Regulation of CD8 T cell differentiation by the RNA-binding protein DDX5

Abstract

The RNA-binding protein DDX5 is a polyfunctional regulator of gene expression, but its role in CD8⁺ T cell biology has not been extensively investigated. In this study, we demonstrate that deletion of DDX5 in murine CD8⁺ T cells reduced differentiation of terminal effector (TE), effector memory (T_EM), and terminal effector memory (t-T_EM) cells while increasing the generation of central memory (T_CM) cells; forced expression of DDX5 elicited the opposite phenotype. DDX5-deficient CD8⁺ T cells exhibited increased expression of genes that promote T_CM cell differentiation, including Tcf7 and Eomes. Together, these findings reveal a role for DDX5 in regulating the differentiation of effector and memory CD8⁺ T cell subsets in response to microbial infection.

Introduction

CD8⁺ T cells are a key component of the adaptive immune response. Activated CD8⁺ T cells differentiate into heterogeneous cell populations that provide efficient pathogen clearance and durable protection against reinfection. Terminal effector (TE) cells are responsible for mediating pathogen clearance through the production of inflammatory cytokines and cytolytic granules, whereas memory cells provide potent recall responses in the event of reinfection (1). The circulating memory cell population is comprised of several distinct subsets, including central memory (T_CM), effector memory (T_EM), and terminal effector memory (t-T_EM) cells that each possess unique functional, migratory, and survival properties (2, 3, 4, 5). T_CM cells express key transcription factors, such as Tcf7, Eomes, and Foxo1, that enable their stem cell-like capacity to expand, differentiate, and self-renew (6). T_EM and t-T_EM cells are characterized by potent effector function upon infectious rechallenge.

RNA-binding proteins are critical regulators of cellular differentiation due to their broad influence on RNA metabolism and gene regulation (7, 8). However, the understanding of their impact on T cell differentiation is still developing. We previously reported that the RNA-binding protein, Dead box protein 5 (DDX5), regulates the differentiation of an effector-like tissue-resident memory cell population (9). These findings, coupled with recent evidence suggesting a role for DDX5 in regulating the effector function of Th17 cells (10), raised the possibility that DDX5 might also regulate differentiation of circulating effector and memory CD8⁺ T cell subsets.

DDX5 is a member of the DEAD box family of RNA helicases that are named for a conserved DEAD (Asp-Glu-Ala-Asp) motif that is important for ATP binding, hydrolysis, and RNA helicase activities (11, 12). DDX5 has been shown to be crucial for cell development, proliferation, and organ maturation, and has been implicated in the tumorigenesis of many cancer types (13). DDX5 has been reported to influence nearly every modality of gene regulation, including chromatin organization and epigenetic remodeling, transcriptional co-activation and co-repression, and alternative splicing of RNA transcripts and RNA metabolism (11). Although DDX5 is a well-known polyfunctional regulator of gene expression, its role in CD8⁺ T cell biology has not been extensively explored.

In this study, we report the consequences of T cell-specific deletion of DDX5 in the differentiation and function of CD8⁺ T cells responding to lymphocytic choriomeningitis virus (LCMV) infection. DDX5 deficiency reduced the proportions of KLRG1^hi CD127^lo TE cells and increased the proportions of the KLRG1^loCD127^hi MP cell population at day 7 post-infection. Furthermore, DDX5 deficiency reduced the proportions of T_EM and t-T_EM cells, and increased the proportion of T_CM cells at day 30 post-infection. DDX5-deficient CD8⁺ T cells exhibited decreased expression of genes associated with terminally differentiated effector cells, including Klrg1, Zeb2, and Cx3cr1, and increased expression of genes associated with T_CM cell differentiation, including Tcf7 and Eomes. Ectopic expression of DDX5 increased the proportions of T_EM and t-T_EM cells and decreased the proportion of T_CM cells, and was associated with reduced expression of Tcf7 and Eomes. Taken together, these findings suggest that DDX5 may regulate CD8⁺ T cell differentiation by virtue of repressing genes that promote the generation of T_CM cells.

Materials and Methods

Mice

All mice were housed under specific pathogen–free conditions in an American Association of Laboratory Animal Care–approved facility at UCSD, and all procedures were approved by the UCSD Institutional Animal Care and Use Committee. C57BL6/J (CD45.1.2⁺ or CD45.2⁺) and P14 TCR transgenic (CD45.1⁺ or CD45.1.2⁺, maintained on a C57BL6/J background) mice were bred at UCSD or purchased from the Jackson Laboratories. Ddx5^fl/fl mice were obtained from Dr. Frances Fuller-Pace’s laboratory (University of Dundee) and have been previously described (14). To generate congenically distinct P14 Ddx5^fl/fl Cd4-Cre⁺ and P14 Ddx5^fl/fl Cd4-Cre⁻ mice, Ddx5^fl/fl mice were crossed to P14 Cd4-Cre⁺ mice (either CD45.1⁺ or CD45.2⁺). All mice were used from 6 to 9 weeks of age, male mice were used as recipients, and male or female mice were used as donors in adoptive transfer experiments. No randomization of blinding was used in infection experiments.

CD8⁺ T cell isolation

For isolation of CD8⁺ T cells from the spleen and peripheral lymph nodes, tissues were collected and filtered through 70 μm cell strainers to yield a single-cell suspension before treatment with Red Blood Cell Lysing Buffer Hybri-Max (Sigma-Aldrich) for 5 minutes. CD8⁺ T cells were then isolated using the CD8a⁺ T Cell Isolation Kit and LS MACS Columns (Miltenyi Biotec) according to the manufacturer’s protocol.

