Long noncoding RNA HITT coordinates with RGS2 to inhibit PD-L1 translation in T cell immunity

HITT promotes T cell immunity. We first compared the anticancer effects of HITT in immune-competent BALB/c mice treated with anti-CD8α antibody to block CD8+ T cell cytotoxicity or the IgG control (Figure 1, A–C). As expected, murine mammary carcinoma 4T1 grew more quickly in mice treated with anti-CD8α antibody than in mice treated with IgG isotype control (Figure 1, A–C). HITT overexpression in 4T1 cells attenuated tumor growth under both conditions (Figure 1, A–C), but it suppressed tumor growth more evidently in the control mice (HITT/vector control: 25%–34%) than in anti-CD8α antibody–treated mice (HITT/vector control: 78%–80%) (Figure 1, A–C). This is not due to the different HITT fold changes (Figure 1D). In line with above data, MTT and BrdU incorporation assays revealed no obvious intrinsic impacts of HITT on cell viability and proliferation in 4T1 cells (Supplemental Figure 1, A and B; supplemental material available online with this article; https://doi.org/10.1172/JCI162951DS1). Because of this observation, the effects of HITT expression by cancer cells on T cell activity were further explored. MDA-231 (breast cancer) and HeLa (cervical cancer) cells stably expressing HITT and vector controls were successfully established and validated by quantitative reverse-transcription PCR (RT-PCR) (Supplemental Figure 1C). CD8+ T cells were isolated from human blood and activated as described previously (25) and then cocultured with the established cancer cell lines (Figure 1E). HITT overexpression by cancer cells elevated cytotoxic T lymphocyte (CTL) activity, as indicated by increased secretion of IL-2 and IFN-γ in the culture medium (Figure 1F). In agreement, HITT-overexpressing cells also exhibited increased vulnerability to CTL attack (Figure 1G). CRISPR/Cas–mediated HITT KO produced opposing results regarding both IL-2 and IFN-γ secretion and T cell–mediated cancer-killing effects (Figure 1, H and I, and Supplemental Figure 1D). Thus, HITT expression by cancer cells plays an important role in promoting T cell immunity.

HITT sensitizes cancer cells to T cell–mediated cytotoxicity.Figure 1

HITT sensitizes cancer cells to T cell–mediated cytotoxicity. (AC) Volume (A), images (B), and weight (C) of 4T1 syngeneic tumors. Vect., vector. (D) HITT levels in 4T1 syngeneic tumors determined by qRT-PCR. (E) Schematic showing crystal violet staining to analyze T cell–mediated tumor cell–killing efficacy. (F) Detection of IL-2 and IFN-γ levels in the supernatants of T cell control and HITT-overexpressing MDA-231 and HeLa cell cocultures by ELISA assays. (G) Detection of the attached MDA-231 and HeLa cells by crystal violet staining after coculture with the activated T cells for 6 hours. Intensities are shown in bar graphs (right). (H) Detection of IL-2 and IFN-γ levels in the supernatants of T cell and MDA-231 and HeLa cell cocultures by ELISA assays. (I) Detection of the attached MDA-231 and HeLa cells by crystal violet staining after coculture with the activated T cells for 6 hours. Intensities are shown in bar graphs (right). Data in A and C are shown as mean ± SD (n = 5). Data in C, D, and FI are derived from 3 independent experiments and are represented as mean ± SEM. *P < 0.05; **P < 0.01; ***P < 0.001; ***P < 0.0001; NS, not significant by 2-way ANOVA (A), 1-way ANOVA (C and FI), and Student’s t test (D).

HITT inhibits PD-L1 expression. To understand how HITT attenuates T cell immunity, we compared mass-spectrum data in the control and HITT knockdown (KD) HeLa cells. Unsupervised hierarchical-clustering analyses showed that the HITT-KO samples were clustered separately with the controls (Supplemental Figure 1E). A volcano plot demonstrates that 69 proteins were differentially regulated by HITT KO using a threshold of P ≤ 0.05 and fold change ≥ 1.8, with PD-L1 as one of the top hits (Supplemental Figure 1F). Therefore, the impacts of HITT on PD-L1 expression were explored. Remarkably, PD-L1 was dramatically reduced in HITT-overexpressing human breast cancer cells (MDA-231, MDA-468, and BT549), mouse mammary cancer cells (4T1), cervical cancer cells (HeLa), and colon cancer cells (HT29) (Figure 2A and Supplemental Figure 1G). In contrast, HITT KO or siRNA-mediated HITT KD led to increased PD-L1 expression (Figure 2B and Supplemental Figure 1G). Restoration of HITT expression abolished HITT KD–mediated PD-L1 elevation (Supplemental Figure 1H), while the expression of another family member, PD-L2, was unaffected (Figure 2, A and B). PD-L1 localization was not changed by HITT (Supplemental Figure 1I). Therefore, HITT mainly regulates PD-L1 by repressing its expression, but not by changing its localization.

