratories, PI-1000) and alkaline phosphatase anti-mouse IgG (H+L) (Vector Laboratories, AP-2000)] was prepared and applied as recommended by manufacturer (Vector Laboratories) followed by the subsequent visualization of AP activity (Vector Red Alkaline Phosphatase Substrate Kit)
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(Vector Laboratories, SK-5100) and HRP activity (Vector DAB Peroxidase Substrate Kit) (Vector Laboratories, SK-4100). Stained tissues were mounted with hematoxylin (SigmaAldrich), dehydrated, covered and imaged via microscope. Statistical analysis Statistics were calculated using GraphPad Prism software (GraphPad Software). If not stated differently, a two-sided t-test was used to determine significance and p30 nuclei for each donor. Fluorescence intensity in arbitrary units (a.u.). Results are mean ± SEM. (C) Distribution of fluorescence intensity (a.u.) strata from 5 RA (red) and 5 control samples (black) quantified in >30 nuclei for each donor. (D) Telomeric ends were analyzed for
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abnormal structures. Examples of double signal, apposition, fusion, and signal-free ends are shown. (E) Distributions of telomeric phenotypes in 10 patients and 10 controls. 150–200 nuclei were examined in each sample. Percentages of each damage pattern are presented. (F) Naïve and memory CD4+ T cells were separated and placed under proliferative stress by repetitive polyclonal stimulation. Loss of telomeric sequences was measured by PCR and, in parallel, telomeric damage foci were analyzed by dual-color immunostaining with antibodies to the DNA damage protein 53BP1 and the telomeric shelterin TRF2. 53BP1/TRF2 colocalization coefficients and telomeric length shortening in individual samples are correlated. *P < 0.05, two-tailed Student’s t-test. See also Figure S1 and Figure S2.
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Figure 2. The aging profile of CD4+CD45RA+ T cells in RA
CD4+CD45RA+ T cells from RA patients and age-matched controls were stimulated for 72 hrs. All data are mean ± SEM. (A, B) CDKN2A, CDKN1A, and TP53 transcript and p16 protein levels were measured in 7 RA-control pairs by RT-PCR and flow cytometry, respectively. (C) Expression of the aging marker CD57 assessed by flow cytometry. (D) SADS foci analyzed by confocal microscopy. Nuclei were hybridized with a sat-II–specific probe (red) and satellite DNA signals were examined in a minimum of 50 nuclei in each sample. Examples of condensed and threadlike distended satellites are shown in inserts. (E)
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Individual nuclei were scored as SADS-positive, if the pericentromeric or centromeric satellite heterochromatin had lost its tight packaging and was extended longitudinally. Data are from 3 RA-control pairs. (F) Telomeres were uncapped by transfecting healthy CD4+CD45RA+ T cells with TERF2 siRNA oligonucleotides. CDKN2A and CDKN1A transcripts were measured by RT-PCR. Results are from 3 independent experiments. (G) Flow cytometry for p16 expression. Representative histograms from control and TERF2 siRNA transfected cells are shown. Fluorescence Minus One control (FMO) is superimposed as grey area. Mean fluorescence intensities (MFI) of p16 are from 3 independent experiments. (H) CD57 expression was assessed by flow cytometry in 3 independent experiments. Representative data from control and TERF2 siRNA transfected cells are shown.*P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s t-test. See also Figure S3.
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Figure 3. MRE11Alow T cells in RA patients
MRE11A protein expression was quantified by intracellular staining of PBMCs in a cohort of RA patients (RA) and age-matched controls (Con) with antibodies against MRE11A and lineage markers (CD4, CD45RA). (A, B) Flow cytometric measurement of MRE11A protein expression in relation to donor age for naïve CD4+CD45RA+ T cells and memory CD4+CD45RO+ T cells. (C) Representative histograms from naïve CD4+CD45RA+ T cells from a healthy individual, an untreated RA patient and a RA patient on therapy. (D) Mean ± SEM of MRE11A MFI are from 5 samples per group. (E, F) Localization of MRE11A to
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the telomere quantified by dual-color immunostaining with anti-MRE11A (green) and antiTRF2 (red) in resting naïve CD4+CD45RA+ T cells of 5 RA patients and 5 age-matched controls. DNA is marked with DAPI (blue). (E) Representative image of immunostaining for MRE11A. (F) Staining intensities for total nuclear MRE11A and MRE11A-TRF2 colocalization measured in >50 nuclei from each of 5 different donors. *P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s t-test. See also Figure S4.
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Figure 4. Genetic or pharmacologic inhibition of MRE11A induces T cell aging
(A) The impact of MRN insufficiency on telomeric stability was tested by transfecting CD4+CD45RA+ T cells from healthy individuals with control, MRE11A, NBN, or RAD50 siRNA oligonucleotides. 48 hrs later, cells were stained with anti-53BP1 and anti-TRF2. Representative images of 53BP1 (green) and TRF2 (red) after knockdown of individual DNA repair proteins. DNA is marked with DAPI (blue). Merged images show colocalization of 53BP1 and TRF2. (B) Staining intensities (a.u.) for total 53BP1 and 53BP1/TRF2 colocalization were measured in individual nuclei. Mean ± SEM values are indicated.
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Results are from 5 independent knockdown experiments. (C) T cells were treated with the MRE11A inhibitor Mirin. Damage foci were analyzed by staining for 53BP1 and for 53BP1/ TRF2 colocalization. Data are from 5 independent experiments. All data are mean ± SEM. (D, E) CDKN2A and TP53 transcripts were measured by RT-PCR after transfecting CD4+CD45RA+ T cells from healthy individuals with control, MRE11A, NBN, or RAD50 siRNA oligonucleotides for 48 hrs (D) or were treated with the MRE11A inhibitor Mirin (E). (F) Flow cytometry for CD57 expression. Representative data are from control and MRE11A siRNA transfected cells. Percentages of CD57 expressing cells from 3 independent experiments are presented as mean ± SEM. (G) T cells were treated with vehicle or the MRE11A inhibitor Mirin at the indicated doses and analyzed for CD57 expression by flow cytometry. Mean ± SEM from 3 independent experiments. (H, I) Sat-II DNA (red) hybridization in control and Mirin-treated T cells. Distention of satellite DNA in the nuclei of Mirin-treated T cells. Nuclei were scored for SADS as in Figure 2 in >40 nuclei per sample and quantified in 3 control and Mirin-treated samples. *P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s t-test. See also Figure S5.
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Author Manuscript Author Manuscript Author Manuscript Figure 5. Overexpression of MRE11A repairs telomeric damage and prevents T cell aging in RA
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Naïve CD4+CD45RA+ T cells from RA patients were transfected with control plasmids or a myc-MRE11A construct. After 48 hrs, transfection efficiency was monitored by qPCR (A) and Western blotting (B). The error bars in (A) represent the 95% confidence internal. (C) Representative images of 53BP1 (green) and TRF2 (red) after overexpression of control plasmids or myc-MRE11A. DNA is marked with DAPI (blue). Merged images show colocalization of 53BP1 and TRF2. (D) Staining intensities (a.u.) for 53BP1/TRF2 colocalization were measured in a minimum of 70 individual nuclei from 3 different patients. Mean ± SEM values are indicated. CDKN2A and TP53 transcript levels (E) and p16 protein levels (F) were measured in control (RA+Vec) and myc-MRE11A (RA +MRE11A) transfected cells from 3 different patients. Results are mean ± SEM. (G) Flow
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cytometry for p16 expression. Representative data are from cells with or without MRE11A overexpression. (H) Percentages of p16 expressing cells analyzed in 4 independent experiments are presented as mean ± SEM. *P < 0.05, **P < 0.01 and ***P < 0.001, twotailed Student’s t-test.
