Abstract
Most enveloped viruses rely on furin for maturation of their surface glycoprotein. In contrast, mammarenaviruses process their glycoprotein precursor (GPC) using host site-1 protease (S1P), yet the biological implications of this unique reliance on S1P remain unclear. Here, we characterized a furin-dependent recombinant form (rCl13-RRRR) of the persistent clone 13 variant of LCMV (rCl13). Although rCl13-RRRR exhibited fitness comparable to rCl13 in cultured cells, it was highly attenuated in vivo and failed to establish persistence in immunocompetent mice. Clearance of rCl13-RRRR required interferon and CD8+ T cells, and immunization with rCl13-RRRR conferred protective immunity against a subsequent lethal LCMV challenge. Our results demonstrate that S1P-mediated processing of GPC is a key determinant of mammarenavirus fitness and immune evasion in vivo and highlight S1P as a promising and druggable target for host-directed antiviral strategies against human pathogenic mammarenaviruses.
Similar content being viewed by others
Introduction
Mammarenaviruses include important human pathogens such as Lassa virus (LASV), and Junín virus (JUNV). The Old-World (OW) mammarenavirus LASV is endemic in West Africa, where it is estimated to infect several hundred thousand people annually1,2,3,4,5,6. About 20% of LASV human infections can result in Lassa fever (LF) disease with symptoms ranging from mild to hemorrhagic fever disease associated with high morbidity and mortality1,2,7. Likewise, the New World (NW) mammarenavirus JUNV is the causative agent of Argentine hemorrhagic fever (AHF) with case fatality rates exceeding 15%, and several other NW mammarenaviruses cause hemorrhagic fever diseases throughout South America8,9. In addition, evidence indicates that the globally distributed mammarenavirus lymphocytic choriomeningitis virus (LCMV) is an underrecognized human pathogen of clinical significance in congenital infections10, and it poses a serious threat to immunocompromised patients11,12,13,14,15, but can also cause severe disease in immunocompetent people16. Despite their impact on human health, there are no FDA-approved mammarenavirus vaccines or antivirals, and current therapeutic options to treat mammarenavirus human infections are limited to the off-label use of ribavirin, whose efficacy remains controversial17,18,19,20,21. The live-attenuated Candid1 strain of JUNV has been shown to be an effective vaccine against AHF22, but outside Argentina, Candid1 has only an investigational new drug status. Thus, the development of effective antivirals and vaccines against mammarenaviruses is of high priority and will be facilitated by a better understanding of the molecular and cellular processes that modulate mammarenavirus–host interactions.
Mammarenaviruses are enveloped viruses with a bi-segmented negative-sense RNA genome. Each genome segment uses an ambisense coding strategy to express two viral proteins. The small (S) segment encodes for the glycoprotein precursor (GPC) and the nucleoprotein (NP), whereas the large (L) segment encodes for the matrix protein (Z) and the viral polymerase (L)23,24. NP encapsidates the viral genome RNA to form a nucleocapsid structure to which the L polymerase associates to form the viral ribonucleoprotein complex (vRNP) responsible for directing replication and transcription of the viral genome25,26. The matrix Z protein is a main driver of viral budding27,28,29,30. GPC is co- and post-translationally processed by host signal peptidase and site-1 protease (S1P) aka subtilisin/kexin-isozyme-1 (SKI-1), respectively31,32,33,34,35,36,37, to generate a stable signal peptide (SSP), and the surface GP1 and membrane-anchored GP2. The mature GP complex, made of SSP/GP1/GP2 heterotrimers, forms the spikes that decorate the virion surface and mediate virus cell entry via receptor-mediated endocytosis. GP1 binds to cellular receptors, whereas GP2 mediates a pH-dependent fusion event between viral and cell membranes within the acidic environment of the late endosome33,35,37,38. Surface GPs of many enveloped viruses are activated via processing by furin and furin-like proprotein convertases39,40,41,42,43,44 or trypsin45. However, all tested mammarenaviruses and some other bunyaviruses use S1P for processing of their GPCs33,34,37,46,47, but the reasons for this mammarenavirus strict dependence on S1P for GPC processing remain unknown. Hence, elucidating the biological implications of the S1P-mediated processing of the mammarenavirus GPC could provide scientific significance.