Antibodies and flow cytometry

Surface proteins were stained for 10 minutes on ice in Hank’s Balanced Salt Solution (Corning) supplemented with 1% Fetal Bovine Serum (FBS) with the following antibodies: Vα2 (B20.1), CD8α (53–6.7), CD45.1 (A20), CD45.2 (104)), KLRG1 (2F1/KLRG1), CD127 (A7R34), CD27 (LG.3A10), CX3CR1 (SA011F11), CD62L (MEL-14), all purchased from BioLegend. Samples were then stained in Fixable Viability Dye eFluor780 (Thermo Fisher Scientific) at 1:1000 on ice for 10 min. Cells were then fixed in resuspended in eBio Fix/Perm (eBioscience) for 30 minutes at room temperature. After fixation, intracellular targets were stained in eBio Permeabilization Buffer (eBioscience) for 8 hours at 4°C with the following intracellular antibodies: TCF1/7 (C63D9), EOMES (Dan11mag), Ki67 (16A8), T-bet (4B10), and Granzyme B (GB11). For assessment of cytokine production, cells were cultured in the presence of LCMV GP33–41 peptide (GenScript) and Protein Transport Inhibitor Cocktail (Thermo Fisher Scientific) for 3 hours at 37°C. After cell surface and viability staining, cells were fixed and permeabilized using BD Cytofix/Cytoperm (BD Biosciences) for 30 min at room temperature before staining with anti–IFNy (XMG1.2), TNF-α (MP6-XT22), and IL-2 (JES6–5H4) antibodies (all from BioLegend) for 30 min on ice.

For DDX5 staining, after surface marker staining and fixation using eBiosciences Transcription Factor Fixation/Permeabilization Kit, isolated lymphocytes were stained with anti-DDX5 (JM52–30) 1:200–400 in 1X Permeabilization Buffer, at room temperature for 2 hours, followed by staining with Alexa Fluor^® 488 Donkey anti-rabbit IgG (minimal x-reactivity) antibody (Biolegend), 1:100 at room temperature for 1–2 hours.

cDNA synthesis and qRT-PCR analysis

Total RNA was extracted with the RNeasy kit (QIAGEN) and reverse-transcribed using the iScript Select cDNA Synthesis Kit (Bio-Rad Laboratories). Real-time reverse transcription polymerase chain reaction (RT-PCR) was performed using iTaq Universal SYBR Green Supermix (Bio-Rad Laboratories). DDX5 expression data were normalized to Gapdh mRNA levels. hDdx5 primer sequences: Forward: ATGTCGGGTTATTCGAGTGACC; Reverse: ACTTCCTCCAAATCGAGGTGC. Gapdh primer sequences: Forward: TGAGTATGTCGTGGAGTCTAC; Reverse: TGGACTGTGGTCATGAGCC.

CD8⁺ T cell transfers and infection

For naïve CD8⁺ T cell transfers, splenocytes were collected from naïve congenically distinct Ddx5^fl/fl-Cd4-Cre⁺ P14⁺ and Ddx5^fl/fl-Cd4-Cre⁻ P14⁺ mice. To quantify the number of Vα2⁺CD8⁺CD45.1⁺ cells, an aliquot of the isolated lymphocytes was stained with antibodies against Vα2, CD8α, and CD45.1, and analyzed using flow cytometry. Vα2⁺CD8⁺CD45.1⁺ cells (1 × 10⁵) were adoptively transferred into CD45.2⁺ WT recipients 1 day before intraperitoneal (i.p.) infection with 2 × 10⁵ plaque-forming units (PFU) of lymphocytic choriomeningitis virus (LCMV)-Armstrong. At days 7 and 30 p.i., donor mice were sacrificed, their spleens were harvested, and the donor cells were analyzed using flow cytometry.

For induced Ddx5 deletion, Vα2⁺CD8⁺CD45.1⁺ P14 cells from naïve congenically distinct Ddx5^fl/fl-Ert2-Cre⁺ P14⁺ and Ddx5^fl/fl-Ert2-Cre⁻ P14⁺ mice were mixed 1:1 and adoptively transferred into CD45.2⁺ host mice before subsequent LCMV infection. To induce Ddx5 deletion, recipient mice were treated i.p. with 1 mg of tamoxifen diluted in sunflower oil on days 4–7 or 30–33 following infection.

For forced DDX5 (pMThy1.1_hDDX5_FL) experiments, retroviral particles were generated using platinum E cells grown in 10 cm plates with full selection media (DMEM 10% FBS (v/v), 2 mM L-Glutamine, 100 U/mL Penicillin-Streptomycin, 1 ug/mL puromycin and 10 ug/mL blasticidin). Eighteen hours before transfection, selection media was replaced with antibiotic-free media (DMEM, 10% FBS (v/v), 2 mM L-Glutamine). For each 10 cm plate, 10 μg of each construct and 5 μg pCL-Eco helper plasmids were mixed in Opti-MEM (ThermoFisher) to a volume of 700 μl. This was combined with 45 μl TransIT-LT1 Reagent (Mirus) and 655 μl Opti-MEM for 20 minutes at room temperature. The mixture was then added dropwise to each 10 cm plate. Twelve hours later, media was replaced with fresh antibiotic-free media supplemented with cholesterol 30uM and Viralboost 1:500 (Alstem Cell Advancements). The viral supernatant was subsequently harvested at 48 and 72 hours. Retroviral supernatant was filtered through a 0.45 μm syringe filter and stored at −80°C. Naive WT P14 CD8⁺ T cells were plated at a density of 1×106 cells/mL in 24 well plates pre-coated with 5 μg/ml each of anti-CD3 (clone 3C11) and anti-CD28 (clone 37.51) with T cell media supplemented with IL-2 (100 U/ml). The cells were then centrifuged for 90 minutes at 2000 rpm. Retroviral supernatant was removed and replaced with stored T cell media and incubated at 37°C for 2 hours before adoptive transfer.

For co-transfer experiments, cells of each construct were mixed at a 1:1 ratio and a total of 5×10⁵ donor cells/mouse was adoptively transferred into CD45.2⁺ male recipient mice. One hour later, mice were infected with 2×10⁵ PFU LCMV. Seven and 30 days after infection, mice were euthanized, spleens were harvested, and the donor cells were analyzed by flow cytometry.

