Purine nucleoside phosphorylase enables dual metabolic checkpoints that prevent T cell immunodeficiency and TLR7-associated autoimmunity

dCK and SAMHD1 mediate the selective lethality of PNP deficiency.

To investigate how PNP inhibition affects human thymopoiesis, we utilized the artificial thymic organoid (ATO) model (14) supplemented with dG and a highly selective and potent PNP inhibitor (PNPi; ulodesine) that rapidly increases serum dG in mice from undetectable levels to 5–20 μM (Supplemental Figure 1A; supplemental material available online with this article; https://doi.org/10.1172/JCI160852DS1), a range similar to that previously observed in PNP-deficient mice and humans (14). In this system, human cord blood–derived hematopoietic stem and progenitor cells (HSPCs) are cultured in defined media as semidry 3D organoids alongside Notch ligand–expressing bone marrow stromal (MS5-hDLL4) cells, which stimulate HSPC differentiation along a developmental trajectory mirroring human thymopoiesis (Figure 1A). Pharmacological PNP inhibition blocks dG catabolism in both stromal and HSPC cell compartments, thereby mimicking dG accumulation in thymi from PNPi-treated mice (Supplemental Figure 1B). To investigate whether dCK plays a role in mediating the effects of PNPi and dG, we used a highly selective and potent dCK inhibitor developed by our group [(R)-DI-87; dCKi] (15). After 5 weeks of culture, most T cell precursors in control ATOs progressed from the CD4/CD8 double-negative (DN) to the CD4/CD8 double-positive (DP) stage (Figure 1B). PNP inhibition triggered a developmental block, evidenced by DN cell accumulation, depletion of DP thymocytes, and an approximately 60% decrease in ATO cellularity (Figure 1B and Supplemental Figure 1C). Concomitant dCK inhibition completely rescued the inhibitory effects of PNPi/dG, indicating that this response requires a functional dCK-dependent nucleoside salvage pathway (Figure 1B and Supplemental Figure 1C).

PNP is expressed across immune lineages (9), but PNP deficiency is associated with T cell immunodeficiency (4). To investigate the metabolic mechanisms responsible for this selective effect, we used T cell acute lymphoblastic leukemia (T-ALL) cells, which are derived from immature T cells and have been shown to be highly susceptible to dG-induced toxicity, alongside B-ALL, peripheral T cell lymphoma (PTCL, derived from mature T cells), and acute myeloid leukemia (AML) cells (16, 17). Similar to our findings using primary human thymocytes (Figure 1B), T-ALL cells were sensitive to PNPi/dG in a dCK-dependent manner (Figure 1C). In contrast, B-ALL, AML, and PTCL cells were uniformly resistant (Figure 1C). Furthermore, PNPi/dG did not impair the proliferation of primary murine CD4⁺ or CD8⁺ T lymphocytes activated ex vivo (Supplemental Figure 1D). To determine whether impaired dCK activity underlies innate resistance to PNPi and dG in non-T-ALL lines, we employed an liquid chromatography–tandem mass spectrometry–multiple reaction monitoring (LC-MS/MS-MRM) approach developed by our group (18) to simultaneously evaluate the contributions of dCK-dependent salvage and ribonucleotide reductase–dependent (RNR-dependent) de novo biosynthesis to DNA replication using [¹⁵N₃]-labeled deoxycytidine ([¹⁵N₃]dC) and [¹³C₆]glucose, respectively (Supplemental Figure 1E). All tested leukemia and lymphoma cells were capable of utilizing [¹⁵N₃]dC for DNA synthesis, indicating that they possess a functional dCK-dependent salvage pathway (Supplemental Figure 1E). Therefore, dCK activity is required but not sufficient for PNPi/dG-mediated lethality.