We have documented the generation and in vitro characterization of a recombinant form of the Armstrong strain of LCMV expressing a GPC with a furin, instead of S1P, recognition cleavage site48. In the present work, we have investigated the biological properties in vivo of a recombinant form of the immunosuppressive variant clone 13 (Cl13) of the Armstrong strain of LCMV49,50,51,52,53 expressing a GPC that is processed by furin instead of S1P (rCl13-RRRR). We found that rCl13-RRRR had a Cl13-like fitness in vitro but was greatly attenuated in mouse models of LCMV infection. Thus, rCl13-RRRR failed to establish persistence in immunocompetent B6 mice49,50,51,52,54,55,56,57 or to cause lethal disease in FVB mice58,59. However, priming immunocompetent mice with rCl13-RRRR induced protective immunity against a subsequent Cl13 lethal challenge. These findings support a unique role of S1P-mediated GPC processing in the mammarenavirus–host interactions that it is unlikely to be replaced by other host proprotein convertases and further solidified S1P as a druggable target for the development of host-directed antivirals to combat infections by human pathogenic mammarenaviruses.
Materials and methods
Cells
Vero E6 (ATCC CRL-1586), A549 (ATCC CCL-185), HEK 293 T (ATCC CRL-3216), and BHK-21 (ATCC CCL-10) cells were maintained in Dulbecco’s Modified Eagle Medium (DMEM) (Thermo Fisher Scientific, Vacaville, CA, USA) supplemented with 10% fetal bovine albumin (FBS), 2 mM L-glutamine, 100 μg/mL streptomycin, and 100 U/mL penicillin. BHK-21 medium also contained 5% tryptose phosphate broth (#18050039, Thermo Fisher Scientific). HAP1 WT, S1P-KO and S2P-KO cells were generously provided by Dr. Sandra Pohl at University Medical Center Hamburg-Eppendorf, Germany. HAP1 WT cells were maintained in Iscove’s Modified Dulbecco’s Medium (IMDM) (Gibco) supplemented with 10% FBS, 2 mM L-glutamine, 100 μg/mL streptomycin, and 100 U/mL penicillin. HAP1-S1P-KO and S2P-KO cell lines were maintained in the same medium with extra lipid supplements (1 mM sodium mevalonate, 20 μM sodium oleate, and 5 μg/ml cholesterol (L4646, Sigma-Aldrich)). LoVo (ATCC CCL-229) and LoVo-furin cell lines were maintained in Ham’s F-12K (Kaighn’s) Medium (Gibco) supplemented with 10% FBS, 2 mM L-glutamine, 100 μg/mL streptomycin, and 100 U/mL penicillin. LoVo-furin cells were generated from LoVo cells with a stable transfection of a plasmid expressing furin.
Viruses
Recombinant LCMV Armstrong strain (rArm) and Cl13 variant (rCl13), as well as recombinant LCMV Cl13 where the furin recognition cleavage site substituted for the S1P recognition cleavage site (rCl13-RRRR) were rescued in BHK-21 cells via reverse genetics as previously described60. Mutations were introduced using the NEB Q5 Site-Directed Mutagenesis kit per manufacturer protocol. The primer sequences used are forward: TAGGAGACGTCGCGGCACATTCACCTG; reverse: GTGAGGAACTTAGTTTTTTC. Stocks of the rescued viruses were prepared by infecting BHK-21 cells at MOI = 0.01, and their genetic identity confirmed by sequencing the viral genome RNA. All studies were approved by the Institutional Biosafety Committee (IBC) and the Institutional Animal Care and Use Committee (IACUC).