For ex vivo re-stimulation assays, P14 CD8⁺ T cells were plated in a U-bottom 96-well plate at 10–15× 10⁶ cells/well in the presence of 1 ng/μl LCMV GP_33–41 peptide (Genscript) and 1X Protein Transport Inhibitor Cocktail (eBioScience) for >3 h at 37°C. Cells were then fixed and permeabilized using BD Cytofix/Cytoperm (BD Biosciences) and stained for IFNγ (XMG1.2), TNF-α (MP6-XT22), and IL-2 (JES6–5H4) antibodies, all purchased from BioLegend, for 30 min on ice. Flow cytometry was performed on a Novocyte (ACEA Biosciences). FACS sorting of cells was done on a FACSAria Fusion or FACSAria2 (BD Biosciences). FlowJo software (BD Biosciences) was used for analysis of flow cytometry data.

CITE-sequencing and analysis

WT CD45.1 and Ddx5^fl/flCd4-Cre⁺ CD8⁺ CD45.1.2 P14 cells were adoptively co-transferred into congenically distinct hosts. The host mice were infected with 2×10⁵ PFU LCMV 24 hours after transfer and sacrificed on days 4, 7, and 30 post-infection. Circulating lymphocytes were isolated from the spleen and resuspended with antibody staining buffer containing TruStain FcX^™ PLUS (anti-mouse CD16/32; Biolegend). Cells were incubated with TotalSeq-A antibody panel (Supplemental Table 1) and fluorescently labeled antibodies targeting CD8, Vα2, CD45.1, CD45.2 for 30 min at 4°C. Stained cells were washed with antibody staining buffer three times before sorting for P14 CD8⁺ T cells of each genotype. About 10,000 cells per sample were loaded into Single Cell G chips (10x Genomics) and partitioned into Gel Bead In-Emulsions in a Chromium Controller (10x Genomics). Single-cell RNA libraries were prepared according to the 10x Genomics Chromium Next GEM Single Cell 3’ v3.1 (Dual Index) User Guide and sequenced on a HiSeq 4000 (Illumina).

Reads from CITE-seq were aligned to mm10 using Cellranger (v.6.0.1). The Unique Molecular Identifier count matrix was used for downstream analysis using Seurat (v4.1.1). In R, samples were subsampled to 2000 cells per sample. Cells were further filtered to exclude cells with < 300 genes per cell and <5% mitochondrial genes. Standard parameters were used for normalization, filtering, PCA reduction, and UMAP creation. The samples were clustered with Louvain clustering using the FindClusters function with a resolution of 1. Differential expression analysis was completed between clusters and conditions using the FindMarkers function. Data are available at Gene Expression Omnibus under GSE230028.

Results

DDX5 may regulate CD8⁺ T cell proliferation and apoptosis

To assess phenotypic changes in circulating CD8⁺ T cell populations resulting from deletion of DDX5, we adoptively co-transferred P14 CD8⁺ T cells, which express a transgenic T cell receptor (TCR) that recognizes the immunodominant epitope of lymphocytic choriomeningitis (LCMV), from mice with a T cell-specific deletion of Ddx5 (Ddx5^fl/fl Cd4-Cre⁺, “Ddx5^cKO”) and congenically distinct control mice (Ddx5^fl/fl Cd4-Cre⁻,“WT”) into CD45.2⁺ recipient mice. Relatively high numbers of P14 cells were transferred (2 × 10⁵ CD8⁺ P14 cells total), which may alter the kinetics and phenotype of the ensuing CD8 T cell response, representing a limitation of the present study (15). One day after cell transfer, recipient mice were infected with the Armstrong strain of LCMV. At days 7 and 30 p.i., lymphocytes were isolated from the spleens of recipient mice and donor cells were analyzed by flow cytometry (Fig. 1A). DDX5-deficient CD8⁺ T cells were reduced compared to their control counterparts at days 7 and 30 post-infection (p.i.), suggesting a possible role for DDX5 in cellular proliferation and/or survival (Fig. 1B). Further, the altered frequencies of DDX5-deficient CD8⁺ T cells compared to control cells remained relatively consistent through the duration of acute infection (Fig. 1C, 1D). Consistent with the notion of DDX5 repressing cellular proliferation, DDX5-deficient CD8⁺ T cells expressed higher levels of Ki-67, a marker of cellular proliferation (Fig. 1E, 1F), compared to control cells. To further elucidate the role of DDX5 in proliferation, we stained WT and DDX5-deficient CD8⁺ T cells with the fluorescent dye CFSE, which dilutes as cells divide, allowing for efficient tracing of cell proliferation patterns. CFSE-labeled WT and DDX5-deficient cells were mixed at a 1:1 ratio and adoptively transferred into congenically distinct recipients. Forty-eight hours after LCMV infection, recipient mice were sacrificed, and donor cells were analyzed by flow cytometry. DDX5-deficient cells exhibited greater CFSE dilution than control cells (Fig. 1G, 1H; Supplemental Fig. 1A, 1B). Lastly, we observed that DDX5-deficient CD8⁺ T cells expressed higher levels of Annexin V, a membrane-bound apoptotic marker at 48 hours p.i. (Fig. 1I, 1J). Having shown that conditional knock-out of Ddx5 accelerated proliferation and led to an early death phenotype, we next investigated whether deletion of DDX5 after the initial proliferative burst of early infection also affected CD8⁺ T cell survival. Importantly, inducible deletion of Ddx5 during the peak of infection (days 4–7 p.i.) did not result in a numerical deficit in DDX5-deficient cells at days 9 and 30 p.i. (Supplemental Fig. 1C, 1D), suggesting that the DDX5 may only play a role in proliferation and survival during the first few days of infection. Taken together, these results indicate that DDX5 may have an important role in inhibiting apoptosis, leading to the observed decrease of DDX5-deficient cells on days 7 and 30 post-infection.