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Author Manuscript Author Manuscript Author Manuscript Figure 6. The nuclease MRE11A controls pro-arthritogenic effector functions
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Synovial inflammation was quantified by gene transcriptional analysis in tissue extracts or by immunohistochemical analysis of tissue sections. (A) TRB transcript levels measured by qPCR to assess T cell accumulation. (B) Representative sections of synovial tissues stained with anti-CD3 antibodies (brown). (C) Percentages of CD3+ T cells in randomly selected fields of synovial tissues sections presented as mean ± SEM. (D) TNFSF11, (E) TNF, IL6, IL1B and (F) TGFB1, IL10 mRNA expression measured by qPCR in tissue extracts. (G) Transcription analysis of the aging markers CDKN2A, CDKN1A, and TP53 by qPCR. (H) p16 (brown) and CD3 (red) were stained by dual-color immunohistochemistry in synovial tissue sections from vehicle and MRE11A inhibitor-treated chimeras. (I) Percentages of p16+CD3+ T cells in randomly selected fields of synovial tissue sections presented as mean ± SEM. (J) T cell migratory capacity was measured in Transwell migration assays in the absence of chemokine gradients. N=6 RA patient-control pairs; n=4 young (65 years) healthy individuals; n=4 healthy control T cells with or without Mirin treatment. All data are mean ± SEM.*P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s ttest.
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Figure 7. Restoring MRE11A expression in RA T cells prevents pro-arthritogenic effector function
Pairs of NSG mice were engrafted with human synovial tissue and assigned to two treatment arms. CD45ROneg PBMC (A–D) and CD45RAnegPBMC (E–I) were prepared from RA patients and were transfected with either control plasmid or plasmid expressing MRE11A, and adoptively transferred into the chimeric mice. The intensity of synovial inflammation was compared by tissue gene expression analysis applying qPCR. (A, E and G) The density of the synovial T cell infiltrate was captured by TRB and TNFSF11 transcript levels. (B, C
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and H) Synovial cytokine production capability was assessed through TNF, IL6, IL1B, TGFB1 and IL10 transcript expression. (D, I) The impact of MRE11A overexpression on the tissue presence of aging markers was examined through CDKN2A and TP53 transcript levels. (F) Representative sections of synovial tissues stained with anti-CD3 antibodies (brown). Original magnification, X600 (insets in F). All data are mean ± SEM from at least 6 different synovial grafts.*P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s ttest. See also Figure S6.
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Immunity. Author manuscript; available in PMC 2017 October 18. Published in final edited form as: Immunity. 2016 October 18; 45(4): 903–916. doi:10.1016/j.immuni.2016.09.013.
Deficient activity of the nuclease MRE11A induces T cell aging and promotes arthritogenic effector functions in patients with rheumatoid arthritis Yinyin Li1, Yi Shen1, Philipp Hohensinner1,2, Jihang Ju1, Zhenke Wen1, Stuart B. Goodman3, Hui Zhang1, Jörg J. Goronzy1, and Cornelia M. Weyand1,4
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1Division
of Immunology and Rheumatology, Stanford University School of Medicine, Stanford, CA
94305 2Department
of Internal Medicine II/Cardiology, Medical University of Vienna, Vienna, Austria
3Department
of Orthopedic Surgery and Bioengineering, Stanford University School of Medicine, Stanford, CA 94305
Abstract
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Immune aging manifests with a combination of failing adaptive immunity and insufficiently restrained inflammation. In patients with rheumatoid arthritis (RA), T cell aging occurs prematurely, but the mechanisms involved and their contribution to tissue-destructive inflammation remain unclear. We found that RA CD4+ T cells showed signs of aging during their primary immune responses and differentiated into tissue-invasive, pro-inflammatory effector cells. RA T cells had low expression of the double-strand-break repair nuclease MRE11A, leading to telomeric damage, juxtacentromeric heterochromatin unraveling, and senescence marker upregulation. Inhibition of MRE11A activity in healthy T cells induced the aging phenotype, whereas MRE11A overexpression in RA T cells reversed it. In human-synovium chimeric mice, MRE11Alow T cells were tissue-invasive and pro-arthritogenic, and MRE11A reconstitution mitigated synovitis. Our findings link premature T cell aging and tissue-invasiveness to telomere deprotection and heterochromatin unpacking, identifying MRE11A as a therapeutic target to combat immune aging and suppress dysregulated tissue inflammation.
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Correspondence To: Cornelia M. Weyand, M.D., Ph.D., Division of Immunology and Rheumatology, Department of Medicine, Stanford University, CCSR Building Room 2225, MC-5166, 269 Campus Drive West, Stanford, CA 94305; Phone: (650) 723-9027, Fax: (650) 721-1251, [email protected]. 4Lead Contact Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain. COMPETING FINANCIAL INTERESTS The authors declare that no conflict of interest exists. Additional experimental procedures are presented in Supplemental Materials. AUTHOR CONTRIBUTIONS YL, JJG and CMW designed the research and analyzed data. YL, YS, PH, JJ, ZW and HZ were responsible for the experimental work. SG recruited patients. CMW, YL and JJG wrote the manuscript.
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INTRODUCTION Preceded by a decade-long period of preclinical disease, rheumatoid arthritis (RA) manifests with a symmetrical polyarthritis causing irreversible cartilage and bone destruction and shortens life expectancy due to accelerated cardiovascular disease. Immune aging affects the general population after 50 years of age, but is accelerated in RA patients (Weyand et al., 2009), where it is already noticeable in antigen-unprimed naïve T cells (Koetz et al., 2000).
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Cells devote a significant proportion of their machinery to DNA surveillance and repair to prevent cellular aging or death associated with genome instability (Chow and Herrup, 2015). Predictable loss of telomeric sequences with each cell replication allows telomeres to serve as molecular clocks. By tallying the number of cell divisions, telomeres are believed to effectively force mutation-harboring cells into cell cycle arrest. Senescent T cells not only remain viable, but actively shape the tissue microenvironment by secreting cytokines and tissue remodeling factors (Weyand et al., 2014). However, despite several senescence features, aging human lymphocytes are not in replicative arrest (Yang et al., 2016) and continue to participate in clonal expansion, distinguishing lymphocyte aging from senescence (Akbar and Henson, 2011; Chou and Effros, 2013; Sharpless and Sherr, 2015). Reversibility of senescence in human end-differentiated effector T cells further supports the model that aging of lymphocytes reflects progressive differentiation more than true senescence (Di Mitri et al., 2011). Whether aging T cells acquire effector functions that mediate tissue inflammation is not understood.
