TRF2 inhibition rather than telomerase disruption drives CD4T cell dysfunction during chronic viral infection

T-cell activation and proliferation are essential for the adaptive immune responses against pathogenic infections and prophylaxis vaccinations (Laidlaw et al., 2016). However, T-cell overactivation and/or proliferation induce DNA damage and telomere erosion, which are considered to be the driving force of T-cell senescence and immune dysfunction – often seen in the setting of chronic inflammatory and infectious diseases (Gustafson et al., 2020). Indeed, we have recently shown that chronic viral (hepatitis C virus, HCV; hepatitis B virus, HBV; or human immunodeficiency virus, HIV) infection causes irreparable telomeric DNA damage, resulting in premature CD4⁺ T (CD4T)-cell aging and apoptotic death (Cao et al., 2021; Dang et al., 2020; Ji et al., 2019; Khanal et al., 2020; Nguyen et al., 2021a,b, 2018; Schank et al., 2020; Zhao et al., 2018, 2019).

Unlike many human somatic cells that do not express telomerase – an enzyme that elongates the telomeric DNA to prevent telomere shortening and protect chromosome ends (Nguyen et al., 2019) – T cells express telomerase during their activation and proliferation (Weng, 2008). Human telomerase consists of two components: the rate-limiting human (h) telomerase reverse transcriptase (hTERT) and the human telomerase RNA component (hTERC), which, along with several shelterin proteins, form a ribonucleoprotein complex to maintain the integrity of telomeres (Nguyen et al., 2019). T-cell-lineage proliferation is thought to be responsible for telomere shortening, as evidenced by the fact of the memory T-cell subset harbors shorter telomeres compared to their naïve counterparts (Rufer et al., 1999; Weng et al., 1995; Roth et al., 2003). Notably, long-term culturing of human T cells in vitro has revealed shortened telomeres and reduced levels of hTERT mRNA and telomerase activity (Rufer et al., 1999; Weng et al., 1995; Roth et al., 2003). In addition, telomere shortening is associated with T-cell senescence and reduced lineage proliferation (Najarro et al., 2015; Patrick et al., 2019). Moreover, activated human T cells exhibit high levels of DNA-damage-induced cell apoptosis (McNallya et al., 2017; Patrick and Weng, 2019). These studies indicate a complex dynamic between telomere or telomerase and T-cell function and homeostasis, which warrant further investigations.

Although telomere length is maintained under most conditions by telomerase, telomere-associated shelterin proteins are also essential for protecting telomeres from an unwanted DNA damage response (DDR) (Nguyen et al., 2019; Weng, 2008). In particular, telomeric repeat-binding factor 2 (TRF2 or TERF2) is a crucial component of the shelterin complex, which protects the integrity of telomeres by forming a telomere loop (t-loop) structure that prevents activation of the DNA repair pathways (Fujita et al., 2010; Okamoto et al., 2013; Vincent Picco et al., 2016; Ye et al., 2010). We have previously shown that inhibition of TRF2 in CD4T cells derived from HCV patients (Nguyen et al., 2021b, 2018) and CD4T cells derived from healthy subjects exposed to the telomere-targeting drug KML001 (Cao et al., 2019; Schank et al., 2020) resulted in accumulated telomeric DNA damage, telomere erosion and T-cell dysfunction. A previous study showed that telomerase recruitment to the telomeres is regulated by TRF2 via interactions with TIN2 and TPP1 (also known as TINF2 and ACD, respectively) (Nandakumar and Cech, 2013). However, whether TRF2 inhibition disrupts telomere integrity and T-cell functions by interrupting the access and/or recruitment of telomerase to telomeres in CD4T cells during chronic viral infection remains unknown.

In this study, we investigated the role of telomerase and TRF2 in telomeric DNA damage and CD4T cell dysfunction in the setting of chronic viral (HCV or HIV) infections. We demonstrated that the expression and activity of telomerase in HCV- or HIV-derived CD4T cells were intact, but TRF2 expression was significantly inhibited at the post-transcriptional level. Silencing TRF2 expression in CD4T cells derived induced telomeric DNA damage and T-cell dysfunction without affecting telomerase activity or telomerase translocation, similar to what we observed in CD4T cells during chronic HCV and HIV infection. These results indicate that premature T-cell aging and dysfunction during chronic viral infection are primarily caused by the inhibition of TRF2 rather than telomerase disruption, leading to DNA damage and T-cell dysfunction. This study suggests that reversing TRF2 protein inhibition could provide a novel approach to prevent telomeric DNA damage, thus rejuvenating T-cell functions during chronic viral infection.