FIGURE 1.

DDX5 may regulate CD8⁺ T cell proliferation and apoptosis. (A) Experimental set-up. Congenically distinct Ddx5^fl/fl Cd4-Cre⁺ (Ddx5^cKO) cells and Ddx5^fl/fl CD4-Cre⁻ (WT) CD8⁺ P14 T cells were mixed at a 1:1 ratio and adoptively transferred i.v. into CD45.2⁺ hosts. One day after transfer, recipient mice were infected with LCMV, sacrificed at days 7 and 30 p.i., and donor cells analyzed by flow cytometry. (B) Representative flow cytometry plot showing the proportions of WT vs Ddx5^cKO cells (left) and ratio of the absolute numbers of Ddx5^cKO to WT CD8⁺ P14 cells at days 7 and 30 p.i., normalized to the input ratio (right). (C) Absolute numbers of WT vs. Ddx5^cKO CD8⁺ P14 cells at days 7, 14, 30 and 60 post-infection. (D) Ratio of Ddx5^cKO to WT CD8⁺ P14 cells at days 7, 14, 30 and 60 post-infection, normalized to the input ratio. (E-F) Representative flow cytometry plots and histograms showing the frequency, absolute numbers, and MFI of Ki67⁺ WT vs, Ddx5^cKO CD8⁺ P14 cells at day 7 post-infection. Gates were determined by Ki67 fluorescence minus one (FMO). (G-H) CFSE proliferation analysis of WT and Ddx5^cKO CD8⁺ T cells at 48 hours p.i., represented as (G) flow cytometry plots (left), histograms (right). (H) Frequency and absolute numbers of CFSE-labeled undivided, Division 1 (Div 1), Division 2 (Div 2), Division 3 (Div 3), and CFSE⁻ WT vs. Ddx5^cKO CD8⁺ P14 cells. (I-J) Representative flow plots and histograms and showing the frequency, absolute numbers, and MFI of Annexin V⁺ WT and Ddx5^cKO CD8⁺ P14 cells at 48 hours post-infection. All data are from one representative experiment out of two independent experiments with n = 5 to 10 per group: ns, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001 (Student’s paired two-tailed t test). Graphs indicate mean ± SEM, symbols represent individual mice.

DDX5 regulates CD8⁺ TE, T_EM, and t-T_EM cell differentiation

We next examined the consequences of T cell-specific deletion of Ddx5 (Ddx5^fl/fl Cd4-Cre⁺, “Ddx5^cKO”) on circulating effector and memory cell populations (Fig. 1A). Ddx5 deletion resulted in reduced proportions and absolute numbers of KLRG1^hiCD127^lo TE-phenotype cells and increased proportions of KLRG1^loCD127^hi MP-phenotype cells at day 7 p.i. (Fig. 2A, Supplemental Fig. 1E). Furthermore, DDX5 deletion resulted in increased proportions of CD62L^hiCD127^hi T_CM cells at day 30 post-infection (Fig. 2B; Supplemental Fig. 1F). Conversely, Ddx5^cKO memory cells displayed reduced proportions and absolute numbers of CD62L^loCD127^hi T_EM cells and CD62L^loCD127^lo t-T_EM cells at day 30 post-infection (Fig. 2B, 2C). Moreover, compared to control cells, DDX5-deficient memory cells exhibited increased expression of CD62L at day 30 p.i. (Supplemental Fig. 2A). Consistent with the observed increase in T_CM cell differentiation, CD127 (Il7r) and CD122 (Il15rb) expression were both increased in the absence of DDX5, suggesting that these cells may be better able to compete for IL-7 and IL-15 (Supplemental Fig. 2B, 2C). Analyses of functional parameters demonstrated that compared to control cells, DDX5-deficient CD8⁺ T cells displayed equivalent frequencies of IFNγ⁺TNF⁺ cells, whereas the frequency of IL-2⁺ DDX5-deficient CD8⁺ T cells was increased compared to controls, upon ex vivo restimulation with GP_33–41 peptide (Fig. 2D, 2E). Moreover, IFNγ, TNF, IL-2 expression and IFNγ⁺ IL-2⁺ co-expression were unaffected in Ddx5^cKO cells, compared to controls (Supplemental Fig. 2D-G). Importantly, analysis of pre-transfer WT and Ddx5^cKO cells showed similar frequencies of naïve CD44^loCD62L^hi CD8⁺ T cells (Supplemental Fig. 3A). In addition, naïve pre-transfer WT and Ddx5^cKO CD8⁺ T cells exhibited similar expression of the transcription factor TCF1, which is expressed by all naïve CD8⁺ T cells, suggesting that the phenotypes observed in Ddx5^cKO cells were not a consequence of altered thymic selection or homeostasis (Supplemental Fig. 3B, 3C). Taken together, these data demonstrate that deletion of DDX5 changes the composition of the antigen-experienced CD8⁺ T cell pool.

FIGURE 2.