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Abnormalities in the DNA damage sensing and repair machinery of RA T cells have raised the question of whether such defects are mechanistically linked to T cell aging and to arthritogenic effector functions (Shao et al., 2009; Shao et al., 2010). The MRN complex, composed of Meiotic Recombination 11 Homolog A (MRE11A), RAD50 and Nijmegen Breakage Syndrome 1 (NBS1), senses DNA double-strand breaks to amplify DNA repair (Lamarche et al., 2010). The core component of the complex, MRE11A, has doublestranded (ds)DNA exonuclease and single-stranded (ss)DNA endonuclease activity in both homologous recombination and nonhomologous end-joining (Xie et al., 2009). MRE11A is recruited to healthy telomeres, where its function is not understood. In S. cerevisiae, RAD50 or MRE11A loss shortens telomeres and abolishes S phase telomerase recruitment (Takata et al., 2005). In high eukaryotes, MRE11A specifically interacts with the shelterin protein TRF2 (Diotti and Loayza, 2011). Conversely, MRN complex hypomorphism in mouse embryonic fibroblasts does not affect telomere length (Attwooll et al., 2009) and reduces telomere dysfunction-induced foci (TIF), revealing the complexity of MRE11A function. Which role MRE11A plays in healthy and stressed human cells, particularly in long-lived lymphocytes, is unknown. Here, we report that in RA T cells, age-related telomeric defects manifested not only as shortening, but also as damage accumulation. Such T cells unfolded higher-order satellite heterochromatin, expressed increased levels of the cell cycle regulators p16 and p21 but lacked p53. Aged RA T cells were low-expressers for the DNA repair nuclease MRE11A and their chromosomal ends were MRE11Alow. In healthy T cells, decreasing MRE11A mRNA by treatment with short interfering RNAs (siRNAs) or pharmacologic inhibition of
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MRE11A’s nucleolytic activity promptly induced telomeric damage and upregulated the senescence markers p16, p21, and CD57, concomitant with unraveling of pericentromeric satellite DNA. Spontaneous or induced deficiency of MRE11A’s nucleolytic function had a profound impact on T cell behavior and rendered T cells tissue-invasive and pro-arthrogenic, whereas reconstitution of MRE11A protein in patient-derived T cells protected synovial tissue from inflammatory attack. These data provide mechanistic evidence for a role of the MRE11A nuclease in not only regulating aging but also differentiation of T cells into tissueinjurious effector cells.
RESULTS Telomeres in RA T cells are not only shortened, but damaged
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Telomeric sequences in RA T cells are shortened relative to T cells from age-matched healthy individuals, and this has been attributed to increased proliferative pressure in an inflammatory environment (Koetz et al., 2000). However, T cell turnover measured by the proliferation marker Ki-67 correlates inversely with telomeric erosion (Schonland et al., 2003), suggesting alternative mechanisms underlying telomeric loss. Telo-FISH staining in metaphase nuclei confirmed that the vast majority of RA naïve CD4+ T cells had lowintensity telomeric signals (Figure 1A and 1B). Compared to healthy individuals, RA patients lacked high-brightness nuclei and almost all of their cells had a weak telomere signal (Figure 1C).
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The structural intactness of telomeric caps was examined through FISH-probe hybridization patterns in metaphase-arrested nuclei, revealing four structural patterns: telomeric apposition (long chromosomal arms opposed), telomeric fragmentation (signal doubling), telomeric loss (signal-free end) and telomeric fusion (physical linkage of two ends of the same or neighboring chromosomes) (Figure 1D). The majority of healthy metaphase T cell nuclei had structurally intact telomeres (56.5%). Thirty-one percent of nuclei displayed signal doublets and 12.5% displayed telomeric apposition. In contrast, RA T cell nuclei consistently contained damaged telomeres. End signal doubling, indicative of telomeric fragility, occurred in 41.9% of cells and 24.1% had opposed telomeres. Albeit encountered infrequently, RA samples showed evidence for severe structural damage, including fusions (2.8%) and total signal loss (1.6%) (Figure 1E).
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To understand the relationship between proliferation-induced telomeric shortening and TIF formation, we monitored TIF evolution in polyclonally expanding T cell populations by quantifying the recruitment of the DNA damage protein P53-Binding Protein 1 (53PB1) to telomeric ends marked through the shelterin protein TRF2. Naïve and memory CD4+ T cells from the same donor were placed under proliferative stress (Figure S1A) by repetitive polyclonal stimulation. Enforced proliferation markedly shortened telomeres in the naïve population, but lengths were relatively stable in memory counterparts, despite similar proliferation rates in both T cell subpopulations (Figure S1A, S1B, S1D). Naïve CD4+ T cells lost >1000 telomeric nucleotides over 3 weeks, but memory CD4+ T cell chromosomes were shortened by 300 base pairs (bp). Parallel monitoring of TIF by quantifying 53BP1 and TRF2 colocalization demonstrated rare foci in naïve CD4+ T cells, even after >1000 bp loss. In contrast, TIF appeared as early as day 7 in memory CD4+ T cells (Figure S1C). Overall, Immunity. Author manuscript; available in PMC 2017 October 18.
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53BP1 and TRF2 colocalization and erosion of telomeric sequences were unrelated (Figure 1F). Together, these data documented that telomeres in RA T cells are shortened, but more importantly, are sites of DNA damage. Premature aging of naïve CD4+ T cells in RA patients is related to telomeric damage
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The presence of circulating and synovial CD4+CD28null T cells in RA patients provides unequivocal evidence for in vivo T cell aging. Such end-differentiated T cells express CD57 and natural killer (NK) cell receptors (Warrington et al., 2001). To better define the aging process in naïve CD4+CD45RA+ T cells, we assessed the aging-associated cell cycle inhibitors p16, p21, and p53 (Sharpless and Sherr, 2015). Compared to healthy controls, RA T cells expressed higher levels of CDKN2A and CDKN1A mRNA and p16 protein (Figure 2A–B), while TP53 expression was reduced, suggesting a p16-dependent and p53independent aging pathway (Figure 2A). The glucuronyltransferase gene family member CD57 is considered a senescence marker on T cells (Focosi et al., 2010). The frequency of CD4+CD45RA+CD57+ T cells was low in healthy individuals (7%), but more than doubled in RA patients (20%) (Figure 2C).