IFN-γ–induced and E2F1-mediated transactivation of HITT attenuates PD-L1 exFigure 2

IFN-γ–induced and E2F1-mediated transactivation of HITT attenuates PD-L1 expression. (A and B) PD-L1 and PD-L2 protein levels analyzed by WB assay in HITT stable overexpression (A) or HITT-KO (B) cells. (C and D) HITT levels determined by qRT-PCR in MDA-231 and HeLa cells treated with different concentrations of IFN-γ for 24 hours (C) or treated for the indicated time periods with 10 ng/ml IFN-γ (D). (E) PD-L1 protein levels analyzed by WB in IFN-γ–treated cells with or without HITT KD. (F and G) HITT promoter luciferase activities determined by luciferase reporter assay in MDA-231 and HeLa cells treated with different concentrations of IFN-γ for 24 hours (F) or the indicated time periods with 10 ng/ml IFN-γ (G). (H) Relative binding potentials between different transcription factors and HITT promoter region were analyzed by UCSC ChIP sequence data. (I) E2F1 protein levels were detected by WB in MDA-231 and HeLa cells with different concentrations of IFN-γ for 24 hours or with 10 ng/ml IFN-γ for different time courses. (J) HITT expression levels and HITT promoter luciferase activities were measured by qRT-PCR and luciferase reporter assay in IFN-γ–treated (10 ng/ml for 24 hours) cells after E2F1 KD. E2F1 KD efficiency was validated by WB (bottom). (K) HITT promoter (full length and MT) controlled luciferase activities were determined after transient transfection of the indicated reporter plasmids together with E2F1 expression plasmid. (L) Binding between HITT promoter region and E2F1 was determined by ChIP assay after IFN-γ treatment (10 ng/ml for 24 hours). PCR band intensities were quantified using ImageJ and are presented in the bar graph (bottom). Data are derived from 3 independent experiments and are shown as mean ± SEM. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001; NS, not significant by 1-way ANOVA (C, D, F, G, and J) and Student’s t test (K and L).

Intriguingly, HITT expression was increased in a dose- and time-dependent manner in response to IFN-γ exposure in MDA-231 and HeLa cells (Figure 2, C and D). In addition, IFN-γ–induced HITT expression was relatively common because treatment led to increased HITT expression in all breast cancer cell lines tested regardless of their genetic features (Supplemental Figure 2A and Supplemental Table 1). IFN-γ–induced HITT expression was also observed in lung cancer cells, such as H23 and H1299 (Supplemental Figure 2A). These data suggest that HITT is a newly identified IFN-γ signal–responsive lncRNA. In addition, we observed that PD-L1 expression was increased by IFN-γ, whereas 2 independent siRNA-mediated HITT KDs augmented IFN-γ–induced PD-L1 expression (Figure 2E). Therefore, HITT plays important roles in attenuating PD-L1 expression under both basal and IFN-γ–stimulated conditions.

E2F1 transactivates HITT upon IFN-γ stimulation. Given the essential role of HITT in regulating PD-L1 expression, we further explored the underlying mechanisms of IFN-γ–induced HITT expression. HITT promoter luciferase reporter and luciferase-HITT reporter were generated (Supplemental Figure 2B). HITT promoter–driven luciferase activity was elevated in a dose- and time-dependent manner following IFN-γ treatment (Figure 2, F and G), while luciferase-HITT reporter activity was unchanged under the same conditions (Supplemental Figure 2C), suggesting that HITT is activated by IFN-γ at the transcriptional level. In line with these results, actinomycin D (ActD), an mRNA synthesis inhibitor, abolished IFN-γ–induced HITT expression (Supplemental Figure 2D).

We then analyzed the UCSC Genome Browser ChIP-sequencing database (Figure 2H). The most potent transcription factors were early growth response 1 (EGR1), TATA-box binding protein associated factor 1 (TAF1), and E2F transcription factor 1 (E2F1) (Figure 2H). IFN-γ treatment barely affected the expression of EGR1 (Supplemental Figure 2E). Despite detection of increased levels of TAF1 in a time-dependent manner after IFN-γ treatment, diminishing its expression by siRNA failed to influence HITT levels (Supplemental Figure 2F). In contrast, E2F1 was remarkably enhanced by IFN-γ in a dose- and time-dependent manner, accompanied by a coordinate increase of HITT expression (Figure 2I). Inhibition of E2F1 expression by 2 independent small interfering E2F1s (si-E2F1s) completely abolished IFN-γ–induced HITT expression and HITT promoter luciferase activity (Figure 2J).

In addition, ectopic E2F1 expression increased HITT levels and HITT promoter–driven luciferase activity in an E2F1 dose–dependent manner (Supplemental Figure 2G), while KD of endogenous E2F1 reduced them (Supplemental Figure 2H). Furthermore, the activity of mutant type 1 (MT1) luciferase reporter, which contains the predicted E2F1-binding sites, was as effective as that of WT reporter in response to E2F1 expression (Figure 2K), whereas MT2 luciferase reporter, without the predicted binding motif, largely lost its response to E2F1. Moreover, binding between E2F1 and the HITT promoter region was verified by a ChIP assay, and binding was increased after IFN-γ treatment (Figure 2L). E2F1 is therefore required for transcriptional activation of its target HITT upon IFN-γ stimulation.

HITT and RGS2 coordinately inhibit PD-L1 translation. Meanwhile, considering the essential role of PD-L1 in immune evasion, we investigated the mechanisms underlying HITT-inhibited PD-L1 expression. First, we found no obvious change in the expression of Cd274 mRNA, encoding for PD-L1, after HITT overexpression or KD (Supplemental Figure 3, A and B). Secondly, neither lysosome inhibitor chloroquine nor proteasome inhibitor MG132 influenced HITT-mediated PD-L1 inhibition (Supplemental Figure 3, C and D). Intriguingly, a click chemistry and l-azidohomoalanine–label (AHA-label) assay revealed that HITT overexpression inhibited newly synthesized PD-L1 protein (Figure 3A), while HITT KD promoted it (Figure 3B), with the newly synthesized HSP90 serving as a negative control (Figure 3, A and B).