DDX5 regulates CD8⁺ TE, T_EM, and t-T_EM cell differentiation. (A) Representative flow cytometry plots (left) and quantification (right) of WT and Ddx5^cKO TE- and MP-phenotype cells. (B) Representative flow cytometry plots (left) and quantification (right) of WT and Ddx5^cKO T_CM, T_EM, t-T_EM cells at day 30 p.i. (C) Ratio of Ddx5^cKO and WT T_CM-, T_EM-, t-T_EM-phenotype CD8⁺ P14 cells at day 30 post-infection. (D) Representative flow cytometry plots (bottom) and quantification (top) of TNF⁺IFNy⁺ WT and Ddx5^cKO cells isolated at day 30 p.i. and restimulated ex vivo in the presence of GP_33–41 peptide for 3 hours. (E) Representative flow plots (left) and quantification (right) of WT and Ddx5^cKO IL-2⁺ CD8⁺ T cells after ex vivo restimulation at day 30 p.i. (F) Experimental set-up. Congenically distinct CD8⁺ T cells were activated and transduced with empty vector (EV) or forced Ddx5 expression (Ddx5^OE) retroviral constructs. Transduced cells were mixed at a 1:1 ratio and adoptively transferred into CD45.2⁺ recipients. Recipient mice were infected with LCMV and sacrificed at days 7 and 30 p.i. (G) Representative flow cytometry plots (left) and quantification (right) of EV and Ddx5^OE TE- and MP-phenotype cells at day 7 p.i. (H) Representative flow cytometry plots (left) and quantification (right) of EV and Ddx5^OE T_CM, T_EM, t-T_EM cells at day 30 p.i. (I) Ratio of Ddx5^OE and WT T_CM-, T_EM-, t-T_EM-phenotype CD8⁺ P14 cells at day 30 post-infection. (J) Representative flow plots (left) and quantification (right) of EV and Ddx5^OE IFN-γ⁺ and TNF⁺ CD8⁺ T cells after ex vivo restimulation at day 30 p.i. (I) Representative flow plots (left) and quantification (right) of EV and Ddx5^OE IL-2⁺ CD8⁺ T cells after ex vivo restimulation at day 30 p.i. All data are from one representative experiment out of two independent experiments with n = 5–10 per group; ns, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001; student’s paired two-tailed t test (A,B,D,E,G,H,J,K), one sample t test (C and I). Graphs indicate mean ± SEM, symbols represent individual mice.

Since DDX5 deletion appeared to inhibit terminal effector cell differentiation and promote the generation of T_CM cells, we next assessed the functional consequences of ectopic expression of DDX5. We isolated and activated congenically distinct CD8⁺ P14 T cells, retrovirally transduced the cells with empty vector (EV) or DDX5 ectopic-expression (Ddx5^OE) constructs, mixed them at a 1:1 ratio, and adoptively co-transferred the cells into CD45.2⁺ mice (Fig. 2F). Recipient mice were sacrificed at days 7 and 30 p.i. and donor cells were analyzed using flow cytometry. In contrast to decreased proportions of TE cells and increased proportions of T_CM cells resulting from Ddx5 deletion, ectopic expression resulted in increased proportions and absolute numbers of TE cells and reduced proportions of MP cells at day 7 p.i., along with decreased proportions of T_CM cells and increased proportions and absolute numbers of T_EM and t-T_EM cells at day 30 post-infection (Fig. 2G, 2H). Furthermore, Ddx5^OE memory cells exhibited reduced CD62L and CD127, while CD122 expression was unaffected (Supplemental Fig. 2H-J). In contrast to conditional knock-out of Ddx5, a greater proportion of Ddx5^OE cells expressed TNF and IFNγ compared to EV controls, and TNF expression was increased among Ddx5^OE cells (Fig. 2K, Supplemental Fig. 2K, 2L). Additionally, while the proportions of IL-2-producing or IL-2 and IFNγ – co-producing memory cells were not affected by ectopic expression of Ddx5, the expression of IL-2 was enhanced in Ddx5^cKO CD8⁺ T cells (Fig. 2L, Supplemental Fig. 2M, 2N). Taken together, these findings indicate that DDX5 promotes the differentiation of effector-like subsets of CD8⁺ T cells.

DDX5 regulates the maintenance of established CD8⁺ t-T_EM cells

Having shown that DDX5 plays a role in the generation of memory cells, we next investigated whether DDX5 also plays a role in established memory cells. We therefore used an inducible Cre recombinase model in which the deletion of Ddx5 is temporally regulated by administration of tamoxifen. Congenically distinct Ddx5^fl/fl Ert2-Cre⁻ (WT) and Ddx5^fl/fl Ert2-Cre⁺ (Ddx5^iKO) and CD8⁺ P14 T cells were adoptively co-transferred into CD45.2⁺ recipients and infected with LCMV 24 hours later. Recipient mice received tamoxifen on days 30–33 p.i. and donor cells were analyzed at day 40 p.i. by flow cytometry (Fig. 3A). A reduction in DDX5 protein and mRNA expression was confirmed by flow cytometry and RT-qPCR of sorted WT and Ddx5^iKO CD8⁺ T cells (Supplemental Fig. 3D-F). Induced deletion of Ddx5 in established circulating memory cells resulted in increased absolute numbers of total DDX5-deficient memory cells relative to control cells (Fig. 3B). Among memory T cell subsets, Ddx5 deletion resulted in increased proportions and absolute numbers of T_CM cells and reduced proportions of t-T_EM cells (Fig. 3C). Furthermore, compared to control cells, reduced proportions of DDX5-deficient CD8⁺ T cells could produce IFNγ and TNF (Fig. 3D). Notably, the production of IFNγ and TNF were largely similar among Ddx5^iKO and control cells among the circulating memory cell subsets, except for reduced expression of IFNγ in the Ddx5^iKO T_CM cell population (Fig. 3E, 3F). The frequency of IL-2 producing Ddx5^iKO memory cells was also reduced compared to WT control cells, while IL-2 production was only significantly decreased within the t-T_EM cell population and increased in the T_CM cell population (Fig. 3H). Together, these findings suggest a role for DDX5 in promoting the maintenance of t-T_EM cells.

FIGURE 3.