We reasoned that the activity of other purine metabolism-related genes could dictate sensitivity to PNPi/dG. By examining the expression of such genes across PNPi/dG-sensitive T-ALL versus resistant B-ALL models, we observed that the deoxyribonucleotide triphosphohydrolase SAMHD1 was expressed at low levels in the majority of cancer cell line encyclopedia–annotated (CCLE-annotated) T-ALL lines relative to B-ALL lines (Figure 1D and Supplemental Figure 1F) (19). Analysis of SAMHD1 expression in T-ALL cells versus B-ALL cells included in PRoXe, a public repository of patient-derived leukemia cells (https://www.proxe.org/), revealed a similar pattern, with most T-ALL cells lacking detectable SAMHD1 expression, and the majority of B-ALL cells expressing SAMHD1; dCK was expressed at similar levels in T-ALL cells and B-ALL cells (Supplemental Figure 1G). Moreover, DN murine and human thymocytes, which are sensitive to PNPi/dG, express much lower levels of SAMHD1 than PNPi/dG-resistant mature CD4⁺ and CD8⁺ single-positive T cells (Figure 1E and Supplemental Figure 1H) (20). In contrast, SAMHD1 expression was detectable throughout B cell development (Supplemental Figure 1H), which is largely unaffected by PNP deficiency (4, 21).

To confirm that SAMHD1 protects against PNPi/dG mediated toxicity, we restored SAMHD1 expression in CCRF-CEM (CEM) T-ALL cells (Supplemental Figure 1I) (22). In SAMHD1-deficient CEM-YFP cells, PNPi/dG abolished proliferation and induced markers of replication stress (CHEK1-S345 phosphorylation), DNA damage (H2A.X-S139 phosphorylation), and apoptosis (annexin V), while SAMHD1-expressing isogenic cells were completely resistant (Figure 1F and Supplemental Figure 1, J and K).

SAMHD1 catalyzes the phosphohydrolysis of deoxyribonucleoside triphosphates (dNTPs) to nucleosides and is a critical regulator of intracellular dNTP abundance (23). To evaluate the effects of PNPi/dG and dCKi on dNTP pools and DNA replication on SAMHD1-proficient and -deficient cells, we performed LC-MS/MS-MRM tracing of stable isotope-labeled glucose ([¹³C₆]glucose)- and dG ([¹⁵N₅]dG)-derived metabolites. As depicted schematically in Figure 2A, dGTP isotopologue fractional enrichment delineates the biosynthetic route by which it was derived, either by de novo synthesis from glucose via RNR (identified by [¹³C₅]dGTP) or by salvage of intact dG via dCK (identified by [¹⁵N₅]dGTP). This approach also traces the contribution of guanine nucleobases liberated from dG by PNP to dGTP pools via hypoxanthine phosphoribosyltransferase–mediated (HPRT-mediated) fusion with glucose-derived phosphoribosyl pyrophosphate (identified by [¹⁵N₅,¹³C₅]dGTP). In SAMHD1-proficient cells, PNPi prevented HPRT-dependent guanine nucleobase salvage, but RNR-dependent de novo synthesis fully compensated for this defect and sustained DNA synthesis (Figure 2B). In SAMHD1-deficient cells, PNPi massively expanded the dGTP pool derived from dCK-mediated salvage of intact [¹⁵N₅]dG. This effect was blocked by dCKi, substantially reduced in CEM-SAMHD1 cells, and absent in NALM6 (B-ALL) and HUT78 (PTCL) cells, which endogenously express SAMHD1. In PNPi-treated SAMHD1-deficient cells, dGTP pool expansion was associated with inhibition of DNA replication. PNPi impaired pyrimidine dNTP synthesis in these cells, as evidenced by a restriction of dCTP abundance, decreased de novo dCTP production, and reduced dCTP utilization for DNA replication (Figure 2C). Mechanistically, the restriction of dCTP pools by PNPi/dG in SAMHD1-deficient cells likely reflects a dGTP-mediated allosteric switch in RNR substrate specificity that favors ADP reduction at the expense of CDP reduction (24).