Antibodies and compounds
PF-429242 dihydrochloride (SML0667) was purchased from Sigma-Aldrich and dissolved in DMSO to make a 10 mM stock solution. A 1 mM working stock was made by diluting 1:10 in Opti-MEM (Gibco). BOS-318 (HY-147140) was purchased from MedChemExpress and dissolved in DMSO to make a 2 mM stock solution. Ribavirin (R9644) was purchased from Sigma-Aldrich and stored in Opti-MEM at 10 mM. Sodium citrate buffer (0.1 M, pH 5.0, sterile) was obtained from bioWorld (#40121003-1, Dublin, OH, USA). The VL4 rat monoclonal antibody against LCMV NP was obtained from Bio X Cell Therapeutics (New Haven, CT, USA). Conjugation of the antibody with AF488 was done with Invitrogen Antibody Labeling Kit (A10235) per the manufacturer's protocol. Human monoclonal antibodies against GPC (37.2D and 12.1F) was made by cloning the published immunoglobulin sequences61 into expression vectors to transfect Expi293F cells included in the Expi293TM Expression System Kit (A14635, Gibco) per manufacturer protocol. 200 mL culture were transfected with a total of 100 µg plasmids expressing light and heavy chains as well as 10 µg of pADVAntage (E1711, Promega). Cell culture media was collected at 6 days post transfection, and antibodies were purified using protein G Sepharose beads (#101243, Invitrogen). PE-Cy7 B220 (RA3-6B2, #103222), PE CD3 (17A2, #100206), BV785 CD8α (53-6.7, #100750), Pacific Blue Siglec-H (551, #129609), BV711 NK1.1 (PK136, #108745), APC-Cy7 CD11b (M1/70, #101226), BV421 CD11c (N418, #117343), PE-fire810 Ly6G (1A8, #127673), PerCP-Cy5.5 I-A/I-E (M5/114.15.2, #107626), APC Ly6C (HK1.4, #128016), PE-Cy7 CD44 (IM7, #103029), PE IFN-γ (XMG1.2, # 505808), and PerCP-Cy5.5 TNFα (MP6-XT22, #506322) rat anti-mouse antibodies for flow cytometry were purchased from BioLegend. BUV395 CD4 antibody (GK 1.5, #563790) was purchased from BD Biosciences. AF660 F4/80 antibody (BM8, #606480180) was purchased from Invitrogen. AF568 CD169 antibody (EPR27102-11, #ab316878) was purchased from Abcam. Anti-CD8 antibody for CD8 depletion (YTS 169.4, #BE0117), anti-CD4 antibody for CD4 depletion (GK 1.5, #BE0003-1), and anti-IFNαR antibody for IFNαR blockade (MAR1-5A3, #BE0241) were purchased from Bio X Cell. Anti-CD169 antibody (3D6.112, #MA5-16508) for frozen section staining were purchased from Thermo Fisher Scientific.
Focus-forming assay
Vero E6 cells were seeded at 2 × 104 cells/well in 96-well plates. The next day, serially diluted viruses were added (100 µL/well) to the cell monolayers. After 90-min adsorption, the virus inoculum was aspirated and fresh media (100 µL/well) containing 2% FBS and 0.5% methyl cellulose were added to the cells. At 24 h post infection, cells were fixed with 4% paraformaldehyde (PFA) for 15 min and stained with rat mAb VL4 against LCMV NP (Bio X Cell) in PBS with 0.3% Triton X-100 and 3% bovine serum albumin (BSA) overnight at 4 °C. Plates were washed twice with PBS and stained with anti-rat IgG HRP-conjugated antibody and subsequently with KPL TrueBlue peroxidase substrate (#5510-0030, SeraCare, Milford, MA, USA). Foci were quantified using CTL Immunospot Analyzer (Cleveland, OH, USA). The limit of detection (LOD) was set at 2 logFFU/mL. Samples with no observable foci were assigned an arbitrary value of 1.5 logFFU/mL below the LOD.