HITT inhibits PD-L1 translation in an RGS2-dependent manner.Figure 3

HITT inhibits PD-L1 translation in an RGS2-dependent manner. (A and B) Affinity purification of biotinylated AHA-labeled acutely synthesized proteins of PD-L1, RGS2, and HSP90 was detected by WB after HITT overexpression with or without RGS2 KD (A) or RGS2 overexpression with or without HITT KD (B). (C) PD-L1 protein levels were analyzed by WB in HITT stable lines with or without RGS2 KD. (D and E) Polysome in the cytoplasm was fractionated through sucrose gradients. The total RNA amount was determined by the intensity at 254 nm (D), and PD-L1 and GAPDH mRNA levels were detected by qRT-PCR (E) in gradient fractions of HITT stable-expression HeLa cells with or without RGS2 KD. Representative data as a percentage of total RNA of interest in the gradient from 3 independent experiments are presented. *P < 0.05; **P < 0.01, Student’s t test (D and E).

It is reasonable to suppose that HITT may fulfil its roles by cooperating with translational regulators. To test this hypothesis, we first utilized the Gene Ontology (GO) database to search translational regulators in the genome. In total, 78 proteins were identified as negatively involved in protein translation. Among them, we identified 15 proteins that have been reported to be directly or indirectly related to T cell immunity via a literature search (Supplemental Table 2). We then used RNA interference techniques to specifically inhibit the expression of those individual genes (Supplemental Figure 4A). KD efficiency was verified in each case by qRT-PCR. Western blot (WB) assay revealed an obvious increase of PD-L1 protein expression in the (RGS2) KD cells, but not others (Figure 3C and Supplemental Figure 4A). Intriguingly, the ability of HITT to regulate PD-L1 expression was largely diminished by RGS2 KD (Figure 3C). RGS2 had little effect on PD-L1 expression on the mouse cell line 4T1, which does not contain HITT, and overexpression of HITT in 4T1 cells restored the effects of RGS2 KD on PD-L1 expression (Supplemental Figure 4B). Furthermore, the click chemistry and AHA-label assay showed that RGS2 KD increased the levels of the newly synthesised PD-L1 protein and also abolished HITT overexpression–inhibited PD-L1 expression (Figure 3A). In contrast, RGS2 overexpression repressed the newly synthesized PD-L1 protein and also rescued HITT KD–induced PD-L1 expression (Figure 3B). Coordinated regulation of PD-L1 translation by RGS2 and HITT was further validated by a chromosome fractionation assay (Figure 3, D and E). Namely, RGS2 and HITT similarly reduced polysome-occupied Cd274 mRNA and no further reduction was observed with their combination (Figure 3E). These data suggest that HITT and RGS2 coordinately regulate PD-L1 translation through the same mechanism.

1,080-1,130 nt HITT is physically associated with F194, Q196, and D197 in the RGS domain of RGS2. Given their coordinated effects on PD-L1 translation, we speculated that HITT may bind with RGS2. Indeed, a UV cross-linking and immunoprecipitation (CLIP) assay (Figure 4A) revealed that HITT and RGS2 physically associate with each other in living cells, and their association was increased after ectopic HITT overexpression (Figure 4B). Consistently, their binding was increased by IFN-γ, while inhibition of IFN-γ–induced HITT expression by si-HITT abolished such an effect (Figure 4C and Supplemental Figure 4C). Direct binding between HITT and RGS2 was also validated by RNA pull-down assay using in vitro–synthesised biotinylated HITT and purified RGS2 protein, and their binding was suppressed by antisense HITT (Figure 4, D and E).

RGS2 is a binding partner of HITT.Figure 4

RGS2 is a binding partner of HITT. (A) Schematic of CLIP assay for binding between RGS2 and HITT in living cells. (B and C) HITT levels determined by qRT-PCR following CLIP RGS2 after HITT overexpression (B) or KD in the presence or absence of IFN-γ treatment (C) in HeLa cells, with GAPDH or 18s mRNA and CLIP IgG as negative controls. (D) Schematic of in vitro RNA pull-down assay to analyze the binding between in vitro–synthesized biotin-labeled HITT and purified RGS2. (E) GST-tagged RGS2 protein coprecipitated with biotin-sense-HITT in the presence or absence of digoxin-antisense-HITT. (F) RGS2 protein coprecipitated by biotin-HITT-F3-1 (1,030–1,247 nt) or its fragments determined by RNA pull-down assay. Schematic showing sequentially fragmented HITT-F3-1 (1,030–1,247 nt). (G) GST-tagged full-length RGS2 or its mutants coprecipitated with biotin-sense-HITT determined by WB. (H) PLA analysis of endogenous RGS2/exogenous HITT or HITT-del (1,080–1,130 nt) in HeLa cells. Data derived from 3 independent experiments are presented as mean ± SEM in the bar graph. ****P < 0.0001; NS, not significant by 1-way ANOVA (B and C). Scale bars: 40 μm (left and center panels); 15 μm (right panels).