Next, we determined whether the synthetic lethal interaction between PNP and SAMHD1 inactivation extends to solid tumors. HCC827 lung adenocarcinoma cells exhibited the lowest SAMHD1 expression among CCLE-annotated solid tumor cells (Supplemental Figure 2, A and B) and were highly susceptible to PNPi/dG in a dCK-dependent manner (Supplemental Figure 2C). Restoring SAMHD1 expression in HCC827 cells rendered them completely resistant to PNPi/dG-mediated toxicity (Supplemental Figure 2, B and C). Conversely, knocking out SAMHD1 in SUIT2 pancreas cancer cells rendered them hypersensitive to the dCK-dependent lethality of PNPi/dG (Supplemental Figure 2, B and C). As SAMHD1 expression was undetectable in a subset of CCLE melanoma cell lines (Supplemental Figure 2A), we extended our analysis to a panel of melanoma cell lines derived at UCLA. We identified 2 lines (M230 and M418) in which SAMHD1 expression was undetectable both at baseline (Supplemental Figure 2E) and after type I IFN (IFN-β) treatment; in contrast, SAMHD1 was expressed at baseline in M297, M257, and M296 lines and was further increased by IFN-β exposure (Supplemental Figure 2F), consistent with SAMHD1 being an IFN-stimulated gene (25). PNPi/dG elicited dCK-dependent proliferation inhibition in SAMHD1-deficient M230 and M418 cells, while having no effects against SAMHD1-proficient M297 cells (Supplemental Figure 2G). In summary, these findings identify critical roles for dCK and SAMHD1 as mediators for the cellular lethality of PNP inactivation in both lymphocyte and nonlymphocyte models.

Metabolic basis for interspecies differences in the effect of PNP deficiency on T cell development.

PNP-KO mice manifest minor alterations in T cell development and fail to recapitulate the severe T cell immunodeficiency observed in humans with inactivating mutations in PNP (8, 26). Consistent with prior studies in PNP-KO mice, but in contrast to our data in the human ATO system (Figure 1B), we did not observe alterations in thymocyte developmental stage distribution in C57BL/6 mice treated with PNPi for 2 weeks (Figure 3A). To investigate a potential metabolic basis for this interspecies difference, we used 2 orthogonal approaches. First, we examined the effects of PNP inhibition on thymopoiesis in a bone marrow, liver, thymus–humanized (BLT-humanized) mouse model generated by engrafting human fetal liver and thymic fragments under the renal capsule and intravenously delivering hematopoietic stem cells into immunodeficient NOD/SCID IL2Rg^null (NSG) or triple-KO (TKO; lacking RAG2, CD47, and X-linked IL2RG) mice (27). As shown in Figure 3B, PNP inhibition significantly impaired human T cell development in the mice, as evidenced by a CD4/CD8 DN-stage developmental block. which resembles the phenotype observed in the ATO system (Figure 1B). Second, we evaluated the effects of (a) conditioned media (CM) derived from murine (MS5) or human (HS5) bone marrow mesenchymal stromal cell lines, (b) CM from murine bone marrow–derived macrophages (BMDMs), and (c) C57BL/6 mouse sera on PNPi/dG response in SAMHD1-deficient CEM cells (Figure 3C). Strikingly, MS5 and BMDM-derived CM as well as mouse sera completely rescued PNPi/dG-induced lethality, while HS5-derived CM lacked such protective effects (Figure 3D).

Together, these data suggest the existence of a soluble factor that renders SAMHD1-deficient thymocytes and T-ALL cells resistant to PNPi/dG. This factor is present in sera from C57BL/6 mice and CM from murine bone marrow stromal cells and BMDMs but not in sera from humanized mice or CM from human bone marrow stromal cells. We focused on nucleosides as potential resistance factors, as previous work by us and others has shown that stromal cells release these metabolites (25, 28). LC-MS/MS-MRM analyses revealed that resistance-conferring MS5 CM contained substantially more nucleosides uridine, cytidine, and deoxycytidine (dC) than the HS5 CM (Figure 3E and Supplemental Figure 3A). dC emerged as the most plausible resistance factor, given that both dC and dG are salvaged via dCK. dC, however, is the preferred dCK substrate, as indicated by the specificity constant (K_cat/K_M) values 23.1 × 10³ and 2.9 × 10³ for dC and dG, respectively (29). dC levels were also elevated in the resistance-conferring CM from BMDMs (Figure 3E). Consistent with a role for dC in mediating PNPi response in vivo, serum dC levels in BLT mice were significantly lower than in naive NSG mice or C57BL/6 mice and approached levels measured in humans and nonhuman primates (Figure 3F) (30). Moreover, humanized mouse sera exhibited significantly increased dC degradation capacity compared with naive NSG sera (Figure 3G), suggesting an important role for catabolism in regulating dC serum levels.