DDX5 regulates the maintenance of established CD8⁺ t-T_EM cells. (A) Experimental set-up. Congenically distinct Ddx5^fl/fl Ert2-Cre⁻ (WT ) and Ddx5^fl/fl Ert2-Cre⁺ (Ddx5^iKO ) CD8⁺ P14 T cells were mixed at a 1:1 ratio and adoptively transferred into CD45.2⁺ recipients intravenously. One day post-transfer, recipient mice were infected with LCMV. At days 30–33 p.i., mice received 1μg tamoxifen i.p. to induce the deletion of Ddx5. Recipient mice were sacrificed at day 40 p.i. and donor cells were analyzed by flow cytometry. (B) Representative flow cytometry plots (left) and quantification (right) of WT and Ddx5^iKO CD8⁺ P14 cells at day 40 p.i. (C) Representative flow cytometry plots (left) and quantification (right) of WT and Ddx5^iKO T_CM, T_EM, and t-T_EM cells at day 40 p.i. (D) Representative flow plots (left) and quantification (right) of WT and Ddx5^iKO IFN-γ⁺ and TNF⁺ CD8⁺ P14 cells after ex vivo restimulation at day 30 p.i. (E-F) IFNγ and TNF median fluorescence intensity in WT and Ddx5^iKO T_CM, T_EM, and t-T_EM cells at day 40 p.i. (G) Representative flow plots (left) and quantification (right) of WT and Ddx5^iKO IL-2⁺ CD8⁺ T cells after ex vivo restimulation at day 40 p.i. (H) IL-2 median fluorescence intensity of WT and Ddx5^iKO T_CM, T_EM, and t-T_EM cells at day 40 p.i. All data are from one representative experiment out of two independent experiments with n = 5 to 10 per group; ns, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001 (Student’s paired two-tailed t test). Graphs indicate mean ± SEM, symbols represent individual mice.

Genes associated with T_CM cell differentiation are upregulated in the absence of DDX5

To investigate potential mechanisms by which DDX5 regulates CD8⁺ T cell differentiation, we employed Cellular Indexing of Transcriptomes and Epitopes by Sequencing (CITE-seq), which allows for measurements of the transcriptome and selected proteins in the same single-cells (16). We adoptively co-transferred control WT and Ddx5^cKO CD8⁺ P14 T cells into congenically distinct recipient mice that were infected with LCMV one day after transfer (Fig. 4A). At days 4, 7, and 30 days p.i., splenic CD8⁺ P14 T cells were isolated by FACS and processed for CITE-seq using the 10x Genomics platform. Antibodies targeting cell surface molecules, some of which have previously implicated in CD8⁺ T cell activation and differentiation, were selected for inclusion in the CITE-seq antibody panel (Supplemental Table 1).

FIGURE 4.

Genes associated with T_CM cell differentiation are upregulated in the absence of DDX5. (A) Experimental set-up. Congenically distinct Ddx5^fl/fl Cd4-Cre⁻ (WT) and Ddx5^fl/fl Cd4-Cre⁺ (Ddx5^cKO) CD8⁺ P14 cells were mixed and adoptively transferred into CD45.2⁺ recipients. One day after transfer, recipient mice were infected with LCMV. Mice were sacrificed at days 4, 7, and 30 p.i. and stained with a TotalSeq-A antibody panel (Supplemental Table 1) before FACS-sorting and processing for CITE-seq. (B) UMAP analyses of WT and Ddx5^cKO CD8⁺ T cells on days 4, 7, and 30 p.i. (C) Proportions of WT (red) and Ddx5^cKO (blue) cells within each cluster. (D-F) T_CM, T_EM, and t-T_EM gene signature scores applied to Day 30 clusters, represented as violin plots. (G-I) T_CM, T_EM, and t-T_EM gene signature scores applied to WT and Ddx5^cKO cells within each Day 30 cluster and represented as violin plots. (J) Heatmap representing the expression of selected genes among cells from each of the Day 30 clusters. (K) Quantification of expression level of selected genes among cells from each of the Day 30 clusters, represented as violin plots. (L) Quantification of selected proteins by Day 30 cells, represented as relative expression plots superimposed on individual cells in the UMAP.

Uniform Manifold Approximation and Projection (UMAP) analyses revealed that CD8⁺ T cells clustered distinctly at all time points post-infection (Fig. 4B), indicating that T cells exhibit distinct gene expression patterns during differentiation, as previously demonstrated (16, 9, 17). Notably, WT and Ddx5^cKO CD8⁺ T cells clustered distinctly at days 7 and 30 p.i. (Fig. 4B). Ddx5^cKO CD8⁺ T cells exhibited increased expression of Sell and Il7r, and decreased expression of genes associated with terminal effector differentiation, such as Zeb2, Klrg1, and Bhlhe40, consistent with the observed bias toward T_CM cell differentiation in the absence of DDX5 (Supplemental Fig. 4A). Moreover, pathway analyses revealed an enrichment of genes regulating cytokine production and cytotoxicity in WT cells relative to Ddx5^cKOcells, as well as of processes related to ribosome biogenesis and cytoplasmic translation, consistent with DDX5’s known involvement in ribosome assembly and function (19) (Supplemental Fig. 4B). We next evaluated circulating memory cell heterogeneity at day 30 post-infection. Clusters 0, 9, 15, and 19 were primarily comprised of day 30 p.i. WT CD8⁺ T cells, whereas clusters 2 and 12 were primarily comprised of Ddx5^cKOCD8⁺ T cells (Fig. 4C). Comparative gene ontology analyses among the six day 30 clusters revealed that the Ddx5^cKO clusters were more similar to each other than to the WT clusters (Supplemental Fig. 4C). Application of T_CM, T_EM, and t-T_EM transcriptional signatures to WT and DDX5-deficient memory cells revealed that the four WT-predominant clusters (0, 9, 15, and 19) tended to score more highly for the T_EM, and t-T_EM transcriptional signatures, whereas the two Ddx5^cKO-predominant clusters (2, 12) tended to score more highly for the T_CM gene signature (Fig. 4D-4F). Furthermore, application of these transcriptional signatures to WT and DDX5-deficient memory cells within each cluster demonstrated that WT cells scored higher for the T_EM and t-T_EM gene signatures, while DDX5-deficient cells scored higher for the T_CM gene signature (Fig. 4G-4I). Analyses focused on specific genes revealed that compared to the four WT clusters (0, 9, 15, 19), the two Ddx5^cKO clusters (2, 12) exhibited increased expression of transcriptional regulators associated with T_CM cells, including Bcl2, Tcf7, Lef1, Eomes, Foxo1, and Bach2 (Fig. 4J, 4K). In contrast, the two Ddx5^cKO clusters (2, 12) exhibited reduced expression of genes associated with effector cell differentiation and function, including Zeb2, Tbx21, Gzma, Gzmb, Prf1, Cx3cr1, and Klrg1, compared to the WT clusters (0, 9, 15, 19) (Fig. 4J, 4K). Using protein expression derived from the CITE-seq protein panel, we observed that the WT clusters (0, 9, 15, and 19) tended to express higher levels of molecules associated with terminal effector cells, including CX3CR1 and KLRG1, whereas the two Ddx5^cKO clusters (2, 12) tended to express higher levels of molecules associated with T_CM cells, such as CD62L and CD127 (Fig. 4L, Supplemental Fig. 4D). Together these findings raised the possibility that DDX5 functions to repress the expression of T_CM cell-associated genes to promote the differentiation of effector-like subsets of CD8⁺ T cells.