The key RGS2-binding region in HITT was initially mapped to F3-1 (1,030–1,247 nt) by in vitro binding assay (Supplemental Figure 5A). After that, this fragment was sequentially truncated to 4,100 nt fragments with 50 nt sequence overlap (F3-1.1~4, Figure 4F). Among those, F3-1.1 (1,030–1,130 nt) and F3-1.2 (1,080-1,180 nt) bound with RGS2 to similar extents, suggesting that their overlapping region mapped to 1,080–1,130 nt contains the key nucleotides in binding RGS2 (Figure 4F). No other HITT F3-1 fragmented mutants (F3-1.3 and F3-1.4) were found to bind with RGS2 (Figure 4F).

By mixing truncated RGS2 protein with HITT, we found that C-terminal RGS2 (80–212 aa), containing the RGS domain, is necessary for its binding with HITT (Supplemental Figure 5B). We further identified the most potential residues by analysis of the top 10 RGS2-HITT (1,080–1,130 nt) models predicted by HDOCK (26). Seven RGS2 residues (W80, S81, Y92, R133, F194, Q196, and D197) were identified as the most potentially binding sites in bridging their interaction because they were predicted by these 10 models for at least 5 times and with a root mean square deviation (RMSD) value of less than 3Å (Supplemental Table 3). Then, each of these amino acids was substituted (W80F, S81T, Y92F, R133K, F194Y, Q196R, and D197A), and the combined substitution was generated (W80FS81T and F194YQ196RD197A) when they were close or next to each other (Supplemental Figure 5C). The following RNA pull-down assay revealed that none of the single substitutions had impact on the interaction between RGS2 and HITT (1,080–1,130 nt). However, their interaction was largely diminished by triple mutation at site F194YQ196RD197A (Figure 4G), suggesting that F194, Q196, and D197 form the surface to interact with HITT. The direct interaction between RGS2 and HITT was verified using the proximity ligation assay (PLA) in cells transfected with HITT, but not those transfected with RGS2 binding defective mutant HITT-del (1,080–1,130 nt) (Figure 4H). Thus, HITT directly binds with RGS2 mainly at F194, Q196, and D197 via its (1,080–1,130 nt) fragment. The interaction may be essential for their regulation of PD-L1 (see below).

K175, R176, and S179 in RGS domain are required for PD-L1–5′-UTR binding. We next asked how the RGS2/HITT complex influences PD-L1 translation. To answer this question, we generated 2 luciferase reporter plasmids, PD-L1–5′-UTR and 3′-UTR luciferase reporters (as shown in the diagram, Supplemental Figure 5D). Strikingly, it was with PD-L1–5′-UTR, but not PD-L1–3′-UTR, that luciferase reporter activity was decreased by HITT overexpression and increased by HITT KD (Supplemental Figure 5, E and F). RGS2 KD enhanced PD-L1–5′-UTR luciferase activity and completely abolished the effect of HITT (Figure 5A), confirming that RGS2/HITT imparts negative regulation of PD-L1 expression through the 5′-UTR.

RGS2 physically binds with PD-L1–5′-UTR.Figure 5

RGS2 physically binds with PD-L1–5′-UTR. (A) PD-L1–5′-UTR–driven luciferase activities determined in HITT stable lines with or without RGS2 KD. (B) PD-L1–5′-UTR levels determined by qRT-PCR following CLIP RGS2 in HITT-overexpressing stable HeLa cells, with GAPDH mRNA and CLIP IgG as negative controls. (C) GST-tagged RGS2 protein coprecipitated with biotin–PD-L1–5′-UTR or biotin-PD-L1–5′-UTR antisense control determined by WB. (D) Schematic of the compensatory mutations in PD-L1–5′-UTR (1–36 nt). GST-tagged RGS2 protein coprecipitated with biotin-PD-L1–5′-UTR (1–36 nt) or its mutants, determined by RNA pull-down assay. (E) PLA analysis of endogenous RGS2/exogenous PD-L1–5′-UTR or 5′-UTR (1-36 nt) MT4 in HeLa cells. (F) GST-tagged RGS2 or mutant proteins coprecipitated with biotin–PD-L1–5′-UTR (1–36 nt) determined by RNA pull-down assay. Data derived from 3 independent experiments are presented as mean ± SEM. **P < 0.01; ***P < 0.001; ****P < 0.0001; NS, not significant by Student’s t test (A) and 1-way ANOVA (B). Scale bars: 40 μm (left and center panels); 15 μm (right panels).

We further explored how RGS2/HITT regulates PD-L1–5′-UTR–dependent PD-L1 expression. It has been proposed before that RGS2 inhibits protein translation by binding with eIF2Bε (16). However, this is unlikely for RGS2-regulated PD-L1 expression (Supplemental Figure 5G). Intriguingly, by using a CLIP assay and RNA pull-down assay, as indicated in Figure 4, A and D, we found that RGS2 not only served as a HITT-binding protein as described above (Figure 4, B and E), but also associated with the PD-L1–5′-UTR both in living cells and in vitro (Figure 5, B and C). The extreme 5′ end (1–36 nt) in the PD-L1–5′-UTR is essential for RGS2 binding because the 1–36 nt and 1–72 nt regions, but not 37–108 nt, in the PD-L1–5′-UTR were found to coprecipitate with RGS2 (Supplemental Figure 5H). We then generated 4 compensatory mutants spanning across 1–36 nt PD-L1–5′-UTR, as depicted in Figure 5D. Intriguingly, when 28–36 nt were substituted with their compensatory sequences (MT4), PD-L1–5′-UTR (1–36 nt) lost its RGS2-binding ability (Figure 5D), suggesting that the intact 28–36 nt is required for PD-L1–5′-UTR’s interaction with RGS2. Consistently, PLA-positive RGS2/PD–L1-5′-UTR complexes, but not RGS2/PD-L1–5′-UTR 1–36 nt MT4 complexes, were detected in HeLa cells (Figure 5E).