To confirm a role for dC catabolism in modulating responses to PNPi/dG, we used the dC-catabolizing enzyme cytidine deaminase (CDA). The addition of recombinant CDA abrogated the resistance to PNPi/dG conferred by dC, MS5/BMDM-derived CM, and C57BL/6 mouse serum (Figure 4A and Supplemental Figure 3B). Similar effects were observed using SAMHD1-deficient CEM and HCC827 cells engineered to stably express CDA (Supplemental Figure 3C). dC supplementation abrogated PNPi/dG lethality in parental cells but ectopic CDA expression prevented this rescue (Figure 4B and Supplemental Figure 3D).

We performed [¹³C₆]glucose, [¹⁵N₅]dG, and [¹⁵N₃]dC tracing and LC-MS/MS analysis in CEM-YFP or CEM-CDA cells to gain additional mechanistic insight into the interplay between dC and CDA in modulating PNPi/dG-mediated toxicity (Figure 4C and Supplemental Figure 3E). Ectopic CDA expression resulted in the depletion of [¹⁵N₃]dC from the media (Supplemental Figure 3F). PNPi expanded the dGTP pool derived from [¹⁵N₅]dG salvage and depleted intracellular dCTP only in CEM-CDA cells and had no effect in the parental cells in the presence of dC (Figure 4C). This indicated that dC blocks the entry of dG into dGTP pools via dCK and thereby acts as an endogenous inhibitor of PNPi/dG-mediated lethality.

To investigate whether increasing CDA levels in vivo sensitizes SAMHD1-deficient cells to PNPi, we established CEM-YFP control and CEM-CDA bilateral subcutaneous tumors in NCG mice and initiated PNPi treatment once tumor volumes reached 100 mm³ (Figure 4D). PNPi treatment eradicated CEM-CDA tumors, while CEM-YFP tumors grew unchecked (Supplemental Figure 3G). LC-MS/MS-MRM analyses of tumor interstitial fluid dG and dC levels showed that PNPi increased dG levels while CDA expression significantly reduced dC levels (Figure 4E). Collectively, these findings indicate that elevated dC levels in mice relative to humans or humanized mice engender resistance to the synthetic lethality between PNP and SAMHD1 and suggest that humanized mice are a suitable model for the metabolic alterations induced by PNP deficiency during T cell development.

PNP inhibition in mice induces immune activation phenotypes.

In addition to T cell immunodeficiency, loss of PNP function associates with various autoimmune and inflammatory phenotypes in humans (5). Moreover, PNP-KO mice exhibit phenotypes indicative of hyperimmune activation, such as splenomegaly (8, 10). Consistently, pharmacological PNP inhibition in C57BL/6 mice for 2 weeks induced a significant increase in spleen size (Figure 5A). This effect occurred independently of dCK activity and was not accompanied by alterations in spleen cellular composition (Figure 5B).

To elucidate the mechanism responsible for this phenotype, we first determined whether acute PNP inhibition in C57BL/6 mice alters serum levels of cytokines associated with immune activation. A single administration of PNPi was sufficient to induce multiple cytokines and chemokines (Figure 5C). In particular, we noted a robust increase in IL-6, an inflammatory cytokine linked to autoimmunity and inflammation in mice and humans (31, 32). To identify the source(s) of IL-6 in PNPi-treated mice, we measured Il6 transcript levels across multiple tissues and found that the spleen exhibited the greatest fold increase in Il6 transcripts, followed by the lymph nodes and the thymus (Figure 5D).

To further investigate the effects of acute PNP inhibition, we performed single-cell RNA-Seq (scRNA-Seq) analysis of splenocytes isolated from vehicle- and PNPi-treated mice. Integrated uniform manifold approximation and projection (UMAP) of splenocytes from control and treated mice identified 23 unique clusters encompassing all major immune cell lineages (Figure 5E). The most profound transcriptional alterations induced by PNP inhibition were observed within an expanded B2 cluster, which corresponds to activated CD19⁺ B cells, and within the CSF1R⁺ macrophage cluster (Figure 5F and Supplemental Figure 4A). Ontological analysis of genes significantly altered by PNPi revealed enrichment of pattern recognition receptor (PRR) signaling–driven NF-κB–mediated cytokine responses and IFN regulatory factors/IFN response signatures across CD19⁺ B cell and CSF1R⁺ macrophage clusters (Figure 5G and Supplemental Figure 4B). PNPi additionally stimulated Myc and E2f transcription factor–regulated responses associated with proliferation in cluster B2 cells (Figure 5G). Furthermore, PNP inhibition stimulated NF-κB–regulated cytokine transcripts, including Tnf, Il10, and Il6 in B cells, alongside Tnf, Il1b, and IFN-response biomarker Cxcl10 in macrophages (Figure 5H). These findings indicate that, in addition to its positive role in T cell development, PNP functions in the periphery as a negative regulator of immune activation initiated by PRR signaling. These immune functions of PNP are likely mediated by different mechanisms involving distinct gene interactions.