DDX5-deficient t-T_EM cells exhibit increased TCF1 and Eomes protein expression

The CITE-seq analyses indicated that a major consequence of DDX5 deletion was increased expression of genes that promote T_CM cell generation. We therefore sought to confirm some of the key transcriptional findings with independent flow cytometry experiments. Indeed, greater proportions of DDX5-deficient TE and MP cells expressed TCF1 and DDX5-deficient TE cells displayed increased expression of TCF1 compared to control cells at day 7 p.i. (Fig. 5A, 5B); conversely, ectopic expression of Ddx5 resulted in decreased proportions of TE and MP cells expressing TCF1 and reductions of TCF1 expression in the Ddx5^OE TE and MP cell populations (Fig. 5C, 5D). Analyses of TCF1 expression among DDX5-deficient circulating memory CD8⁺ T cell subsets revealed increased expression of TCF1 at day 30 post-infection (Fig. 5E, Supplemental Fig. 4E). The observed increase in TCF1 expression among circulating memory CD8⁺ T cells was also evident within the T_CM, T_EM, and t-T_EM cell populations and DDX5-deficient CD8⁺ T cells displayed increased proportions of T_CM, T_EM, and t-T_EM cells expressing TCF1 (Fig. 5F, Supplemental Fig. 4F, 4G). Conversely, forced expression of DDX5 resulted in decreased expression of TCF1 among circulating memory CD8⁺ T cells (Fig. 5G), which was significant within the T_CM and T_EM cell subsets (Fig. 5H). Importantly, forced expression of Ddx5 also resulted in decreased proportions of T_CM, T_EM, and t-T_EM cells expressing TCF1 (Supplemental Fig. 4H, 4I). Similarly, Eomes expression was increased in DDX5-deficient CD8⁺ T cells as well as within the T_CM and t-T_EM cell populations (Fig. 5I, 5J); moreover, increased proportions of DDX5-deficient t-T_EM cells expressed Eomes (Supplemental Fig. 4J-L). Conversely, ectopic expression of Ddx5 resulted in reduced proportions of cells expressing Eomes in the total circulating memory CD8⁺ T cell pool and among T_CM, T_EM, and t-T_EM cell subsets (Fig. 5K, 5L). Moreover, ectopic expression of Ddx5 resulted in reduced proportions of Eomes^hi T_CM, T_EM, and t-T_EM cells (Supplemental Fig. 4M, 4N). Furthermore, DDX5-deficient CD8⁺ T cells exhibited increased co-expression of Eomes and TCF1 (Supplemental Fig. 4O). Together, these data suggest that DDX5 may promote the differentiation and maintenance of t-T_EM cells by repressing genes associated with generation of T_CM cells.

FIGURE 5.

DDX5-deficient t-TEM cells exhibit increased TCF1 and Eomes protein expression. (A) Representative flow cytometry plots (left) and quantification (right) of proportions of WT and Ddx5^cKO TE and MP CD8⁺ T cells expressing TCF1 at day 7 p.i. (B) Representative histograms (left) and median fluorescence intensity (right) of TCF1 expression in WT and Ddx5^cKO TE and MP cells at day 7 p.i. (C) Representative flow cytometry plots (left) and quantification (right) of proportions of WT and Ddx5^OE TE and MP CD8⁺ T cells expressing TCF1 at day 7 p.i. (D) Representative histograms (left) and median fluorescence intensity (right) of TCF1 expression in WT and Ddx5^OE TE and MP cells at day 7 p.i. (E) Representative flow cytometry plots (left) and histograms (middle) and quantification (right) of TCF1 MFI in WT and Ddx5^cKO CD8⁺ P14 cells at day 30 p.i. (F) Representative histograms (left) and quantification (right) of proportions of WT and Ddx5^cKO T_CM, T_EM, t-T_EM cells expressing TCF1. (G) Representative flow cytometry plots (left), histograms (middle), and quantification (right) of TCF1 MFI in WT and Ddx5^OE CD8⁺ P14 cells at day 30 p.i. (H) Representative histograms (left) and quantification (right) of proportions of WT and Ddx5^OE T_CM, T_EM, t-T_EM cells expressing TCF1. (I) Representative flow cytometry plots (left) and histograms (middle) and quantification (right) of Eomes MFI in WT and Ddx5^cKO CD8⁺ P14 cells at day 30 p.i. (J) Representative histograms (left) and quantification (right) of proportions of WT and Ddx5^cKO T_CM, T_EM, t-T_EM cells expressing Eomes.(K) Representative flow cytometry plots (left), histograms (middle), and quantification (right) of Eomes MFI in WT and Ddx5^OE CD8⁺ P14 cells at day 30 p.i. (L) Representative histograms (left) and quantification (right) of proportions of WT and Ddx5^OE T_CM, T_EM, t-T_EM cells expressing Eomes. All data are from one representative experiment out of two independent experiments with n = 5 to 10 per group; ns, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001 (Student’s paired two-tailed t test). Graphs indicate mean ± SEM, symbols represent individual mice.