We also mapped the key PD-L1–5′-UTR–binding residues in RGS2. Similarly to HITT, PD-L1–5′-UTR also bound to RGS2 (80–212 aa), as revealed by the in vitro RNA-binding assay (Supplemental Figure 5I). Following approaches similar to those described in Figure 4G, we predicted a set of residues, D85, N149, K175, R176, and S179, that may mediate RGS2’s binding with PD-L1–5′-UTR using HDOCK (Supplemental Figure 5J and Supplemental Table 3). We tested the binding ability of the single mutants at each of these sites or triple-mutant K175RR176KS179T (Figure 5F) and found that K175RR176KS179T remarkably reduced its binding with PD-L1–5′-UTR. Therefore K175, R176, and S179 provide the major PD-L1–5′-UTR–binding sites of RGS2 (Figure 5F).

HITT forms an RNA-RNA duplex with the PD-L1–5′-UTR. The newly identified binding mechanisms of RGS2/HITT and RGS2/PD-L1–5′-UTR and the coordinated inhibitory effect of HITT and RGS2 on PD-L1 translation inspired us to explore how HITT contributes to RGS2-regulated and 5′-UTR–dependent PD-L1 translation. To this end, we first compared the binding of RGS2/PD-L1–5′-UTR in cells with different expression levels of HITT. The results showed that IFN-γ elevated HITT expression, which was accompanied by increased RGS2/PD-L1–5′-UTR binding (Figure 6A and Supplemental Figure 4C), while inhibition of IFN-γ–induced HITT expression dramatically reduced RGS2/PD-L1–5′-UTR complex levels (Figure 6A). Arbitrarily, expression of HITT produced an effect similar to that of IFN-γ–mediated endogenous HITT overexpression (Figure 6A). These data suggest that HITT facilitates binding between RGS2 and PD-L1–5′-UTR.

HITT forms RNA-RNA duplex with PD-L1–5′-UTR.Figure 6

HITT forms RNA-RNA duplex with PD-L1–5′-UTR. (A) PD-L1–5′-UTR levels determined by qRT-PCR following CLIP RGS2 under IFN-γ treatment with or without HITT KD, with GAPDH mRNA and CLIP IgG as negative controls. (B) Schematic showing in vitro RNA-RNA binding assay to detect the binding between in vitro–synthesized unlabeled HITT and biotin–PD-L1–5′-UTR. (C) HITT and HITT fragments pulled down by biotin–PD-L1–5′-UTR, biotin-PD-L1–5′-UTR fragments, or biotin–antisense–PD-L1–5′-UTR control determined by qRT-PCR with or without RNase H, RNase A, or RNase III. (D) FISH showing colocalization between HITT and PD–L1–5′-UTR in PBS or IFN-γ–treated HeLa cells. (E) Schematic showing complementary sequence (BSs) between HITT and PD-L1–5′-UTR according to the prediction of an online bioinformatic tool (http://rna.informatik.uni-freiburg.de/IntaRNA/Input.jsp). Three PD-L1–5′-UTR mutations, which lost the complementarity site of PD-L1–5′-UTR at BS1 (BS1-MT), BS2 (BS2-MT), and both BS1 and BS2 (BS1+2-MT) were generated and are shown in the diagram. (F) HITT coprecipitated by biotin–PD-L1–5′-UTR (WT or mutants) or biotin–antisense–PD-L1–5′-UTR control determined by qRT-PCR. (G) GST-tagged RGS2 pulled down by biotin-HITT and biotin-antisense-HITT control in the presence of unlabeled FL PD-L1–5′-UTR or PD-L1–5′-UTR mutants determined by WB in an in vitro RNA pull-down assay. Data derived from 3 independent experiments are presented as mean ± SEM. ****P < 0.0001; NS, not significant by 1-way ANOVA (A, C, and F). Scale bars: 20 μm (left 3 panels); 5 μm (right 2 panels).

We further explored how HITT fulfills such a task by testing whether it forms an RNA-RNA complex with PD-L1–5′-UTR. In this RNA-RNA binding assay (27), we found that in vitro–synthesised HITT (unlabeled) was associated with biotin-labeled PD-L1–5′-UTR, but not biotin-labeled antisense PD-L1–5′-UTR (Figure 6, B and C). Remarkably, HITT antisense RNA disrupted the binding between HITT and PD-L1–5′-UTR (Supplemental Figure 6A). In addition, their binding was completely abrogated by RNase III or RNase A, but not RNase H (Figure 6C), suggesting the double-stranded RNA (HITT/PD-L1–5′-UTR) is formed. Furthermore, the colonization of HITT/PD-L1–5′-UTR was detected by FISH using Cy3-labeled HITT probe and FAM-labeled PD-L1–5′-UTR probe in cells under both basal and IFN-γ–treated conditions (Figure 6D).