We have previously shown that chronic HCV or HIV infection causes irreparable telomeric DNA damage and telomere erosion, leading to T-cell senescence and dysfunction (Cao et al., 2021; Dang et al., 2020; Ji et al., 2019; Khanal et al., 2020; Nguyen et al., 2021a,b, 2018; Schank et al., 2020; Zhao et al., 2018, 2019). To investigate whether telomere erosion in CD4T cells from HCV patients or people living with HIV (PLWH) is due to a telomerase deficiency, we examined the expression of hTERT at the transcriptional level by quantitative real-time reverse transcriptase PCR (RT-qPCR) in CD4T cells derived from HCV patients, PLWH and healthy subjects (HS). As shown in Fig. 1A, in the absence of T-cell receptor (TCR) stimulation, the levels of hTERT mRNA were almost similar in CD4T cells derived from HCV patients or PLWH compared to those in CD4T cells from HS.

Resting peripheral T cells express relatively low levels of hTERT protein, and TCR stimulation increases hTERT expression and telomerase activity (Patrick et al., 2019). Thus, we further analyzed hTERT protein levels in TCR-stimulated CD4T cells from HS. Following TCR stimulation, the hTERT protein levels increased on day 1, peaked on day 3 and then declined on day 4 (Fig. 1B). However, there were no significant differences in hTERT protein levels in TCR-stimulated CD4T cells from HCV patients or PLWH compared to those in TCR-stimulated CD4T cells from HS (Fig. 1C). We then measured telomerase activity using the telomeric repeat amplification protocol (TRAP) (Herbert et al., 2006) and found that, although telomerase activity was significantly increased in CD4T cells from all subjects after in vitro TCR stimulation for 3 days, there were no significant differences in the telomerase activity in CD4T cells (with or without TCR stimulation) from HCV patients or PLWH compared to that in CD4T cells from HS (Fig. 1D). Taken together, these results suggest that hTERT protein expression and activity are almost similar in CD4T cells from virally infected subjects (HCV patients or PLWH) and HS, regardless of the presence or absence of TCR stimulation.

The hTERT enzyme can be detected in both cytoplasmic and nuclear compartments in human T lymphocytes (Patrick et al., 2019). To further investigate the telomerase activity during T-cell activation, we isolated the cytoplasmic and nuclear proteins from CD4T cells derived from HS following TCR stimulation and determined the levels of hTERT by immunoblotting. As shown in Fig. 2A, the hTERT protein was detected only in the cytoplasm in resting CD4T cells. Upon TCR stimulation, hTERT was shuttled from the cytoplasm into the nucleus. Notably, DNA damage occurred along with TCR stimulation, as demonstrated by the increase in γH2AX levels (a marker for DNA damage) in both the cytoplasm and the nucleus. To determine whether telomeric DNA was damaged during TCR activation, we stimulated healthy CD4T cells with anti-CD3/CD28 antibodies for 24 h, followed by examining γH2AX levels at the telomeres using Amnis ImageStream flow imaging. As shown in Fig. 2B, without TCR stimulation, γH2AX signals were not detected at telomeres. Upon TCR stimulation, we observed γH2AX signals (red fluorescence) at the telomeres (green fluorescence) in the nucleus (DAPI staining). These data suggest that CD4T-cell activation promotes telomerase translocation into the nucleus, which corresponds to the induction of telomeric DNA damage.

To determine whether telomere erosion during chronic HCV or HIV infection is due to defective telomerase activity, we examined hTERT protein in the cytoplasmic and nuclear compartments in 3-day TCR-stimulated CD4T cells from HCV patients or PLWH and from HS using immunoblotting. As shown in Fig. 2C,D, there were no significant differences in the levels of hTERT protein in the cytoplasm and in the nucleus in activated CD4T cells derived from HCV patients or PLWH compared to those in activated CD4T cells derived from HS. These results indicate that hTERT protein levels in the cytoplasm and nucleus are almost similar in TCR-stimulated CD4T cells from virally infected subjects (HCV patients or PLWH) and from HS.