PNP controls TLR7 at the level of guanosine ligand availability.

Recent studies of TLR7 and TLR8 activation mechanisms provide potential clues about how PNP may function as a negative regulator of PRR activation (33, 34). TLR7 activation requires binding of single-stranded RNA (ssRNA) containing a nonterminal uridine residue to “site 2” followed by binding of a guanine nucleoside to “site 1” (33). Dual ssRNA/guanine nucleoside binding initiates TLR7 dimerization, activation of downstream signaling, and transcriptional initiation of cytokine responses. We reasoned that PNP inactivation could initiate TLR7 signaling by increasing the levels of “site 1” ligands. Consistent with this model, acute inhibition of PNP in mice substantially increased guanine nucleoside levels in the spleen and lymph nodes, which correlated with the observed induction of IL-6 (Figure 6A).

To investigate a role for PNP as a negative regulator of TLR7, we utilized TLR7-expressing BMDMs stimulated with combinations of TLR7 site 1 (dG and rG) and site 2 (21-mer poly-uridine RNA complexed with DOTAP [polyU]) ligands with or without PNPi. Both dG and rG induced Il6 expression in PNPi- and polyU-treated BMDMs (Figure 6B). PNP inhibition was required for the ability of guanosine nucleosides to induce IL-6 expression in polyU-treated BMDMs (Figure 6B). In BMDMs, PNPi and guanosine nucleosides were also essential for the ability of various polyU concentrations to induce Il6 expression (Supplemental Figure 5A) and phosphorylation of downstream TLR7 signaling mediators, IKKβ, IκBα, and NF-κB (Supplemental Figure 5B). Inhibition of TAK1, IRAK4, or IKKβ, the kinases responsible for propagating TLR7 signaling, restricted Il6 induction upon PNPi+dG/rG+ssRNA treatment (Supplemental Figure 5C). While polyU induced Ifnb1 in expression in BMDMs in a dose-dependent manner, PNPi and guanine nucleosides significantly increased this response at all examined polyU concentrations (Supplemental Figure 5D). PNP inhibition also modulated ssRNA recognition in other TLR7-expressing immune cells. In FLT3-differentiated bone marrow–derived dendritic cells, PNPi+dG/rG significantly potentiated polyU-induced Il6 expression (Supplemental Figure 5E). In primary murine B cells, PNPi+dG/rG induced Il6 expression independently of polyU (Supplemental Figure 5E), indicating that these cells contain sufficient amounts of endogenous ssRNA to initiate TLR7 signaling in the presence of elevated levels of site 1 guanosine ligands.

Next, we used genetic approaches to unequivocally demonstrate that PNP inhibition triggers TLR7 activation. Chronic TLR7 signaling in mice is associated with reductions of B cell numbers in peripheral blood and expansion of myeloid cell populations (35). Consistently, PNPi treatment in mice restricted circulating B220⁺ B cells while expanding CD11b⁺ and CD11c⁺ myeloid populations (Figure 6C). Genetic inactivation of MyD88, an essential TLR7 signaling adaptor, prevented these alterations. Additionally, we observed that (a) increases in serum IL-6 and TNF-α by PNPi were abrogated in Tlr7-KO mice (Figure 6D), (b) PNPi+dG/rG did not induce Il6 expression in polyU-transfected Tlr7-KO BMDMs or B cells (Figure 6E), and (c) PNPi+dG/rG induced phosphorylation of downstream TLR7 effectors IKKα/β and NF-κB p65 alongside polyU only in TLR7-expressing BMDMs (Figure 6F).

Collectively, these findings suggest a model in which PNP regulates TLR7 activation by restricting the availability of site 1 guanosine ligands in endolysosomes (Figure 6G). PNP inactivation stabilizes guanosine nucleosides, thereby facilitating their transit to endolysosomes.