The RNA-RNA binding assay also revealed that HITT F3 (1,030–2,050 nt) and F3-1 (1,030–1,247 nt), but not other mutant fragments, contributed to PD-L1–5′-UTR binding (Figure 6C). The binding motif between F3-1 (1,030–1,247 nt) and PD-L1–5′-UTR was further analyzed using an RNA-RNA interaction bioinformatic tool, IntaRNA. The highest potential binding site between 2 RNA molecules was predicted to be 83–89 nt (binding site 1 [BS1]) and 97-105 nt (BS2) in PD-L1–5′-UTR (Figure 6E). To validate this bioinformatic result, point mutations on the PD-L1–5′-UTR that aimed to disrupt the RNA-RNA duplex were synthesized, as shown in Figure 6E. No binding was detected between HITT and the biotin-labeled BS2-MT and BS1+2-MT PD-L1–5′-UTRs in the in vitro binding assay (Figure 6F), whereas WT and BS1-MT PD-L1–5′-UTRs, both of which retained the ability to bind with HITT, were found to dramatically improve RGS2’s binding with the streptavidin magnetic beads to pull down biotin-HITT. However, the BS2-MT and BS1+2-MT PD-L1–5′-UTRs, the 2 HITT binding-defective mutants, failed to do so (Figure 6G). Neither BS1 nor BS2 influenced PD-L1–5′-UTR’s binding with RGS2 (Supplemental Figure 6B), which is consistent with above data showing that 1–36 nt is essential for PD-L1–5′-UTR/RGS2 binding (Supplemental Figure 5H). In addition, HITT strengthened the binding between RGS2 and PD-L1–5′-UTR-WT or BS1-MT, but not the binding between RGS2 and PD-L1–5′-UTR-BS2-MT or BS1+2-MT (Supplemental Figure 6B). These data show that HITT bridges and strengthens the interaction of PD-L1–5′-UTR with RGS2 by direct interaction with PD-L1–5′-UTR at BS2 (Supplemental Figure 6C).

HITT/PD-L1–5′-UTR/RGS2 interactions are essential for PD-L1 inhibition. To validate a model where 3 molecules interact to inhibit PD-L1 translation, anti-biotin–conjugated beads were used to pull down biotin-labeled PD-L1–5′-UTR and its possible binding partners in the mixture. As shown, coprecipitated HITT was gradually increased with rising doses of digoxin-labeled HITT in the mixture (Figure 7A). Intriguingly, despite the same amount of RGS2 protein in the mixture, its binding with PD-L1–5′-UTR was also gradually increased with rising doses of HITT (Figure 7A). Therefore, the increased HITT not only enhances its own binding with PD-L1–5′-UTR, but also facilitates the binding of RGS2 with PD-L1–5′-UTR, suggesting the 3 molecules form one complex. We also found that HITT lost its ability to improve the binding between PD-L1–5′-UTR and PD-L1–5′-UTR binding–deficient RGS2 (K175RR176KS179T) (Figure 7A), suggesting that HITT recruits RGS2 to the complex and also promotes direct binding between RGS2 and PD-L1–5′-UTR (Supplemental Figure 6C).

RGS2, HITT, and PD-L1–5′-UTR interaction is required for PD-L1 inhibition.Figure 7

RGS2, HITT, and PD-L1–5′-UTR interaction is required for PD-L1 inhibition. (A) The interactions between RGS2, HITT, and PD-L1–5′-UTR RNA determined by RNA pull-down assays. (B) PD-L1 protein levels after transfection with HITT, F3-1, F3-1.1, F3-1.2, F3-1.3, and F3-1.4 into HeLa cells. HITT and its mutant overexpression efficiencies were measured by qRT-PCR, and PD-L1 intensities were quantified and are shown in bar graph. (C and D) Reporter activities of the indicated luciferase reporters before and after RGS2 overexpression (C) or HITT overexpression (D). Data derived from 3 independent experiments are presented as mean ± SEM.*P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001; NS, not significant by 1-way ANOVA test (BD).

We then tested the essential roles of their interaction in regulating PD-L1 expression. First, the impact of the binding of RGS2 with HITT or PD-L1–5′-UTR was tested after overexpression of RGS2 WT, RNA-binding defective mutants (M2, K175RR176KS179T and M2, 194YQ196RD197A) and the combined mutant (M3, K175RR176KS179T-194YQ196RD197A) in HeLa cells. The expression of PD-L1 was examined by WB. The HITT or PD-L1–5′-UTR–binding defective mutants repressed PD-L1 expression, despite a relatively low efficiency when compared with WT RGS2 (Supplemental Figure 6D), whereas the combined substitution of all 6 amino acids completely abolished RGS2’s ability to inhibit PD-L1 (Supplemental Figure 6D). These data suggest that both bindings (RGS2/HITT and RGS2/PD-L1–5′-UTR) are essential for RGS2-mediated PD-L1 inhibition.

Second, the essential roles of HITT-mediated RGS2 binding were validated by another assay. As shown in Figure 7B, the fragments containing 1,080–1,130 nt HITT, such as full-length HITT, F3-1, F3-1.1, and F3-1.2, were able to inhibit PD-L1 expression (Figure 7B). The other fragments (F3-1.3 and F3-1.4) failed to do so (Figure 7B), further suggesting that the physical interaction between HITT and RGS2 is required for HITT-regulated PD-L1 inhibition.