We further investigated the translocation of telomerase to telomeres in activated CD4T cells from HCV patients or PLWH and from HS using Amnis ImageStream flow imaging. Specifically, the CD4T cells were cultured with or without TCR stimulation for 3 days and then subjected to flow imaging analysis. Fig. 3 shows the gating strategy, representative images of CD4T cells and summary data for the mean bright detail similarity of colocalization of the hTERT signal (red) with the telomere probe (TelC, green). These results showed significant increases in telomere and hTERT colocalization signals in the TCR-stimulated cells compared to those in non-stimulated cells. However, there were no significant differences in the colocalization signals between TCR-stimulated CD4T cells from HCV patients or PLWH and those from HS. Taken together, these results indicate that telomerase is recruited to telomeres at a remarkably high level following CD4T cell activation, but the rate of telomerase translocation to telomeres is similar in activated CD4T cells from HCV patients or PLWH and HS.

Telomere shortening occurs during chronic HCV or HIV infection despite intact telomerase activity (Nguyen et al., 2021b, 2018; Zhao et al., 2018), indicating that mechanisms other than changes in telomerase activity are involved in telomere erosion and T-cell dysfunction in patients with these infections. Notably, the integrity of telomeric DNA is maintained by a telomere–shelterin complex, which is composed of six telomere-bound proteins: telomeric repeat-binding factors 1 and 2 (TRF1/TERF1 and TRF2, respectively), TRF1-interacting nuclear protein 2 (TIN2), repressor/activator protein 1 (RAP1 or TERF2IP), telomere protection protein 1 (TPP1) and protection of telomeres protein 1 (POT1). The primary function of the telomere shelterin proteins is to safeguard telomeric DNA from unwanted DDR (Nguyen et al., 2019; Weng, 2008). We have previously shown that TRF2 is the only shelterin protein that is downregulated in CD4T cells during chronic HCV infection (Nguyen et al., 2018). To assess the role of the shelterin proteins in telomere erosion and T-cell dysfunction during HIV infection, we examined shelterin protein levels in CD4T cells from PLWH and those from age-matched HS following cell culture for 4 days using immunoblotting. Similar to CD4T cells derived from HCV patients, we found that TRF2 was the only shelterin protein that was significantly downregulated in CD4T cells derived from PLWH, whereas the expression of all other components of the shelterin complex (including TRF1, POT1, TIN2, TPP1 and RAP1) remained unchanged compared to the expression of these proteins in CD4T cells from HS (Fig. 4A). In addition, we examined total TRF2 protein levels in naïve (CD45RA⁺) and memory (CD45RO⁺) CD4T cells from PLWH and HS by flow cytometry. The results showed that TRF2 was significantly downregulated in both cell subsets from PLWH compared to HS (Fig. 4B). Also, similar to what we previously found in HCV patients (Nguyen et al., 2018), there were no significant differences in the mRNA levels of TRF2 or other members of the shelterin complex between CD4T cells from PLWH and those from HS (Fig. 4C). Moreover, we examined TRF2 protein levels in the cytoplasmic and nuclear compartments in 3-day TCR-stimulated CD4T cells from HCV patients and PLWH versus those from HS and found that TRF2 was only present in the nucleus and that its levels were significantly lower in CD4T cells from HCV patients (Fig. 2C) and PLWH (Fig. 2D) compared to those in CD4T cells from HS. In essence, these results indicate that TRF2 expression is downregulated at the post-transcriptional level in CD4T cells during chronic HIV infection – similar to our previous observations in CD4T cells during chronic HCV infection.