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DNA condensation into senescent-associated heterochromatin foci is a hallmark of cellular senescence, but early stages of cellular aging can be associated with the unravelling of constitutive heterochromatin into visibly extended structures (Swanson et. al., 2013). To identify heterochromatin changes, we analyzed CD4+CD45RA+ T cell populations for evidence of unraveling of pericentromeric and centromeric satellite DNA. Hybridization of a probe to satellite-II (sat-II) DNA in stimulated healthy CD4+CD45RA+ T cells yielded compact, round and centric satellite signals (Figure 2D). Conversely, RA T cells had distended pericentromeric and centromeric satellites, usually affecting multiple sites within a nucleus. Eighty percent of RA cells had clearly unraveled satellites. In contrast, 80% of control cells had tightly packaged condensed satellites (Figure 2E), demonstrating that pericentromeric and centromeric satellite heterochromatin in RA T cells undergoes decondensation, consistent with entry into early senescence. To investigate whether aging-associated T cell phenotypes are mechanistically linked to telomeric damage, we used siRNAs to decrease telomere-protecting shelterin TERF2 mRNA. Transfecting TERF2 siRNAs or a TERF2 mutant plasmid reduced TERF2 mRNA (Figure S3A); induced robust TIF formation (Figure S3B) and doubled CDKN2A and CDKN1A mRNA transcripts (Figure 2F, Figure S3C). TERF2 knockdown was sufficient to induce p16 and CD57 expression (Figure 2G–H), recapitulating conditions in RA patients. T cells from RA patients are MRE11A deficient
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The presence of telomeric damage foci and satellite DNA distention in RA T cells raised the question whether the DNA repair machinery was fully intact. We examined RA T cells for the expression of key molecules involved in the DNA double strand break repair, including Ataxia Telangiectasia and Rad3-Related Protein (ATR), Ataxia Telangiectasia Mutated (ATM), MRE11A, NBS1, RAD50 and DNA-dependent protein kinase catalytic subunit (DNA-PKcs) by flow cytometry. Intracellular staining of peripheral blood mononuclear cells (PBMCs) demonstrated reduced MRE11A protein expression in naïve and memory CD4+ T
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cells from RA patients (Figure 3A–B). In both T cell subpopulations, MRE11A protein expression declined in an age-dependent fashion in RA patients as well as healthy donors. However, MRE11A protein was consistently lower in RA patients’ cells; reduced by 40– 50% in the naïve population and 35% in the memory population. CD4+ T cells of patients with the inflammatory polyarthritis, PsA, were indistinguishable from healthy T cells (Figure S4A). In RA patients, but not in PsA patients, CD8+ T cells, CD14+ monocytes and CD19+ B cells were also MRE11Alow expressors (Figure S4B, S4C, S4D). MRE11A lowexpression was most pronounced in T cells from untreated RA patients, eliminating immunosuppressive therapy as the causative factor (Figure 3C–D). Total nuclear and telomere-bound MRE11A was analyzed by dual-color immunostaining with anti-MRE11A and anti-TRF2. Staining intensity for MRE11A protein was markedly reduced in RA T cell nuclei, in which the signal for MRE11A and TRF2 colocalization was especially low, suggesting that MRE11Alow RA T cells have telomeric tips almost depleted of the nuclease (Figure 3E–F). Thus, aging is associated with progressive decline in the expression of the double strand break repair nuclease MRE11A, a process which is notably accelerated in RA T cells. Diminished MRE11A activity causes telomeric damage
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The exonuclease and endonuclease MRE11A partners with the ATPase RAD50 and NBS1 to form the MRN complex, a DNA break sensor critically involved in multiple repair pathways (Lamarche et al., 2010). siRNA-mediated knockdown of each of the three MRN components, MRE11A, RAD50 and NBS1, produced partial depletion of both transcripts and protein (Figure S5A–S5B) and induced widespread DNA damage (Figure 4A). The amount of unrepaired DNA breaks was quantified by measuring the intensity of 53BP1binding to damaged chromatin sites, where this checkpoint protein forms foci and regulates end-joining-mediated repair processes. MRE11A loss was most devastating for DNA integrity; 53BP1 was recruited to numerous DNA break sites in almost every single nucleus. In contrast, NBS1 knockdown left some nuclei unaffected. Partial loss of RAD50 was tolerated the best, producing the lowest 53BP1 signal. 53BP1 and TRF2 colocalization studies confirmed the functional relevance of MRE11A at the telomere (Figure S5C). Overall, transfection of MRE11A-specific siRNA produced robust DNA damage, with almost 50% of the 53BP1 signal localizing to telomeric ends. Only one-third of the 53BP1 signal copositioned with the shelterin protein TRF2 in NBS1 or RAD50 insufficient cells (Figure 4B). Alternatively, the function of MRE11A was impaired by treating with the small molecule inhibitor Mirin, which inhibits MRE11A’s exonuclease activity while sparing the endonuclease activity (Dupre et al., 2008; Shibata et al., 2014). Inhibition of exonuclease activity in healthy naïve CD4+ T cells resulted in massive DNA damage, of which a substantial proportion localized to the telomere (Figure 4C and Figure S5D). These data confirmed that the nucleolytic activity of MRE11A is particularly important in protecting telomeric integrity. Impairing MRE11A activity induces the aging profile of RA T cells To investigate whether MRE11Alow T cells are prone to accelerate the aging process, we targeted MRE11A, RAD50 and NBS1 with interfering RNA technology. MRE11A knockdown potently induced CDKN2A, NBS1 knockdown resulted in a significant, but Immunity. Author manuscript; available in PMC 2017 October 18.
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small, increase in CDKN2A mRNA and RAD50 knockdown left CDKN2A expression unchanged. TP53 expression was unaffected by reducing MRE11A expression (Figure 4D). Treatment with the MRE11A inhibitor Mirin led to similar results (Figure 4E), indicating that MRE11A’s nucleolytic activity protected T cells from entering the aging program. Overall, the pattern of p16 induction paralleled the degree of telomeric damage induced by the genetic inhibition of the three molecules (Figure 2), with MRE11A trumping the two other components in TIF induction and p16 upregulation. Weaning of the inhibitory effects of the siRNAs and recovery of MRE11A expression were sufficient to reverse the upregulation of CDKN2A (encoding p16) and CDKN1A (encoding p21) (Figure S5E–S5F). CD57 expression in MRE11A siRNA-treated cells (Figure 4F) recapitulated the spontaneous aging profile in patient-derived T cells (Figure 2). Similarly, Mirin treatment more than doubled the frequency of CD57+CD4+ T cells (Figure 4G) and rapidly induced heterochromatin unpacking (Figure 4H). More than 80% of Mirin-treated cells had distended satellites, reproducing the constitutive satellite DNA unraveling in RA T cells (Figure 4I). These experiments mechanistically linked the nuclease MRE11A to the aging phenotype of T cells, including the upregulation of cell cycle regulators and the unraveling of satellite chromatin. Restoring MRE11A expression repairs telomeric damage and prevents T cell aging
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To examine whether MRE11A restoration is sufficient to revert the aging phenotype, an MRE11A construct was overexpressed in RA CD4+ naïve T cells (Figure 5A and 5B). Increasing the MRE11A protein concentration prevented telomeric 53BP1 recruitment; 53BP1 and TRF2 colocalization declined by 75% (Figure 5C and 5D), suggesting a direct role of MRE11A in telomere repair. Other components of the T cell aging program, such as p16 expression, were similarly affected (Figure 5E–5G). Sixty percent of control-transfected RA T cells were p16pos and frequencies were reduced to 20% after MRE11A overexpression (Figure 5H). Consistent with the data in Figure 4D, p53 expression appeared independent of MRE11A protein concentrations. These data confirmed that MRE11A has anti-aging function by securing telomeric intactness and controlling cell cycle regulators and that the aging signature of RA T cells is reversible. Inhibiting MRE11A’s nucleolytic activity renders T cells tissue-invasive and proinflammatory
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CD4+CD28null T cells participate in the inflammatory lesions in RA joints (Schmidt et al., 1996), but whether their accelerated aging is mechanistically connected to their proinflammatory functions is unknown. To establish tissue inflammation, T cells must migrate into the tissue site where they display proinflammatory effector functions, such as cytokine and chemokine release, activation of endothelial cells, stromal cells and innate immune cells. All phases of this process are captured in a humanized mouse model in which human synovial tissue is engrafted into immunodeficient mice and human CD45RO− PBMCs, either deriving from healthy individuals or from patients with RA, are adoptively transferred. Chimeric mice were reconstituted with either control or RA PBMCs and were treated with the MRE11A inhibitor Mirin or vehicle (Figure 6). Adoptively transferred PBMCs from
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healthy individuals formed sparse T cell infiltrates in the synovial grafts (Figure 6A). Inhibition of MRE11A’s nucleolytic activity enhanced T cell accumulation in the synovial tissue. MRE11Alow RA PBMCs spontaneously had a much higher degree of tissue invasiveness. Here, MRE11A inhibition marginally enhanced T cell recruitment into the synovium. Synovial T cell accumulation was similar in Mirin-treated control PBMCs and untreated RA PBMCs (Figure 6B–C). Inhibiting MRE11A activity resulted in upregulated expression of TNFSF11 (encoding RANKL), a molecule critically involved in rheumatoid bone destruction (Figure 6D). Also, the intensity of tissue inflammation was increased, as measured by the expression of the pro-inflammatory cytokines TNF, IL6 and IL1B (Figure 6E). Conversely, reducing MRE11A activity prompted increased tissue expression of the regulatory cytokines TGFB1 and IL10 (Figure 6F). Exposure of synovial tissue to the MRE11A inhibitor in the absence of transferred PBMCs had no impact on cytokine expression (Figure 6E–F).