Third, using luciferase reporter assays, we found that RGS2 binding defective mutant PD-L1–5′-UTR-MT4 (compensatory mutation at 28–36 nt), but not the other mutant reporter, failed to respond to RGS2 overexpression (Figure 7C), which provides additional evidence that RGS2/PD-L1–5′-UTR binding is essential for RGS2-mediated PD-L1 inhibition.

Fourth, the critical roles of HITT/PD-L1–5′-UTR interactions in regulating PD-L1 expression were also examined. We found that HITT inhibited the activities of PD-L1–5′-UTR luciferase reporters with intact HITT BS2, such as WT and PD-L1–5′-UTR-BS1-MT reporter, and failed to change the luciferase reporter activities of PD-L1–5′-UTR-BS2-MT or BS1+2-MT (Figure 7D). These data suggest that the intact HITT BS2 is necessary for HITT-mediated PD-L1 inhibition. These data show that the 3-way interaction among HITT, PD-L1–5′-UTR, and RGS2 is critical for the inhibition of PD-L1 translation.

HITT inhibits T cell immunity in a PD-L1–dependent manner. Given the essential role of HITT in inhibiting PD-L1 expression, we compared the killing effects of CTLs before and after blocking PD-L1 signaling via anti–PD-1 antibody in foreign antigen chicken OVA-expressing 4T1 cells (4T1-OVA). We consistently detected an increased killing effect of OT-I T cells after coculture with HITT-overexpressing 4T1-OVA cells (Figure 8A). Anti–PD-1 antibody increased the killing effect of CTLs, as reported previously (28). The HITT-regulated CTL killing effect was completely abrogated by blocking PD-L1 signaling (Figure 8A). Consistently, a similar effect of HITT on the killing effect of human CTLs after coculture with HITT overexpressing MDA-231 and HeLa cells was observed (Figure 8B and Supplemental Figure 7, A and B). Anti–PD-1 antibody or PD-L1 KD increased the killing effect of CTLs. The HITT-regulated CTL killing effect was completely abrogated by blocking PD-L1 signaling (Figure 8, B and C, and Supplemental Figure 7, A and B). In contrast, PD-L1 overexpression repressed CTL-mediated cancer cell killing effects, and it also abolished HITT-induced killing effects of CTL (Supplemental Figure 7C). In line with these data, HITT lost its ability to regulate expression levels of IL-2 and IFN-γ after anti–PD-1 treatment (Supplemental Figure 7D). These data demonstrate that HITT mainly regulates T cell immunity by suppressing PD-L1 expression. Consistently, HITT KD increased the binding of PD-1 protein to the surfaces of cancer cells, as shown in a PD-1–binding assay (Figure 8D). Thus, HITT markedly enhances T cell cytotoxicity by inhibiting PD-L1 expression in cancer cells, leading to reduced interaction between PD-L1 and PD-1.

HITT enhances T cell–mediated tumor cell–killing efficacy in a PD-L1–dependFigure 8

HITT enhances T cell–mediated tumor cell–killing efficacy in a PD-L1–dependent manner. (A) Detection of the attached 4T1-OVA cells by crystal violet staining after coculture with the activated mouse OT-I T cells for 2 days in the presence of anti–PD-1 antibody or IgG control. Intensities are shown in bar graph. (B and C) Detection of the attached MDA-231 and HeLa cells by crystal violet staining after coculture with the activated T cells for 6 hours in the presence of anti–PD-1 antibody or IgG control. Intensities are shown in bar graphs. (D) Immunostaining of PD-1 (fused to Ig-Fc) on HITT KD MDA-231 cells. PD-L1 fluorescence intensities at cell edge were quantified, and relative levels are shown in bar graph (right). HITT KD efficiency was determined by qRT-PCR. Data derived from 3 independent experiments are presented as mean ± SEM. *P < 0.05; **P < 0.01; *** P < 0.001; **** P < 0.0001; NS, not significant by Student’s t test (AC) and 1-way ANOVA (D). Scale bars: 10 μm.

HITT inhibits tumor growth in vivo by preventing PD-L1–mediated T cell deactivation. We next explored whether HITT promotes T cell immunity in vivo using the 4T1/immune-competent BALB/c orthotopic model of murine mammary carcinoma. HITT-overexpressing orthotopic tumors grew relatively slowly compared with control tumors (Figure 9, A–C). Anti–PD-1 antibody dramatically suppressed tumor growth compared with the corresponding controls. Intriguingly, the effect of HITT was compromised, but not completely abolished, by anti–PD-1 (Figure 9, A–C). The above data were validated using HITT-expressing lentivirus administration in PD-L1–KO tumors (Supplemental Figure 8, A–F). In contrast to HITT, PD-L1–5′-UTR binding defective HITT mutant (HITT-Mut) elicited little antitumor effect. Such a striking difference was completely abolished by PD-L1 KD (Supplemental Figure 8, D–F). HITT-overexpressing 4T1 tumor–bearing mice and anti–PD-1–treated mice survived significantly longer compared with control 4T1 tumor–bearing mice treated with IgG control (Figure 9D). Anti–PD-1–treated HITT-overexpressing 4T1 tumor–bearing mice survived longest among the 4 groups (Figure 9D). These data suggest that blocking PD-L1–mediated T cell inactivation by either anti–PD-1 antibody and/or HITT increases the survival of mammary tumor–bearing mice by suppressing tumor growth with low toxicity (Figure 9E).