The t-loop structure formed by the shelterin complex can negatively affect the binding of telomerase to telomeres, preventing its telomere-elongation activity (Smogorzewska et al., 2000). Thus, we hypothesized that the accessibility of telomerase to telomeres might be altered in TRF2-deficient cells in the setting of T-cell activation. To test this possibility, we investigated the translocation and activity of telomerase after TRF2 knockdown (KD) in CD4T cells from HS. The efficiency of TRF2 knockdown is shown in Fig. 5A. We did not observe any significant change in the levels of hTERT and phospho-S6 ribosomal protein (P-S6, a T-cell activation marker) in TRF2 KD cells compared to the control scrambled shRNA-transduced cells. Although TRF2 KD increased the levels of γH2AX compared to those in control cells, it did not affect telomere length or telomerase activity (Fig. 5B). However, the expression of the cellular proliferation marker Ki67 significantly decreased in TRF2-KD cells (Fig. 5C). Correspondingly, the frequency of apoptotic cells was higher in TRF2-KD cells (Fig. 5D). The elevated levels of apoptosis of TRF2-KD cells were further demonstrated by a significant increase in the expression of the apoptosis markers p53 and cleaved caspase-3, compared to the scrambled shRNA-transduced cells (Fig. 5E). Furthermore, TRF2 KD resulted in a significant decrease in the production of the cytokines IL-2 and IFN-γ compared to control cells (Fig. 5F). However, there was no significant difference in the telomeric translocation of telomerase in CD4T cells following TRF2 KD (Fig. 5G). Taken together, these results show that TRF2 inhibition induces telomeric DNA damage and T-cell apoptosis and dysfunction, and indicate that telomerase levels and activity are not affected by TRF2 KD in CD4T cells from HS. These results recapitulate what we observed in CD4T cells from patients with chronic HCV or HIV infections.

Cellular senescence in T lymphocytes is a state of extensive, replication-induced, accelerated telomeric DNA damage and telomere shortening, and is considered to be the driving force inducing aging-associated immune dysfunction in the elderly (Goronzy and Weyand, 2019), autoimmune disorders (Fujii et al., 2009; Wagner et al., 2018; Lee et al., 2016) and chronic viral infections (Nguyen et al., 2018; Zhao et al., 2018). Despite the importance of T-cell proliferation for the adaptive immune response, immune-activation-induced T-cell proliferation has been shown to negatively regulate telomerase expression and telomere length, resulting in T-cell senescence. This further leads to decreased T-cell lineage proliferation and increased cellular apoptosis (Patrick and Weng, 2019; Patrick et al., 2019). Indeed, telomere length has been reported to be shortened in older compared to younger individuals (Tedone et al., 2019) and in differentiated memory T cells compared to naïve T cells (Patrick et al., 2019). Recently, the dynamics of T-cell telomere length have been applied to predict disease severity in coronavirus disease 2019 (COVID-19) patients; in this study, patients with short telomeres have been reported to have higher risks of severe disease outcomes and death compared to those with longer telomeres (Froidure et al., 2020; Aviv, 2020). These studies suggest that disrupted telomere integrity and shortened telomeres in T lymphocytes are the hallmarks of immune aging and play a vital role in predicting disease severity across a wide range of viral infections.

Here, we showed that telomerase expression and activity were quite similar in CD4T cells from HCV patients or PLWH when compared to those in CD4T cells from HS. However, CD4T cells from HCV patients or PLWH still showed telomere erosion, as demonstrated by an increased level of telomeric DNA damage compared to that in HS. Our previous studies have shown that, upon TCR stimulation, CD4T cells exhibit increased cellular exhaustion, senescence and expression of apoptosis markers (Cao et al., 2021; Dang et al., 2020; Ji et al., 2019; Khanal et al., 2020; Nguyen et al., 2021a,b, 2018; Schank et al., 2020; Zhao et al., 2018, 2019). In addition, we demonstrated that CD4T cells derived from HCV patients or PLWH exhibit a similar phenomenon of premature aging due to T-cell overactivation and proliferation. These changes in activated CD4T cells were also accompanied by accelerated telomere erosion, as evidenced by the shortening of telomere length as well as accumulated telomeric DNA damage.

In the present study, we further investigated the expression of hTERT as well as telomeric DNA damage in both cytoplasmic and nuclear compartments in activated CD4T cells from HCV patients, PLWH and HS. Our data showed that, upon TCR stimulation, telomerase was translocated from the cytoplasm to the nucleus, and cells exhibited increased levels of the DNA damage marker γH2AX (Fig. 2A). We obtained further evidence of the telomeric DNA damage (Fig. 2B) as increased colocalization of telomeres and γH2AX was observed in the nucleus of activated cells compared to non-activated control cells. However, the telomerase expression, activity and translocation into the nucleus did not differ between activated CD4T cells derived from HCV patients or PLWH and those from HS (Fig. 2C,D and Fig. 3).