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To examine a possible link between tissue invasiveness, inflammatory capability and cellular aging, we quantified tissue expression of CDKN2A, CDKN1A and TP53 transcripts and measured p16 protein expression by dual-color immunohistochemistry (Figure 6G–I). Mirin treatment left the cell cycle regulator expression in noninflamed synovial cells unaffected, but promptly induced the CDKN2Ahigh, CDKN1Ahigh, TP53low profile in tissue-infiltrating cells (Figure 6G). Loss of MRE11A function rendered T cells and non-T cells p16-positive (Figure 6H–I). RA T cells formed dense synovial infiltrates, with a visible enrichment of CD3+ p16+ cells. Also, MRE11Alow RA T cells had a hypermigratory phenotype in Transwell assays performed in the absence of chemokines (Figure 6J). Hypermotility was shared by T cells from old individuals and T cells treated with the MRE11A inhibitor Mirin (Figure 6J). Overall, healthy T cells with inhibited MRE11A nuclease activity resembled RA T cells. These experiments assigned a mechanistic role to the nuclease MRE11A in several disease-relevant capabilities of RA T cells; specifically, their tissue-invasive and aggressive inflammatory behavior. Restoring MRE11A in RA cells prevents pro-arthritogenic effector functions
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To test whether restoring MRE11A expression was sufficient to correct the pathogenic behavior of RA CD4+ T cells, we overexpressed MRE11A in naïve and memory CD4+ T cells prior to adoptive transfer. Forced MRE11A overexpression was strongly antiinflammatory (Figure 7). Transfer of MRE11Ahigh RA T cells minimized tissue TRB transcript levels, diminished the density of the T cell infiltrate, reduced TNF, IL6, IL1B expression, increased TGFB1 and IL10 mRNA and prevented CDKN2A induction. Correcting MRE11A protein concentrations in the transferred T cells did not affect T cell lineage commitment, as indicated by the expression of the T cell transcription factors TBX21, GATA3, RORC and FOXP3 (Figure S6). These experiments confirmed that MRE11A is a critical regulator of pathogenic effector functions in tissue-residing RA T cells and that restoring MRE11A protein expression is sufficient to modify arthritogenic capabilities in naïve and memory CD4+ T cells.
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DISCUSSION
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RA is an HLA class II-associated autoimmune disease, in which arthritogenic T cells drive progressive inflammatory destruction of cartilage and bone. Patients with RA have a signature of premature immune aging, providing an excellent model system to study the interrelationship between failing T cell immunity and unremitting tissue inflammation. Here, we present data implicating defective DNA damage responses in accelerating T cell aging, which renders such T cells susceptible to differentiate into tissue-invasive, pro-athrogenic effector cells. Specifically, low expression of the double strand break repair nuclease MRE11A in RA T cells diminishes dsDNA exonucleolytic and ssDNA endonucleolytic activity, which appears critically important in maintaining intact telomeres, preventing entry into the aging program and averting tissue invasion of inflammation-biased T cells. These findings support the concept that telomeric erosion in MRE11Alow T cells is a consequence of insufficient repair and not proliferative stress and that telomeric damage is mechanistically linked to cellular behavior.
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In this study, we defined the T cell aging program in RA, which is distinct from cellular senescence (Chou and Effros, 2013). How T cells age and whether immune cells follow a “universal” aging program is insufficiently understood, but RA T cells and T cells from aged humans lack the irreversible cell cycle arrest that is considered a cardinal feature of senescence (Goronzy and Weyand, 2013; Sharpless and Sherr, 2015). In contrast, cell cycle passage of RA T cells is shortened and the entire population of patient-derived CD4 T cells is hyperproliferative (Fujii et al., 2009). Synovial T cells have all converted into effector memory T cells, but naïve CD4+ T cells in RA already display abnormalities, e.g., due to increased apoptotic susceptibility clonal expansion is relatively futile (Schonland et al., 2003). Long before differentiating into effector memory cells, RA T cells fail to repair DNA damage (Shao et al., 2009), are energy deprived (Yang et al., 2013) and chronically activate the DNA-PKcs-JNK pathway (Shao et al., 2010). The downregulation of MRE11A was a global phenomenon in RA T cells affecting the entire population and was not selective for a subpopulation. Flow cytometric quantification of MRE11A protein produced unimodal distribution patterns in both CD45RA+ and CD45RA− populations, indicating a broadly distributed defect amongst T cells. MRE11A overexpression rescued the pathogenic effector functions of both, naïve CD45RA+ and memory CD45RO+ CD4+ T cells, when adoptively transferred into chimeric mice. Experiments with isolated CD45RA+CD4+ T cells allowed focusing on early phases in the life cycle of T cells, as they begin to make the transition into effector and memory T cells, that eventually mediate the pathogenic functions in the tissue environment. The human CD8+, but not the CD4+ T cell compartment contains small populations of T cells that increase in frequency with age and that can reacquire a naïve-like phenotype after in vivo stimulation while maintaining the ability to rapidly respond to restimulation (Fuertes Marraco et al., 2015; Pulko et al., 2016). In healthy humans, such CD8 naïve-like T cells with effector function have relatively long telomeres, longer than memory T cells, indicating that they do not have a defect in telomeric maintenance. Cellular senescence, genomic instability, epigenetic alterations, stem cell exhaustion, loss of proteostasis, mitochondrial dysfunction and deregulated nutrient sensing are all critical components in organismal aging (Lepez-Otin et al., 2013). Current data defined the T cell
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aging signature as p16high, p21high and p53low. p16INK4A accumulation is characteristic of aging in other lymphoid and nonlymphoid cells (Chen et al., 2011; Cosgrove et al., 2014; Janzen et al., 2006; Liu et al., 2011; Molofsky et al., 2006; Signer et al., 2008; Sousa-Victor et al., 2014). More difficult to understand is the p21-gain and p53-loss combination, as DNA-damage-induced ATM-p53-p21 signaling typically drives growth arrest of senescent T cells (Bartkova et al., 2006; Fumagalli et al., 2014; Nakamura et al., 2008; Rodier et al., 2011). However, in cancer cells p21 expression can occur in the absence of p53 function (Jeong et al., 2010; Macleod et al., 1995). In RA T cells, p53 is consistently low (Maas et al., 2005; Shao et al., 2009), associating aging with a p53-independent, p16-dependent mechanism. Naïve RA CD4+ T cells include a considerable population of CD57+ cells. How CD57 alters the behavior of old versus young CD4+ T cells is unknown, but it could possibly affect tissue homing.