Western blot
Whole cell lysates were collected in lysis buffer (50 mM Tris-HCl, pH 7.5, 1 mM EDTA, 0.5% NP-40, and 150 mM NaCl) with HaltTM protease and phosphatase inhibitor cocktail (Thermo Fisher Scientific) and centrifuged to collect supernatant. Samples were denatured for 5 min at 95 °C, then 16 µg per sample were loaded onto an SDS-PAGE stain-free gel (Bio-Rad). Separation was done by electrophoresis at 200 V for 30 min. Transfer of protein to a low-fluorescence PVDF membrane (Bio-Rad) was done at 1.3 A, 25 V for 12 min. The membrane was blotted with primary antibody overnight at 4 °C, then with secondary antibody for 1 h at room temperature. Bands were visualized with SuperSignal™ West Pico PLUS Chemiluminescent Substrate (#34580, Thermo Fisher Scientific).
Virus growth kinetics
Cells were seeded at 1 × 105 cells/well (A549) or 2 × 105 cells/well (HAP1 and LoVo) in 24-well plates. The next day, cells were infected (0.25 mL/well; MOI = 0.01). After 90 min adsorption at 37 °C, the virus inoculum was then aspirated and replaced with fresh growth media. Cell culture supernatant (CCS) was collected at 24, 48, 72, and 96 hpi, and virus titers were determined by focus-forming assay (FFA). For HAP1-S1P-KO and S2P-KO cells, media was supplemented with 1 mM sodium mevalonate, 20 μM sodium oleate, and 5 μg/ml cholesterol.
Dose-response assay
A549 cells were seeded at 2 × 104 cells/well in 96-well plates. The next day, cells were infected (100 µL/well; MOI = 0.01) for 90 min. Virus inoculum was then aspirated and replaced with fresh growth media (100 µL/well) containing serially diluted compounds. Media containing DMSO or 100 µM ribavirin (RBV) were used as controls. The medium of samples treated with the S1P inhibitor PF-429242 was supplemented with lipids (1 mM sodium mevalonate, 20 μM sodium oleate, and 5 μg/ml cholesterol). Cells were fixed with 4% PFA at 72 hpi and stained with rat mAb VL4 against LCMV NP (Bio X Cell), then with anti-rat AF488-conjugated antibody. CellTiter 96 AQueous One Solution (G3580, Promega) and DAPI staining were used to assess the effect of PF-429242 and BSO-318 on cell viability. CellTiter assay was done following the manufacturer’s protocol. Briefly, 10 µL of the assay reagent was added to 100 µL of the media already on the cells at the 72 hpi. Then the plates were allowed to incubate at 37 °C for 20 min before reading the absorption at 490 nm minus the background at 700 nm. Plates were read on a Biotek Cytation 5 plate reader (Agilent Technologies, Santa Clara, CA, USA). Readouts were normalized to vehicle-treated wells, which were assigned a value of 100%. Six replicates per dilution per compound were used to plot the dose-response curve. Effective concentration 50 (EC50) and cytotoxic concentration 50 (CC50) were calculated on GraphPad Prism using a nonlinear regression model with equation log(inhibitor) vs. response – Variable slope (four parameters). The selective index (SI) was calculated using the formula SI = CC50 / EC50.
Fusion assay
HEK293T cells were seeded at 2.5 × 105 cells/well in six-well plates. The next day, each well was transfected with 1 µg pCAGGS plasmids expressing WT or mutant GPCs or an empty plasmid as a negative control, together with 50 ng pCAGGS plasmid expressing GFP. Compounds were added at the indicated time and concentrations. At 48 h post transfection, media was transferred to a clean plate and the cell monolayers were treated with citrate buffer (pH 5.0, 50 mM sodium citrate, 5 mM KCl, 2 mM CaCl2, and 90 mM NaCl) or PBS (pH 7.0) for 15 min at 37°C. After treatment, the original media was added back to the matching wells, and the plate was returned to the incubator. Fusion activity was monitored every 10 min using a fluorescent microscope. Once fusion was observed in a specific well, all other wells were given 5 extra minutes to initiate fusion. After 10 min of the first observed fusion activity, all wells were fixed with 4% PFA at the same time, then permeabilized and stained with human mAb to GPC, followed by anti-human AF568-conjugated antibody. Images were taken with a Zeiss LSM 780 Confocal laser scanning microscope at 10x magnification.