TRF2 is a crucial component of the shelterin complex that protects the ends of telomeres by forming a t-loop structure (Diotti and Loayza, 2011; Lange, 2005). The expression of TRF2 is downregulated in CD4T cells derived from patients chronically infected with HCV or HIV. The deficiency of TRF2 is associated with the accumulation of telomeric DNA damage and telomere erosion (Nguyen et al., 2021b, 2018). Similar to our previous findings showing decreased TRF2 expression in CD4T cells from HCV patients, we found a significant decrease in TRF2 protein levels in CD4T cells from PLWH (Fig. 4). The TRF2 KD in CD4T cells from HS provided further evidence of the impact of TRF2 deficiency on CD4T cell function as well as on the translocation of telomerase into the nucleus, but TRF2 KD did not affect telomerase levels. Specifically, our data showed that TRF2 KD resulted in increased telomeric DNA damage and apoptosis with decreased cytokine production and cellular proliferation; however, telomerase expression, activity and nuclear translocation remained unchanged compared to the control (Fig. 5). These results regarding TRF2 KD in CD4T cells from HS recapitulate what we observed in patients with HCV or HIV infection.

Based on these findings, we conclude that CD4T cell dysfunction during chronic HCV or HIV infection is not a function of perturbations in telomerase expression, activity or translocation, but rather a function of the inhibition of the shelterin complex protein TRF2. A model depicting this process is shown in Fig. 6. We propose that chronic viral (HCV or HIV) infection leads to an inhibition of TFR2 with subsequent telomere deprotection, thus exposing the telomeres to DNA damage and driving cell apoptosis, leading to impaired cytokine expression and cellular dysfunction. At the core of our studies is the role of TRF2 in overall telomere homeostasis. We show that chronic HCV or HIV infection can stimulate T-cell overactivation and/or proliferation, which cause TRF2 inhibition, telomeric DNA damage and T-cell exhaustion or senescence, leading to T-cell dysfunction and a decreased ability to proliferate. In turn, decreased TRF2 further increases telomeric DNA damage and decreases T-cell proliferation. Thus, this malicious cause–effect relationship eventually impairs T-cell function and leads to viral persistence. Support for this paradigm and the role of activation-induced TFR2 inhibition can be found in the studies of HIV-1-specific cytotoxic T-cell responses during progressive HIV infection, wherein Lichterfeld et al. (2012) demonstrated a selective reduction of TFR2 (and TPP1) in these cells and linked this reduction to accelerated aging during viral infection. In addition, recent studies of telomeric dysfunction in Epstein–Barr virus (EBV)-associated Hodgkin's lymphoma by Knecht and Mai (2017) demonstrated a progressive disruption of shelterin activity by the downregulation of TRF2, leading to telomeric dysfunction, which culminated in the progression of chromosomal rearrangements that underlie the pathogenesis of classic Hodgkin's lymphoma (Knecht and Mai, 2017). Notably, TRF2 is also the key shelterin protein affected in EBV-infected lymphoblastic cells and is considered to be involved in the mechanism for telomere deprotection (Kamranvar et al., 2013). Taken together, these studies suggest that TRF2 inhibition is a shared mechanism underlying both telomeric DNA damage and T-cell dysfunction during chronic viral infection.

The molecular mechanisms that induce TRF2 downregulation in CD4T cells in patients with chronic viral infection are still not completely understood. Our previous studies showed that TRF2 is inhibited in naïve CD4T cells in HCV patients due to increased proteasomal degradation that is associated with the activation of the p53-SIAH1 ubiquitination pathway (Nguyen et al., 2018). In addition, we have shown that persistent immune activation, as demonstrated by the hyperactivation of phosphoinositide 3-kinase/Akt/mammalian target of rapamycin (mTOR) signaling, is the cause of TRF2 inhibition and the bias toward pro-inflammatory differentiation of total CD4T cells in HCV patients (Nguyen et al., 2021b). Furthermore, several studies have investigated the mechanisms that regulate TRF2 inhibition using cell lines and suggested that TRF2 expression is tightly regulated by post-translational mechanisms via the regulation of the chromatin state, such as by SIRT6-mediated deacetylation in colorectal cancer cells (Rizzo et al., 2017) or by long non-coding RNAs and microRNAs in fibroblast and breast cancer cells, respectively (Hu et al., 2018; Luo et al., 2015). However, the mechanisms that induce TRF2 inhibition in CD4T cells during chronic viral infection warrant further investigation.