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Epigenetic changes to chromatin are essential to cellular aging. Senescence-associated distension of satellites (SADS) formation is a unifying process in most models of early senescence, believed to signify the loss of higher-order chromatin packaging (Swanson et al., 2013). The packaging of pericentromeric and centromeric satellite DNA, especially satII DNA, in RA T cells was visibly altered. Whether distensions in sat-II DNA are associated with transcription at these loci will need examination. So far, pericentromeres are considered transcriptionally inert chromatin conformations, but recent evidence emphasizes active and regulated transcription, especially in the context of development, tumorigenesis, senescence and cellular stress (Dejardin, 2015; Saksouk et al., 2015). MRE11A inhibition induced pericentromere unraveling, linking satellite packaging with DNA repair and emphasizing the nuclease’s role in protecting the epigenetic landscape. The MRN complex is a multifunctional enzyme conglomerate known to activate cell cycle checkpoints when sensing DNA breakage (Stracker and Petrini, 2011). In addition to a critical role in ATM function and double strand break repair, MRN is also implicated in suppression of lymphomagenesis (Balestrini et al., 2015) and regulating metastatic breast cancers (Gupta et al., 2013). In humans, MRE11A mutations cause an ataxia-telangiectasia-like disease, with considerable variability in clinical phenotypes (Taylor et al., 2015). In primary human T cells, MRE11A appears to have primarily a protective function in telomere maintenance. Reducing MRE11A abundance by 50% rapidly induced the formation of telomeric damage foci, long before such T cells shortened telomeric length. MRE11A deficiency was more effective in inducing telomeric damage foci than the knockdown of RAD50 or NBS1. Vice versa, replenishing MRE11A protein by forced overexpression promptly reversed telomeric damage. These data suggest MRE11A’s participation in telomere repair and protection in human T cells.
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MRE11A is an abundant protein in human T cells. A 50% reduction, however, had profound consequences, revealed by the reversal of several phenotypes after MRE11A overexpression. Although overexpression prevented TIF formation and suppressed p16, p21 and CD57 expression, the mechanistic connection between telomeric damage and T cell aging awaits further clarification. Pharmacological inhibition of MRE11A’s nucleolytic activity recapitulated all abnormalities of aged T cells, placing insufficient exonuclease activity at the pinnacle of the T cell aging program. Systemic inflammation was insufficient to explain
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MRE11A downregulation. In PsA, an aggressive multi-organ inflammatory syndrome, MRE11A expression was normal. Untreated RA patients had the lowest MRE11A concentrations; immunosuppressive therapy improved but did not normalize the nuclease’s expression, implicating mechanisms other than systemic inflammation in MRE11A suppression.
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In previous reports, the failure to degrade cytoplasmic DNA induced inflammatory responses in bone marrow macrophages to eventually induce polyarthritis (Kawane et al., 2006). Current data link DNA damage in T cells, not macrophages, to functional abnormalities driving synovitis. Mechanistically, MRE11A’s nuclease activity appears to be related to in vivo T cell trafficking and T cell control of the synovial microenvironment. Human synovium and human T cells in chimeric mice enabled us to investigate how insufficient MRE11A activity regulates nonlymphoid tissue infiltration and inflammation. RA T cells have been reported to be spontaneously hypermigratory (Hwang et al., 2015) and we found that nuclease inhibition rendered them hyper-motile and tissue-invasive. Once in the tissue, MRE11Alow T cells supported p16 and p21 expression, left p53 unaffected and greatly increased the abundance of classical pro-inflammatory cytokines. MRE11A restoration did not change T cell lineage commitment but appeared to regenerate an antiinflammatory tissue environment, suggestive for a role of T cells in regulating joint-residing cells, such as synovial fibroblasts.
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The current study has implication for the understanding of the immune aging process in humans and could change the approach towards treating senescence-associated tissue inflammation. Experimental data focused attention on DNA damage sensing and repair as a key event in immune aging, particularly in RA. Impaired DNA damage responses affected T cells in their naïve state and fundamentally change their differentiation program, biasing them towards tissue-invasive and pro-inflammatory behavior. Inflammation per se, as in PsA patients, was insufficient to repress MRE11A expression and the associated T cell aging profile. MRE11A insufficiency appeared involved in multiple aspects of T cell aging, from unraveling of heterochromatin, to telomeric damage, changed expression of cell cycle regulators and altered tissue homing. Restoring MRE11A expression was sufficient to reverse aging features of RA T cells, identifying DNA damage repair, and particularly MRE11A, as a promising therapeutic target to treat immune-aging related disease.
EXPERIMENTAL PROCEDURES Patients and control individuals
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RA patients recruited into the study fulfilled the 2010 diagnostic criteria for RA and were positive for anti-CCP antibodies and for rheumatoid factor (Table S1). Healthy controls were matched for age and gender. The following were exclusion criteria for their enrollment: personal or family history of autoimmune disease, malignancy, chronic infection or another inflammatory syndrome. The Institutional Review Board approved the study and written informed consent was obtained from all participants.
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T cell purification and culture
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Peripheral blood mononuclear cells (PBMCs) were separated from whole blood using Lymphocyte Separation Medium (Corning). Naïve CD4+ T cells were isolated from PBMCs using a human naïve CD4+ T cell isolation kit according to the manufacturer’s instructions (STEMCELL Technologies). This purification procedure minimizes the contamination of CD4+CD45RA+ cells with end-differentiated CD4+CD28− T cells, which accounted for 0.4% of cells in both patients and controls (Figure S2). Naïve CD4+ T cells were cultivated in RPMI 1640 medium (Fisher Scientific) supplemented with 10% FBS (Atlanta Biologicals) and Pen/Strep (Fisher Scientific). Cells were stimulated for 72 hrs using antiCD3/CD28 beads (Life Technologies) at a ratio of 1 bead per 2 cells. Telomere FISH
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Telomere FISH was performed using a PNA probe (Panagene, Daejeon, Korea) as described (Batista et al., 2011; Gu et al., 2012). Proliferating T cells were treated overnight with 0.2 µg/ml colcemid (Life Technologies), swollen in KCl buffer (12.3 mM HEPES, 0.53 mM EGTA, 64.4 mM KCl), fixed in methanol/acetic acid (3:1) and dropped onto glass slides. Metaphase spreads were rehydrated in PBS, fixed in 4% formaldehyde and dehydrated in an alcohol series. Slides were incubated with hybridization mixture (70% formamide, 10 mM NaHPO4, pH 7.4, 10 mM NaCl, 20 mM Tris buffer, pH 7.5), placed on a heating block at 80°C for 5 min to denature chromosomal DNA and incubated for 30 min to 2 hrs at room temperature with the PNA probe (0.05 µg/ml). After washing, slides were mounted with ProLong Gold Antifade with DAPI (Life Technologies) and analyzed with a Leica microscope (Leica, Heidelberg, Germany). To visualize MRE11A binding, metaphase spreads were first incubated with anti-MRE11A (Cell Signaling Technology, 4847P; overnight, 4°C) and secondary Alexa Fluor® 488 goat anti-rabbit IgG (H+L) antibody (Life Technologies, A-11008) for 1 hr. After washing, slides were dehydrated in an alcohol series, hybridization mixture was applied and slides were placed on a heat block (80°C) for 5 min followed by 1 hr incubation as above. In situ DNA hybridization for SADS For DNA hybridizations, cells were denatured by incubating at 70°C for 2 min in 70% formamide. Preparations were immediately dehydrated through cold 70%, 95%, and 100% ETOH for 5 min each and then air-dried. Cells were hybridized with directly labeled oligonucleotides and heated at 70°C to 80°C for 10 min. Cells were incubated at 37°C in a humidified chamber overnight (Lawrence et al., 1988). Oligo sequences were sat-II: Cy3 direct label 5′-ATTCCATTCAGATTCCATTCGATC-3′.