Mice and in vivo infection
Husbandry and handling of mice conformed to guidelines set forth by the National Institute of Health Guide for the Care and Use of Laboratory Animals and Department of Animal Resources at The Scripps Research Institute (TSRI). Mice experiments were approved under the TSRI Institutional Animal Care and Use Committee (IACUC) protocol #09-0098 and #09-0137. C57Bl6/J, IFNαR-KO, IL27αR-KO and FVB/N mice were obtained from the rodent breeding colony at Scripps Research. Analgesics and anesthetics were used as determined by the veterinarian staff. Anesthetic: isoflurane (3% inhaled with precision vaporizer). Analgesics (based on veterinarian assessment): buprenorphine (0.1 mg/kg) or flunixin (2.5 mg/kg). For intravenous (IV) infections, mice were injected with 2 × 106 FFU retro-orbitally unless otherwise stated. For intracranial (IC) infection, viruses were first diluted in sterile PBS to the appropriate titers depending on the dosage (1000 FFU unless otherwise stated), then the viruses were injected using a special stepper tool, which held the syringe and injected only 20 µL each time. At experimental endpoints, Mice were euthanized by an overdose of inhalant anesthetic (isoflurane), followed by cervical dislocation, a method consistent with the recommendations of the most recent (2020) AVMA Guidelines for the euthanasia of animals. Bleeding was done retro-orbitally using heparinized capillary tubes. Blood was collected in 1.5 mL microcentrifuge tubes and spun at 3000 ×g for 5 min to isolate the serum. Tissues were harvested from euthanized mice and homogenized in 1 mL of DMEM with 2% FBS, 2 mM L-glutamine, 100 μg/mL streptomycin, and 100 U/mL penicillin. Homogenates were clarified by centrifugation at 10,000×g for 10 min at 4°C, and 800 µL of clarified homogenates were collected from each sample for the titration of infectious virus.
Flow cytometry
Spleens were harvested from euthanized mice and mechanically dissociated through a 70-µm filter. The single-cell suspensions were then erythrocyte-depleted in ammonium-chloride-potassium (ACK) lysing buffer (A1049201, Thermo Fisher Scientific) for 3 min at room temperature. The remaining cells were resuspended in FACS buffer (PBS with 2% FBS and 1 mM EDTA) and counted. To quantify NP+ splenocytes, cells were stained with antibodies against indicated surface markers on ice for 1 hour, fixed, permeabilized, and then intracellularly stained with AF488-conjugated rat mAb VL4 against LCMV NP.
For the T cell stimulation assay, as previously described62, splenocytes were incubated with LCMV GP33-41 peptides for 1 h at 37 °C. Then Brefeldin A (1 mg/mL) was added to stop Golgi export. After 5 h of incubation at 37 °C, cells were stained with antibodies against B220, CD8α, CD4, and CD44 on ice for 1 h, fixed, permeabilized, and then intracellularly stained with fluorescently conjugated antibodies against IFNγ and TNFα. Flow cytometry was performed on Cytek Aurora and analyzed using FlowJo software.
Spleen sectioning
Spleens were harvested from euthanized mice and fixed in 4% PFA at 4 °C for 12 h, then dehydrated in 30% sucrose solution at 4 °C, flash frozen in OCT at −80 °C. Embedded tissues were cut into 6–8 µm sections and stored at −80 °C. Before staining, sections were baked to 56 °C for 30 min, then washed with water to remove OCT. Slides were permeabilized with 0.5% Triton X-100 in PBS for 5 min, then blocked with 5% BSA + 0.1% Triton X-100 in PBS at room temperature for 1 h. Staining was done with anti-CD169 and anti-GPC antibodies at 4 °C overnight, then fluorescently conjugated secondary antibodies for 1 h at room temperature in 1% BSA + 0.1% Triton X-100 in PBS. Nuclear staining was done with DAPI for 15 min. Slides were then mounted with Fluoromount-G and imaged using a Zeiss LSM 780 Confocal laser scanning microscope.