Based on our results and published reports, it appears that telomere integrity or homeostasis is essential to the maintenance of normal cellular function and the aging process. During chronic viral infection, TRF2 functions as a key mediator or biomarker for telomere homeostasis and, thus, telomere attrition or erosion can lead to significant telomeric DNA damage and cellular dysfunction. Therefore, restoring TRF2 protein inhibition might provide a novel approach to rescue telomere loss and prevent DNA damage and premature T-cell aging, thus rejuvenating CD4T cell functions in chronic viral diseases.

This study was approved by the joint institutional review board of East Tennessee State University, TN and James H. Quillen Department of Veteran Affairs Medical Center, TN. Written informed consent was obtained from all participants, and all investigations involving human subjects were conducted according to the principles expressed in the Declaration of Helsinki. All HCV patients were positive for HCV RNA for at least 6 months prior to antiviral treatment. All PLWH were on antiretroviral therapy for at least 1 year with an undetectable viral load (HIV RNA). Blood samples from HS were obtained from BioIVT (Biological Specialty Company, Gray, TN) and were negative for HBV, HCV and HIV infections. The demographic characteristics of all subjects included in this study are shown in Table S1.

Peripheral blood mononuclear cells (PBMCs) were isolated from fresh heparinized whole blood by Ficoll-Paque density centrifugation (Cytiva, Marlborough, MA). Total CD4T cells were isolated from PBMCs using the CD4T Cell Isolation Kit (Miltenyi Biotec, Cambridge, MA). The cells were cultured in RPMI-1640 complete medium (Fisher Scientific; Pittsburgh, PA) containing 10% fetal bovine serum (Atlanta Biologicals, Flowery Branch, GA), 100 IU/ml penicillin and 2 mM L-glutamine (Thermo Fisher Scientific, Waltham, MA) at 37°C and 5% CO₂. For the differentiation of naïve CD4T cells to effector CD4T cells, the cells were incubated with Dynabead-coated human anti-CD3/CD28 antibodies for T-cell activation and expansion (Thermo Fisher Scientific) at a one-bead-to-one-cell ratio for 3 days, unless indicated otherwise. For measurement of cytokine production, cells were stimulated with phorbol 12-myristate-13-acetate/ionomycin with Brefeldin A (BioLegend, San Diego, CA) for 4–6 h before the detection of IL-2 and IFN-γ.

Total RNA was extracted from 1×10⁶ cells using the RNeasy Mini kit (QIAGEN, Germantown, MD) and reverse transcribed to cDNA using the High-Capacity Reverse Transcription Kit (Applied Biosystems, Foster City, CA) according to the manufacturer's instructions. RT-qPCR was performed in triplicate using iTaq Universal SYBR Green Supermix (Bio-Rad, Hercules, CA) according to the manufacturer's protocol. Gene expression was calculated using the 2^−ΔΔCt method (Livak and Schmittgen, 2001), normalized to GAPDH levels, and is presented as fold change. The PCR primer sequences used are listed in Table S2.