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Confocal microscopy Cells were fixed in 4% paraformaldehyde, permeabilized with 0.05% Triton X-100 and incubated with primary antibodies to 53BP1 (Cell Signaling Technology, 4937S) and TRF2 (Abcam, ab13579) at 4°C for overnight. Incubation with secondary antibodies was performed at room temperature for 1 hr using Alexa Fluor 488 labeled goat anti-rabbit (Life Technologies, A-11008) and Alexa Fluor 546 labeled goat anti-mouse antibodies (Life
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Technologies, A-21123). The images were analyzed using the LSM710 microscope system with the ZEN 2010 software (Carl Zeiss) and a 63× oil immersion objective (Carl Zeiss). Telomere length measurement Genomic DNA was extracted directly from CD4+ T cells using a Mini Genomic DNA Kit (IBI Scientific) according to manufacturer’s instructions. Telomere length was determined as described (Cawthon, 2002). The primer sequences were as follows: tel 1, 5′-GGTTTTTGAGGGTGAGGGTGAGGGTGAGGGTGAGGGT-3′; tel 2, 5′-TCCCGACTATCCCTATCCCTATCCCTATCCCTATCCCTA-3′; 36B4u, 5′-CAGCAAGTGGGAAGGTGTAATCC-3′; 36B4d, 5′-CCCATTCTATCATCAACGGGTACAA-3′
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Synovitis induction in chimeric mice
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NOD.Cg-PrkdcscidIl2rgtm1Wjl/SzJ (NSG) female mice (10 to 14 weeks old) (The Jackson Laboratory) were used as previously described (Seyler et al., 2005). Pieces of human synovial tissue were placed into a subcutaneous pocket on the upper dorsal midline. In this model, complete engraftment is reached within 1 week. Following engraftment, mice were injected intravenously with 10 million cells of CD45RO− PBMCs. Chimeric mice from the same litter and carrying the same synovial tissue were randomly assigned to one of two treatment arms: (A) vehicle (DMSO) control and (B) treatment with Mirin, 1 mg/kg/day. All treatments were delivered by daily intraperitoneal injection over a period of 9 days. For the MRE11A overexpression experiments, mice from the same litter were randomly assigned to two treatment arms and engrafted with aliquots of the same synovial tissue. CD45RO− PBMCs were prepared from RA patients and transfected with either a control plasmid (10 million cells transferred into group A chimeras) or a MRE11A overexpression plasmid (10 million cells transferred into group B chimeras). Transfected cells were rested for 24 hrs before the adoptive transfer. At the completion of the experiments, mice were sacrificed and synovial tissues were harvested and embedded in OCT (Tissue-Tek; Sakura Finetek) for histological studies or were shock frozen in liquid nitrogen for RNA extraction. Immunohistochemical staining
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Frozen tissue blocks were cut at 5 µm thickness and picked up on X-tra® Slides (Leica Biosystems). After air-drying for 30 min at room temperature, sections were immediately fixed with acetone at −20°C for 20 min. Slides were rehydrated in 1× PBS for 10 min. To quench endogenous peroxidase activity, sections were incubated with peroxidase reagent (3% H2O2 in 1× PBS) for 15 min and gently washed twice in 1× PBS for 5 min. Slides incubated with a primary antibody cocktail overnight at 2°C to 8°C. The cocktail contained both monoclonal mouse anti-human CD3, (Clone F7.2.38; 1:100, Dako) and anti-p16 ARC antibody (EP1551Y; 1:100, Abcam, ab51243). Upon finishing and rinsing, the secondary antibody cocktail [peroxidase anti-rabbit IgG (H+L) (Vector Laboratories, PI-1000) and alkaline phosphatase anti-mouse IgG (H+L) (Vector Laboratories, AP-2000)] was prepared and applied as recommended by manufacturer (Vector Laboratories) followed by the subsequent visualization of AP activity (Vector Red Alkaline Phosphatase Substrate Kit)
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(Vector Laboratories, SK-5100) and HRP activity (Vector DAB Peroxidase Substrate Kit) (Vector Laboratories, SK-4100). Stained tissues were mounted with hematoxylin (SigmaAldrich), dehydrated, covered and imaged via microscope. Statistical analysis Statistics were calculated using GraphPad Prism software (GraphPad Software). If not stated differently, a two-sided t-test was used to determine significance and p30 nuclei for each donor. Fluorescence intensity in arbitrary units (a.u.). Results are mean ± SEM. (C) Distribution of fluorescence intensity (a.u.) strata from 5 RA (red) and 5 control samples (black) quantified in >30 nuclei for each donor. (D) Telomeric ends were analyzed for
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abnormal structures. Examples of double signal, apposition, fusion, and signal-free ends are shown. (E) Distributions of telomeric phenotypes in 10 patients and 10 controls. 150–200 nuclei were examined in each sample. Percentages of each damage pattern are presented. (F) Naïve and memory CD4+ T cells were separated and placed under proliferative stress by repetitive polyclonal stimulation. Loss of telomeric sequences was measured by PCR and, in parallel, telomeric damage foci were analyzed by dual-color immunostaining with antibodies to the DNA damage protein 53BP1 and the telomeric shelterin TRF2. 53BP1/TRF2 colocalization coefficients and telomeric length shortening in individual samples are correlated. *P < 0.05, two-tailed Student’s t-test. See also Figure S1 and Figure S2.
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Figure 2. The aging profile of CD4+CD45RA+ T cells in RA
CD4+CD45RA+ T cells from RA patients and age-matched controls were stimulated for 72 hrs. All data are mean ± SEM. (A, B) CDKN2A, CDKN1A, and TP53 transcript and p16 protein levels were measured in 7 RA-control pairs by RT-PCR and flow cytometry, respectively. (C) Expression of the aging marker CD57 assessed by flow cytometry. (D) SADS foci analyzed by confocal microscopy. Nuclei were hybridized with a sat-II–specific probe (red) and satellite DNA signals were examined in a minimum of 50 nuclei in each sample. Examples of condensed and threadlike distended satellites are shown in inserts. (E)
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Individual nuclei were scored as SADS-positive, if the pericentromeric or centromeric satellite heterochromatin had lost its tight packaging and was extended longitudinally. Data are from 3 RA-control pairs. (F) Telomeres were uncapped by transfecting healthy CD4+CD45RA+ T cells with TERF2 siRNA oligonucleotides. CDKN2A and CDKN1A transcripts were measured by RT-PCR. Results are from 3 independent experiments. (G) Flow cytometry for p16 expression. Representative histograms from control and TERF2 siRNA transfected cells are shown. Fluorescence Minus One control (FMO) is superimposed as grey area. Mean fluorescence intensities (MFI) of p16 are from 3 independent experiments. (H) CD57 expression was assessed by flow cytometry in 3 independent experiments. Representative data from control and TERF2 siRNA transfected cells are shown.*P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s t-test. See also Figure S3.