Unique transcriptional responses to PNPi-stabilized guanosine nucleosides and imidazoquinolines.

Imiquimod (R837) and resiquimod (R848) are synthetic imidazoquinoline TLR7 site 1 ligands that bind independently of ssRNA (33). To determine if transcriptional alterations triggered by PNPi with guanosine nucleosides overlap or diverge from those elicited by synthetic TLR7 ligands, we performed RNA-Seq analysis of BMDMs, following acute stimulation with PNPi+dG/rG, polyU, PNPi+dG/rG+polyU, or R848. Principal component analysis of significantly altered transcripts (>10 fold change, <0.01% FDR) revealed a significant separation among these groups (Figure 7A and Supplemental Data File 1). K-means clustering and ontology analysis of induced transcripts reveled that both TLR7 activation modalities elicited NF-κB–associated responses, but a type I IFN response was induced exclusively by ssRNA and PNPi-stabilized guanosine ligands (Figure 7, B and C). A titration of R848 confirmed that, unlike endogenous coligands, this synthetic TLR7 agonist does not induce type I IFN expression in BMDMs (Figure 7D). These findings indicate that PNP tunes TLR7-driven transcriptional responses to ssRNA and reveal that natural or synthetic TLR7 ligands elicit distinct transcriptional profiles in BMDMs.

PNP is a metabolic immune checkpoint that controls germinal center formation and autoimmunity.

Our studies indicated that PNP inhibition triggers TLR7 activation in B cells both in vivo and in cell culture (Figure 5H) and Supplemental Figure 5E). B cell–intrinsic TLR7 signaling and IL-6 production promote the development of spontaneous germinal centers (GCs) in secondary lymphoid tissues, which are associated with autoinflammation and autoimmunity (31, 36). To investigate a role for PNP in regulating the GC reaction, we evaluated C57BL/6, BALB/c, and NOD mice treated with PNPi daily for 2 weeks. Splenomegaly and increased spleen cellularity were observed in all treated mice (Supplemental Figure 6A). PNPi stimulated the expansion of the CD19⁺GL7⁺CD95⁺PNA⁺ GC B cells in the spleen and inguinal lymph nodes (Figure 8A and Supplemental Figure 6, B and C). PNPi also significantly increased the abundance of GC-participating TCRβ⁺CD4⁺CD62L^–CXCR5⁺PD1⁺ T follicular helper cells. Moreover, the expression of the GC-promoting cytokine IL-6 increased in splenic B cells from BALB/c mice following PNPi treatment (Supplemental Figure 6D).

Unlike the T cell developmental defects induced by PNP inhibition (Figure 1B), the effects of PNPi treatment on secondary lymphoid organs and GC formation were independent of dCK activity, as dCKi coadministration did not effect PNPi-induced alterations in spleen weight, GC B cell, or T follicular helper cell expansion (Figure 8B). Moreover, dCK inhibition did not affect Il6 upregulation by PNPi+dG/rG in B cells in vitro (Figure 8C). Together, these results indicate that (a) PNP inhibition in mice triggers the formation of spontaneous GCs in the absence of foreign antigen and (b) dCK/SAMHD1- and TLR7-dependent effects of PNP inactivation can be functionally uncoupled by dCK inhibition, which restricts PNP inactivation-induced dGTP pool expansion while retaining TLR7-driven cytokine responses.

Constitutive TLR7 signaling is associated with the development of systemic lupus erythematosus–like autoimmune phenotypes in mice and humans (36). Additionally, various autoimmune pathologies are characteristic of PNP deficiency in humans (6). To determine if PNP inactivation in mice engenders similar autoimmunity-associated phenotypes we used MRL-LPR mice, which are prone to the development of systemic autoimmunity and B cell–dependent nephritis (37). We treated MRL-LPR mice with PNPi for 35 days and used ultrasound imaging to monitor increases in spleen size as an indicator of disease progression. PNPi-induced splenomegaly was apparent as early as day 22 and was sustained until the experimental endpoint (Figure 8D and Supplemental Figure 6E). This finding was corroborated by measurements of spleen mass (Figure 8E). PNPi treatment also increased serum cytokine levels and accelerated the onset of other manifestations of autoimmunity in the MRL-LPR model, including lymphadenopathy, proteinuria, and glomerulonephritis (Figure 8, E–H, and Supplemental Figure 6, F and G). Activation of splenic B cells by PNPi was evidenced by increased PNA binding in CD19⁺ cells (Figure 8I).