Immunoblotting was performed as previously described (Zhao et al., 2018). The following primary antibodies were used: anti-telomerase catalytic subunit (600-401-252S, 1:500) from Rockland Immunochemicals, Limerick, PA (Eitan et al., 2012; Kabacik et al., 2018); anti-TRF2 (13136S, 1:1000), anti-phospho-S6^Ser235/236 ribosomal protein (5316S, 1:1000), anti-RAP1 (5433S, 1:1000), anti-TPP1 (14667S, 1:1000), anti-β-actin HRP-conjugated (4970S, 1:2000), anti-caspase-3 (9662S, 1:1000), anti-cleaved-caspase-3 (9661S, 1:1000) and anti-lamin A/C (2032S, 1:1000) from Cell Signaling Technology; anti-GAPDH (0411, 1:500) and anti-P53 (sc-126, 1:500) from Santa Cruz Biotechnology; anti-TRF1 (NBP141217, 0.5 µg/ml) and anti-TIN2 (NBP2-55709, 0.5 µg/ml) from Novus Biologicals; and anti-POT1 (MAB5299, 2 µg/ml) from R&D systems. The following horseradish peroxide-conjugated secondary antibodies were used at a dilution of 1:3000: anti-rabbit (7074S) and anti-mouse (7076S) from Cell Signaling Technology. Protein bands were visualized with the Amersham ECL Prime Western Blotting Detection Reagent (GE Healthcare Bio-Sciences) and captured and quantitatively analyzed by Chemi DocTM MP Imaging System (Bio-Rad).

Cytoplasmic and nuclear extracts were isolated following a previously published protocol (Gagnon et al., 2014) and the cell pellet was resuspended in cell extraction (CE) buffer (10 mM Tris-HCl, pH 7.4; 2 mM Na₃VO₄; 100 mM NaCl; 1% Triton X-100; 1 mM EDTA; 10% glycerol; 1 mM EGTA; 0.1% SDS; 1 mM NaF; 0.5% deoxycholate; 20 mM Na₄P₂O₇) with protease inhibitor cocktail (Thermo Fisher Scientific, Waltham, MA). The mixture was kept on ice for 10 min and then centrifuged at 800 g for 8 min. The supernatant was collected as the cytoplasmic fraction. The pellet was washed with CE buffer and resuspended with RIPA buffer (BP-407; Boston BioProducts, Milford, MA) mixed with protease inhibitor cocktail. The mixture was then sonicated at 50 kHz for 2 s and then centrifuged at 18,000 g for 15 min at 4°C. The supernatant was collected as the nuclear fraction.

The activity of the telomerase enzyme was measured using telomeric repeat amplification protocol (TRAP assay) as previously described (Herbert et al., 2006). The primers used in this experiment are listed in Table S2.

Staining for cell surface markers, intracellular proteins and cytokines was performed as described previously (Nguyen et al., 2018; Zhao et al., 2018). The following fluorochrome-conjugated antibodies were used: anti-TRF-2 (NB100-56506PE, 0.1 µg/10⁶ cells; Novus Biologicals, Littleton, CO); anti-H2A.X Phospho (Ser139) (613412, 5 µl/10⁶ cells), anti-IL-2 (500310, 5 µl/10⁶ cells), anti-IFN-γ (502509, 5 µl/10⁶ cells) and anti-Ki-67 (350520, 5 µl/10⁶ cells) from BioLegend, San Diego, CA. The probe TelC-AF488 (F1004, 0.5 µg probe/ml; PNA Bio, Newbury Park, CA) was used for fluorescence in situ hybridization (FISH). For apoptosis analysis, cells were stained with Annexin V and 7-AAD from the PE Annexin V apoptosis detection kit (BD Biosciences, San Jose, CA). Cells were assayed with a BD Accuri C6 Plus Flow Cytometer (BD Biosciences) and the data were analyzed by FlowJo software (v10.0).