DISCUSSION

RNA-binding proteins are important regulators of cellular differentiation due to their unique ability to operate at multiple levels of gene regulation, but their role in immune cell biology has not been extensively explored (19, 20). Here, we identify the RNA-binding protein DDX5 as a regulator of effector and circulating memory CD8⁺ T cell differentiation in response to microbial infection.

While DDX5 has been implicated in nearly every aspect of genetic regulation from chromatin organization to RNA metabolism and suggested to broadly regulate cell differentiation, the role of DDX5 in T cells during acute infection is only beginning to be explored (21, 22, 23). We show here that DDX5-deficient CD8⁺ T cells exhibited accelerated proliferation and reduced survival compared to controls, implicating DDX5 in the regulation of CD8⁺ T cell proliferation. However, the substantial CFSE dilution was accompanied by a small difference in the proportion of Ki67⁺ cells, suggesting that factors other than cell cycle entry might be contributing to the observed phenotype. Furthermore, consistent with our previously reported role for DDX5 in regulating the differentiation of an effector-like subset of tissue-resident memory cells (9), we demonstrate that deletion of DDX5 resulted in a reduction in the proportions of TE, T_EM, and t-T_EM cells, along with increased proportions of MP and T_CM cells. In established memory cells, Ddx5 deletion promoted maintenance of the T_CM population while simultaneously limiting the t-T_EM population. Some observed differences between WT and Ddx5^iKO cells were modest, representing an important limitation of the present study. Further, DDX5-deficient CD8⁺ T cells exhibited increased expression of key genes known to be involved in T_CM cell differentiation, such as Tcf7 and Eomes. Together, these findings raise the possibility that DDX5 regulates differentiation of effector-like CD8⁺ T cell subsets by virtue of repressing genes that promote T_CM cell generation.

There are several potential mechanisms by which DDX5 may repress expression of genes associated with T_CM cells. Multiple TCF1 isoforms have been identified in T cells (24, 25); the long isoforms possess a beta-catenin binding domain and are dispensable for TE cell differentiation, but are necessary for the generation of T_CM cells (26). Our data suggest that DDX5 may repress TCF1 expression, raising the possibility that DDX5 could regulate the differentiation of TE and t-TEM cells through alternative splicing of TCF1 and shifting the composition of the TCF1 isoform pool to favor the short isoforms (27–29); such a possibility will be investigated in future studies. Furthermore, prior studies have described the intrinsic histone-modifying capabilities of TCF1 and its homolog, LEF1 (30), suggesting the possibility that repression of TCF1 expression by DDX5 may shift the epigenetic landscape to favor expression of genes associated with TE cell differentiation. Another potential mechanism by which DDX5 could repress expression of genes associated with T_CM cells is through an association with the Polycomb Repressive Complex 2 (PRC2), thereby mediating the subsequent silencing of memory genes through deposition of repressive H3K27me3 marks. Indeed, recent reports have posited a role for long non-coding RNAs (lncRNAs) in enabling targeted PRC2-mediated gene silencing (27, 28). Recently published global RNA interactions with DNA by deep sequencing (GRID-seq) data revealed that the lncRNA Malat1 interacts at genomic loci associated with H3K27me3 deposition in TE cells; furthermore, shRNA-mediated knockdown of Malat1 inhibited TE and t-T_EM cell differentiation (32). Coupled with the well-established capacity of DDX5 to bind lncRNAs, these findings raise the possibility that lncRNAs, such as Malat1, may guide DDX5/PRC2 activity for sequence-specific silencing of T_CM cell-associated transcripts in TE and t-T_EM cells. Future studies exploring the putative relationships between DDX5, PRC2, and lncRNAs may reveal previously unknown mechanisms of sequence-specific PRC2-directed CD8⁺ T cell differentiation.

The possibility of DDX5-mediated repression of the T_CM cell-associated transcriptional program may be especially pertinent in the context of chronic infections and cancer. In the current study, deletion of DDX5 appeared to de-repress Tcf7 expression, potentially allowing for the preferential differentiation of T_CM cells. The ability of TCF1 to preserve cellular plasticity has recently garnered a lot of attention, as many groups have shown that this property is essential for maintaining a stem-like population of CD8⁺ T cells that may play a critical role in chronic infections and cancer (29, 30, 31, 32). Furthermore, high expression of DDX5 has been correlated with aggressive tumor features, including treatment resistance, and has been shown to be predictive of poor clinical outcomes in many cancer types (33, (38). Together, these insights suggest that targeting DDX5 in cancer may inhibit DDX5-driven tumorigenesis and drug-resistance within the tumor, as well as potentially skewing the immune response towards TCF1-expressing stem-like CD8⁺ T cells. Future studies evaluating the efficacy of DDX5-deficient CD8⁺ T cells in controlling tumor growth may provide support for DDX5 as a target for cancer therapy.

Overall, our study has identified a role for DDX5 in promoting the differentiation of TE, T_EM, and t-T_EM cells, potentially through repression of T_CM cell-associated gene expression. These findings advance our understanding of the role of RNA-binding proteins in regulating CD8⁺ T cell differentiation and serve as the basis for future studies aimed at improving CD8⁺ T cell responses in infection and cancer.

KEY POINTS.

The RNA-binding protein DDX5 regulates CD8⁺ T cell differentiation.

DDX5 represses the generation of central memory CD8⁺ T (T_CM) cells.

DDX5-deficient T cells exhibit increased expression of T_CM-associated genes.