These findings indicate that PNP functions as a metabolic immune checkpoint that restricts inappropriate TLR7 activation, controls the formation of spontaneous GCs, and participates in the maintenance of B cell tolerance.

Limitations of the study.

Our findings underscore the importance of considering metabolic and immune differences between mice and humans in the design of translational research studies. Interspecies differences can be addressed through the development of mouse models that possess human nucleoside levels without the inherent limitations of the BLT mice, which are not readily amenable for genetic studies of T cell development. Our data offer guidance on future directions; one possibility is the development of mice that possess humanized systemic CDA activity. Regarding the role of PNP as a negative regulator of TLR7 signaling, the lack of functional TLR8 expression in mice (48) may confound the autoimmune and inflammatory phenotypes caused by PNP inhibition. Future studies in humanized TLR8-expressing mice (49) will be required to address this limitation.

Cell culture

Cell culture was performed as previously described (50). All cancer cell cultures were maintained between passages 3 and 20 and maintained in antibiotic-free media plus 10% FBS at 37°C in 5% CO₂. All leukemia and lymphoma models were maintained in RPMI plus 10% FBS. Primary T cell cultures were maintained in RPMI with 10% FBS and 30 U/mL recombinant IL-2. Primary B cell cultures were maintained in RPMI with 10% FBS and 1× sodium pyruvate and 0.1% β-mercaptoethanol. Solid tumor cultures were maintained in DMEM with 10% FBS. All cell culture experiments were performed using media supplemented with 10% dialyzed FBS. Cell cultures were routinely monitored for mycoplasma contamination using the PCR-based Venor Mycoplasma kit. Cell lines were acquired either from a commercial vendor (ATCC and DSMZ) or from collaborators. Cell line identity was independently authenticated by PCR (Laragen). Methods for culture and analysis of primary cells are included in Supplemental Methods.

Drugs

Drugs for in vitro studies were prepared as previously described (50). Drug stocks were prepared in neat DMSO or H₂O, stored at –80°C and diluted fresh in cell culture media for experiments. 21-mer polyU ssRNA was complexed with DOTAP before treatment.

Animal studies

For treatment studies, ulodesine was formulated in 100% sterile distilled water and administered as indicated in a volume of 100 μL/20 g mouse; (R)-DI-87 was formulated in 40% captisol and administered at 75 mg/kg p.o. b.i.d. or 100 mg/kg p.o. q.d. in a volume of 100 μL/20 g mouse. For ad libitum administration of PNPi, ulodesine was formulated in drinking water at 0.13 mg/mL, which was intended to deliver a daily dose of 20 mg/kg. Urine protein was measured using the BCA assay (Supplemental Table 1).

For subcutaneous tumor models, 6- to 8-week-old male NOD-Prkdc^em26Cd52;Il2rg^em26Cd22;NjuCrl coisogenic immunodeficient (NCG) mice were injected subcutaneously on the flanks with 1 × 10⁶ CEM cells resuspended 1:1 in PBS/Matrigel (50 μL/50 μL). Subcutaneous tumor volumes were calculated by micro-computed tomography (μCT) analysis using a G8 PET/CT scanner. All tumor volume measurements were performed by trained technicians blinded to experimental conditions.

NSG and TKO BLT-humanized mice were derived by the UCLA CFAR Humanized Mouse Core Facility. The generation of these mice has been previously described (51). For these mice, human fetal liver and autologous thymus fragments were coimplanted under the renal capsule of NSG or TKO mice and human HSCs were engrafted to the bone marrow by intravenous injection. Human lymphocyte reconstitution was evaluated by flow cytometry analysis of percentage of human CD45-reactive peripheral blood cells and confirmed to be more than 30% for all mice before enrollment for treatment studies. Humanized mouse treatment studies were initiated 4 months after tissue engraftment.