CD4T cells from HS were left unstimulated or stimulated with Dynabeads coated with anti-CD3/CD28 antibodies at a one-bead-to-one-cell ratio for 24 h. The cells were fixed with fixation buffer (BioLegend) for 15 min at room temperature, washed and resuspended in flow cytometry buffer [containing phosphate-buffered saline (PBS), pH 7.2, 0.5% bovine serum albumin (BSA) and 2 mM EDTA]. The fixed cells were permeabilized with Foxp3/transcription factor staining buffer (Thermo Fisher Scientific) for 45 min at room temperature, followed by washing once with flow cytometry buffer and then staining with anti-γH2AX antibody (1:150) in 1× permeabilization buffer (Thermo Fisher Scientific) at 37°C for 1 h. The cells were washed with 1× permeabilization buffer, fixed in fixation buffer for 15 min at room temperature and washed once with flow cytometry buffer. Next, the cells were resuspended in hybridization buffer (70% formamide, 1% bovine serum albumin, 150 mM NaCl and 20 mM Tris-HCl pH 7.2). The telomere probe TelC-AF488 was used to hybridize cellular telomeres at a concentration of 0.5 µg probe/ml. The cells were incubated at room temperature for 10 min in the dark and hybridized at 82°C for 12 min. After incubating overnight in the dark at room temperature, the cells were washed twice with post-hybridization buffer (70% formamide, 0.1% bovine serum albumin, 150 mM NaCl, 10 mM Tris-HCl pH 7.2 and 0.1% Tween-20) and once with flow cytometry buffer. Then, the cells were stained with goat anti-rabbit IgG-Alexa Fluor 647 antibody at 1:100 (Jackson ImmunoResearch Laboratories, West Grove, PA; 111-605-003) in 1× permeabilization buffer at 37°C for 1 h, followed by washing once with 1× permeabilization buffer and once with flow cytometry buffer. DAPI staining was performed in Dulbecco′s phosphate-buffered saline (DPBS) at room temperature for 15 min, followed by washing twice with flow cytometry buffer. The cells were resuspended in flow cytometry buffer and subjected to ImageStream flow cytometry using an Amnis ImageStream Mk II Imaging Flow Cytometer (Luminex, Austin, TX) with 60× magnification. Approximately 5000 cells per sample were acquired and the images were analyzed using IDEAS software version 6.2 (Luminex). The colocalization of fluorescent probes was measured using the Mean Similarity Detail Feature in IDEAS, which identifies the small punctate staining in image pairs and then measures how well those images correlate.

For lentivirus packaging, human embryonic kidney cells expressing SV40 large T antigen (HEK293T) at 80% confluency were transfected with 2.5 μg of pMD2.G plasmid (Addgene #12259), 7.5 μg of psPAX2 plasmid (Addgene #12260) and 10 μg of pLKO.1-puro Non-Mammalian shRNA control (Sigma-Aldrich, St Louis, MO, SHC002) or pLK0.1-puro-TERF2 shRNA (Sigma-Aldrich, SHCLNG-NM_005652, TRCN0000004811) for TRF2 knockdown using the Transporter 5 Transfection Reagent (Polysciences, Warrington, PA). The lentiviruses were harvested, filtered using a 0.45 µM sterile syringe filter (Sigma-Aldrich, SLHV004SL) and concentrated using PEG-it Virus Precipitation Solution (System Biosciences, Palo Alto, CA) following the manufacturer's protocol. The lentiviruses were resuspended in sterile DPBS and stored at −80°C. CD4T cells from HS were seeded at 2.5×10⁶ cells/ml in RPMI-1640 complete medium supplemented with anti-CD3/CD28 antibody-coated beads at a one-cell-to-0.5-beads ratio for 24 h. The cells were harvested and transduced with shRNA scramble or shRNA-TERF2 encoded viruses. TransDux MAX Lentivirus Transduction Reagent (System Biosciences) was used to enhance transduction efficiency. Briefly, the stimulated CD4T cells were resuspended in 400 mL RPMI-1640 complete medium containing 100 μl of MAX Enhancer, 2 μl of TransDux^MAX and 4 μl of 1 M HEPES buffer. The cell suspension was then seeded into a 24-well plate with the desired lentivirus amount. The plates were spun at 1500 g and 32°C for 1.5–2 h. The supernatants were discarded and the cells resuspended in 1 ml RPMI-1640 complete media containing anti-CD3 (1 µg/ml) and anti-CD28 antibodies (2 µg/ml). After 48 h, the transduced cells were collected and used for further analysis.

Data were obtained from at least three independent experiments and analyzed by GraphPad Prism version 8, and are presented as mean±s.e.m. Statistical outliers were removed by the ROUT method (Q=1%). Differences between two groups were analyzed by two-tailed unpaired independent Student's t-test, Mann–Whitney U-test, two-tailed paired t-test or Wilcoxon matched-pairs signed-rank test. Differences among multiple groups were analyzed by one-way ANOVA with Tukey's multiple comparisons post hoc test. P<0.05 was considered statistically significant, and P<0.01, P<0.001 and P<0.0001 were considered very significant.

This publication is the result of work supported with resources and the use of facilities at the James H. Quillen Veteran Affairs Medical Center.

The authors declare no competing or financial interests.