HITT inhibits tumor growth by attenuating PD-L1–mediated T cell deactivatioFigure 9

HITT inhibits tumor growth by attenuating PD-L1–mediated T cell deactivation in vivo. (AC) Volume (A), images (B), and tumor weight (C). Each dot represents an evaluation in an individual tumor. (D) Kaplan-Meier survival curve of mice bearing syngeneic 4T1 tumor with treatment of IgG or anti–PD-1. (E) Body weights of BALB/c mice measured with treatments. (F) PD-L1 protein levels determined by WB. (G) Immunostaining of CD8+ IFN-γ+ in CD3+ T cell populations from isolated tumor-infiltrating lymphocytes in syngeneic tissues. Each dot represents an evaluation in an individual tumor. (H) HITT levels in 4T1 syngeneic determined by qRT-PCR. Data in A, CE, and G are shown as mean ± SD. *P < 0.05; **P < 0.01; ***P < 0.001; ***P < 0.0001; NS, not significant by 2-way ANOVA (A and E, n = 6 mice per group), 1-way ANOVA (C and G, n = 6 mice per group), log-rank test (D, n = 10 mice per group), and Student’s t test (H). Data derived from 3 independent experiments are presented as mean ± SEM.

Furthermore, HITT inhibited PD-L1 expression in orthotopic 4T1 tumors (Figure 9F and Supplemental Figure 8, G and H). In addition, a significant increase of the activated tumor-infiltrated CD8+ T cell population (CD3+CD8+IFN-γ+) was detected in HITT-overexpressing tumors (Figure 9G). Anti–PD-1 antibody had no obvious effects on HITT or PD-L1 expression (Figure 9H), while treatment led to a significant increase in the activated tumor-infiltrated CD8+ T cell population (Figure 9G). Anti–PD-1 antibody failed to further enhance the tumor-infiltrated CD8+ T cell population in HITT-overexpressing 4T1 tumors (Figure 9G). Unlike in the CD8+ T cell population, tumor growth and mouse survival were both further decreased or prolonged by the combination of anti–PD-1 and HITT overexpression (Figure 9, A–C).

The association between HITT/RGS2 and PD-L1 in breast cancer tissues. qRT-PCR assay revealed that HITT was downregulated in breast cancer tissues compared with the adjacent normal controls (Figure 10A), while PD-L1 protein levels were increased in breast cancer tissues, as indicated by WB assays (Figure 10, B and C). The decreased HITT and increased PD-L1 were both associated with advanced stages of breast cancers (Figure 10, D and E). In addition, a negative association between the fold changes of HITT and those of PD-L1 protein was detected (Figure 10F). RGS2 was also found to be decreased in breast cancer tissues, and its downregulation was more evident in the advanced breast cancers (Figure 10, B, G, and H). Similarly to what occurred with HITT, RGS2 fold change exhibited a negative correlation with PD-L1 protein fold change (Figure 10I). Neither HITT nor RGS2 correlated with the mRNA levels of PD-L1 (Figure 10, J and K). Therefore, RGS2/HITT may contribute to PD-L1 regulation in vivo in human cancer tissues.

RGS2, HITT, and PD-L1 are associated with each other in vivo.Figure 10

RGS2, HITT, and PD-L1 are associated with each other in vivo. (A) Expression of HITT in human breast tumors (T) and their paired adjacent normal controls (N) (n = 38) determined by qRT-PCR. (B and C) Representative WB (B) and quantification of PD-L1 proteins (C) in 38 pairs of breast cancer tissues and their adjacent normal controls. (D and E) The correlation between the fold change of HITT (D) and PD-L1 protein (E) and stages. (F) Lineal correlation analysis of the fold changes of HITT expression versus those of PD-L1 protein expression (P = 0.021). (G) Quantification of RGS2 proteins in 38 pairs of breast cancer tissues and their adjacent normal controls. (H) Correlation between fold change of RGS2 protein and TNM stages. (I) Lineal correlation analysis of fold changes of RGS2 protein expression versus those of PD-L1 protein expression (P = 0.012). (J) Lineal correlation analysis of fold changes of HITT expression versus those of PD-L1 mRNA expression. (K) Lineal correlation analysis of fold changes of RGS2 protein expression versus those of PD-L1 mRNA expression. (L) Schematic diagram of RGS2/HITT/PD-L1–regulated interaction between cancer cells and T cells to modulate tumor immunity. IFN-γ secreted by activated T cells or others triggers E2F1-mediated transactivation of lncRNA HITT in cancer cells, where HITT directly binds with RGS2 and PD-L1–5′-UTR. This function of HITT also strengthens the direct interaction between RGS2 and PD-L1–5′-UTR. These interactions among HITT, RGS2, and PD-L1–5′-UTR lead to a retarded translation of PD-L1 and elevated T cell activation. Such activity of HITT is impaired in cancer cells due to the reduced expression of HITT. Activating HITT in cancer cells is a potential treatment for elevating T cell immunity. Data derived from 3 independent experiments are presented as mean ± SEM (A and CK). **P < 0.01, Student’s t test (A, CE, G, and H). Correlations were calculated according to Pearson’s correlation (F and IK).

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