Ultrasound imaging was performed using the Vevo 2100 Ultrasound from Visual Sonics equipped with an ultrahigh frequency linear probe. Prior to imaging, animals underwent hair removal with clippers and depilatory cream. 3D scans were obtained and exported for analysis using the companion software, Vevo Lab version 5.5.1. Spleen volume measurements were performed using the multislice setting and a step size factor of 1 and were reported in mm³.

ATO model

RNA-Seq

scRNA-Seq

Methods for scRNA-Seq data acquisition and analysis are included in Supplemental Methods.

Immunoblot analysis

PBS-washed cell pellets obtained by trypsinization of cells and centrifugation at 450g for 5 minutes were resuspended in cold RIPA buffer supplemented with protease and phosphatase inhibitor cocktails. Sample protein content was determined using the BCA assay. Samples were normalized by RIPA and 4× Laemmli loading dye dilution, resolved on 4%–12% Bis-Tris gels, and electrotransferred to nitrocellulose membranes. After blocking with 5% nonfat milk in TBS 0.1% Tween-20 (TBS-T) for 30 minutes at room temperature, membranes were incubated overnight in primary antibodies diluted 1:1000 in 5% BSA in TBS-T at 4°C. Membranes were washed with TBS-T and incubated with HRP-linked secondary antibodies prepared at a 1:2500 dilution in 5% nonfat dry milk/TBS-T for 1 hour at room temperature. HRP was activated by incubating membranes with a mixture of SuperSignal Pico and SuperSignal Femto ECL reagents (33:1 ratio). Exposure of autoradiography film was used for detection. Antibodies are reported in Supplemental Table 1. Catalog information for all reagents is included as Supplemental Table 1. See complete unedited blots in the supplemental material.

RT-PCR

RT-PCR was performed as previously described (50). Total RNA was isolated from cell cultures using the NucleoSpin RNA kit (Supplemental Table 1) and quantified by Nanodrop. Total RNA was isolated from 30 to 80 mg of snap frozen tissues by homogenization in 500 μL extraction buffer in Omni Hard Tissue homogenization vials using an Omni Bead Ruptor Elite (8 cycles of 15 seconds on, 30 seconds off, speed 8) chilled to 4°C, and isolated using the NucleoSpin RNA kit. Reverse transcription was performed using the High Capacity cDNA Reverse Transcription kit (Supplemental Table 1). Quantitative PCR was performed using EvaGreen qPCR Master Mix on the QuantStudio3 system (Supplemental Table 1). RNA expression values were normalized to housekeeping gene (ACTIN, Actb) expression, calculated using the ^ΔΔCt method, and reported as relative expression to vehicle control-treated samples. Primer sequences are indicated in Supplemental Table 1.

Cell proliferation analysis

For Cell Titer Glo (Supplemental Table 1) analysis, cells were seeded at 1 × 10³ cells per well in 50 μL media with 10% dialyzed FBS in white 384-well plates and treated as described. After 72 hours, 50 μL Cell Titer Glo reagent (diluted 1:5 in PBS) was added to each well and luminescence was measured using a Synergy H1 reader (Supplemental Table 1).

Generation of isogenic cell line models

Luminex analysis

Blood was collected in 1.5 mL Eppendorf tubes by the retro-orbital technique using heparin-coated capillary tubes and incubated at room temperature for 30 minutes. Samples were centrifuged at 2000g for 15 minutes at 4°C, and serum was transferred to a new tube for 32-plex Luminex analysis performed by the UCLA Immune Assessment Core.

Flow cytometry

Methods for flow cytometry data acquisition and analysis are included in Supplemental Methods.

Mass spectrometry

Methods for mass spectrometry data acquisition and analysis are included in Supplemental Methods.

Statistics

Statistical analysis was performed as previously described (50). Data are presented as mean ± SD, with the number of biological replicates indicated. Comparisons of 2 groups were evaluated using either unpaired 2-tailed Student’s t test or 2-tailed Mann-Whitney test, and P values of less than 0.05 were considered significant. Comparisons of more than 2 groups were evaluated using either 1-way ANOVA, followed by Bonferroni’s multiple comparisons test or Kruskal-Wallis test with Dunn’s multiple comparisons correction, and P values less than 0.05/m, where m is the total number of possible comparisons, were considered significant.

Study approval

All animal studies were approved by the UCLA Animal Research Committee.

Neonatal umbilical cord blood was obtained under UCLA IRB-approved protocols or exemptions.

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