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Figure 3. MRE11Alow T cells in RA patients
MRE11A protein expression was quantified by intracellular staining of PBMCs in a cohort of RA patients (RA) and age-matched controls (Con) with antibodies against MRE11A and lineage markers (CD4, CD45RA). (A, B) Flow cytometric measurement of MRE11A protein expression in relation to donor age for naïve CD4+CD45RA+ T cells and memory CD4+CD45RO+ T cells. (C) Representative histograms from naïve CD4+CD45RA+ T cells from a healthy individual, an untreated RA patient and a RA patient on therapy. (D) Mean ± SEM of MRE11A MFI are from 5 samples per group. (E, F) Localization of MRE11A to
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the telomere quantified by dual-color immunostaining with anti-MRE11A (green) and antiTRF2 (red) in resting naïve CD4+CD45RA+ T cells of 5 RA patients and 5 age-matched controls. DNA is marked with DAPI (blue). (E) Representative image of immunostaining for MRE11A. (F) Staining intensities for total nuclear MRE11A and MRE11A-TRF2 colocalization measured in >50 nuclei from each of 5 different donors. *P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s t-test. See also Figure S4.
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Figure 4. Genetic or pharmacologic inhibition of MRE11A induces T cell aging
(A) The impact of MRN insufficiency on telomeric stability was tested by transfecting CD4+CD45RA+ T cells from healthy individuals with control, MRE11A, NBN, or RAD50 siRNA oligonucleotides. 48 hrs later, cells were stained with anti-53BP1 and anti-TRF2. Representative images of 53BP1 (green) and TRF2 (red) after knockdown of individual DNA repair proteins. DNA is marked with DAPI (blue). Merged images show colocalization of 53BP1 and TRF2. (B) Staining intensities (a.u.) for total 53BP1 and 53BP1/TRF2 colocalization were measured in individual nuclei. Mean ± SEM values are indicated.
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Results are from 5 independent knockdown experiments. (C) T cells were treated with the MRE11A inhibitor Mirin. Damage foci were analyzed by staining for 53BP1 and for 53BP1/ TRF2 colocalization. Data are from 5 independent experiments. All data are mean ± SEM. (D, E) CDKN2A and TP53 transcripts were measured by RT-PCR after transfecting CD4+CD45RA+ T cells from healthy individuals with control, MRE11A, NBN, or RAD50 siRNA oligonucleotides for 48 hrs (D) or were treated with the MRE11A inhibitor Mirin (E). (F) Flow cytometry for CD57 expression. Representative data are from control and MRE11A siRNA transfected cells. Percentages of CD57 expressing cells from 3 independent experiments are presented as mean ± SEM. (G) T cells were treated with vehicle or the MRE11A inhibitor Mirin at the indicated doses and analyzed for CD57 expression by flow cytometry. Mean ± SEM from 3 independent experiments. (H, I) Sat-II DNA (red) hybridization in control and Mirin-treated T cells. Distention of satellite DNA in the nuclei of Mirin-treated T cells. Nuclei were scored for SADS as in Figure 2 in >40 nuclei per sample and quantified in 3 control and Mirin-treated samples. *P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s t-test. See also Figure S5.
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Author Manuscript Author Manuscript Author Manuscript Figure 5. Overexpression of MRE11A repairs telomeric damage and prevents T cell aging in RA
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Naïve CD4+CD45RA+ T cells from RA patients were transfected with control plasmids or a myc-MRE11A construct. After 48 hrs, transfection efficiency was monitored by qPCR (A) and Western blotting (B). The error bars in (A) represent the 95% confidence internal. (C) Representative images of 53BP1 (green) and TRF2 (red) after overexpression of control plasmids or myc-MRE11A. DNA is marked with DAPI (blue). Merged images show colocalization of 53BP1 and TRF2. (D) Staining intensities (a.u.) for 53BP1/TRF2 colocalization were measured in a minimum of 70 individual nuclei from 3 different patients. Mean ± SEM values are indicated. CDKN2A and TP53 transcript levels (E) and p16 protein levels (F) were measured in control (RA+Vec) and myc-MRE11A (RA +MRE11A) transfected cells from 3 different patients. Results are mean ± SEM. (G) Flow
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cytometry for p16 expression. Representative data are from cells with or without MRE11A overexpression. (H) Percentages of p16 expressing cells analyzed in 4 independent experiments are presented as mean ± SEM. *P < 0.05, **P < 0.01 and ***P < 0.001, twotailed Student’s t-test.
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Author Manuscript Author Manuscript Author Manuscript Figure 6. The nuclease MRE11A controls pro-arthritogenic effector functions
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Synovial inflammation was quantified by gene transcriptional analysis in tissue extracts or by immunohistochemical analysis of tissue sections. (A) TRB transcript levels measured by qPCR to assess T cell accumulation. (B) Representative sections of synovial tissues stained with anti-CD3 antibodies (brown). (C) Percentages of CD3+ T cells in randomly selected fields of synovial tissues sections presented as mean ± SEM. (D) TNFSF11, (E) TNF, IL6, IL1B and (F) TGFB1, IL10 mRNA expression measured by qPCR in tissue extracts. (G) Transcription analysis of the aging markers CDKN2A, CDKN1A, and TP53 by qPCR. (H) p16 (brown) and CD3 (red) were stained by dual-color immunohistochemistry in synovial tissue sections from vehicle and MRE11A inhibitor-treated chimeras. (I) Percentages of p16+CD3+ T cells in randomly selected fields of synovial tissue sections presented as mean ± SEM. (J) T cell migratory capacity was measured in Transwell migration assays in the absence of chemokine gradients. N=6 RA patient-control pairs; n=4 young (65 years) healthy individuals; n=4 healthy control T cells with or without Mirin treatment. All data are mean ± SEM.*P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s ttest.
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Figure 7. Restoring MRE11A expression in RA T cells prevents pro-arthritogenic effector function
Pairs of NSG mice were engrafted with human synovial tissue and assigned to two treatment arms. CD45ROneg PBMC (A–D) and CD45RAnegPBMC (E–I) were prepared from RA patients and were transfected with either control plasmid or plasmid expressing MRE11A, and adoptively transferred into the chimeric mice. The intensity of synovial inflammation was compared by tissue gene expression analysis applying qPCR. (A, E and G) The density of the synovial T cell infiltrate was captured by TRB and TNFSF11 transcript levels. (B, C
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and H) Synovial cytokine production capability was assessed through TNF, IL6, IL1B, TGFB1 and IL10 transcript expression. (D, I) The impact of MRE11A overexpression on the tissue presence of aging markers was examined through CDKN2A and TP53 transcript levels. (F) Representative sections of synovial tissues stained with anti-CD3 antibodies (brown). Original magnification, X600 (insets in F). All data are mean ± SEM from at least 6 different synovial grafts.*P < 0.05, **P < 0.01 and ***P < 0.001, two-tailed Student’s ttest. See also Figure S6.
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