IFNαR blockade, CD4+ and CD8+ T cells depletion
IFNαR blockade was done by intraperitoneal injection of 2 mg of anti-IFNαR antibody (MAR1-5A3) on day -1 prior to infection. For CD4+ T cell depletion, mice were intraperitoneally treated with 300 µg of anti-CD4 antibody (GK 1.5) 1 day prior to infection. For CD8+ T cell depletion, mice were intraperitoneally treated with 300 µg of anti-CD8 antibody (YTS 169.4) or rat IgG2b isotype control 2 days prior to and on the same day of infection.
Results
Characterization of a recombinant Cl13 (rCl13-RRRR) with furin-mediated GPC processing
To test if Cl13 could utilize any other host protease instead of S1P for its GPC processing, we rescued a recombinant Cl13 that had a furin cleavage site (262RRRR265) in place of its natural S1P cleavage site (262RRLA265) in the GPC gene. The rescue was done employing the described reverse genetics procedures48,60. Briefly, we transfected BHK-21 cells with mouse pol-I driven expression plasmids for the L and a mutant (GPC: RRLA to RRRR) S genome RNAs, together with two pCAGGS plasmids expressing the minimal viral trans-acting factors L and NP required for replication and transcription of the viral genome. Infectious particles were detected in the cell culture supernatant (CCS) at 4 days post transfection. We termed this newly generated virus rCl13-RRRR. We also rescued a wild type (WT) clone 13 (rCl13) and a rCl13 containing the S1P cleavage site from LASV (262RRLL265) termed rCl13-RRLL.
Results from multi-step growth kinetics in HAP1 cells revealed that rCl13-RRRR did not have fitness disadvantage compared to rCl13 (Fig. 1a). Growth of both rCl13 and rCl13-RRLL, but not rCl13-RRRR, was inhibited in CRISPR-generated S1P-KO HAP1 cells63,64 (Fig. 1b). Importantly, rCl13 growth was not affected in S2P-KO HAP1 cells demonstrating that processing of LCMV GPC is S1P-specific (Fig. 1c). As predicted, rCl13-RRRR growth also was not affected in SP2-KO HAP1 cells (Fig. 1c). To biochemically confirm that GPC processing of rCl13-RRRR was independent of S1P, we analyzed the cells lysates from WT, S1P-KO and S2P-KO HAP1 cells infected with rCl13, rCl13-RRRR or rCl13-RRLL by western blot (Fig. 1d). GPCs from all these three viruses were processed in WT and S2P-KO HAP1 cells, whereas only the GPC from rCl13-RRRR was processed in S1P-KO HAP1 cells. To further confirm the S1P or furin-dependent processing of the different viral GPCs, we compared their multi-step growth kinetics in furin-deficient LoVo65,66,67 and furin-reconstituted LoVo-furin cells (Fig. 1e). As predicted, rCl13 and rCl13-RRLL exhibited similar growth kinetics in LoVo and LoVo-furin cells, where rCl13-RRRR growth was greatly restricted in LoVo cell. To further confirm the expected phenotype of rCl13-RRRR, we tested its susceptibility to the furin (BOS-318) and S1P (PF-429242) inhibitors. As predicted, rCl13-RRRR, but not rCl13, was inhibited by BOS-318 in a dose-dependent manner (EC50 values of >2 µM and 0.161 µM for rCl13 and rCl13-RRRR, respectively) (Fig. 1f), while rCl13, but not rCl13-RRRR, was inhibited by PF-429242 in a dose-dependent manner (EC50 values of 2.46 and 28 µM for rCl13 and rCl13-RRRR, respectively) (Fig. 1g). The effect of BOS-318 and PF-429242 on cell viability was assessed by DAPI staining and CellTiter 96 AQueous One Solution. We observed a very good correlation between cell viability results obtained from both methods. PF-429242 and BOS-318 had CC50 values of >40 µM and >2 µM, respectively, which were the highest concentrations tested. Based on their EC50 and CC50 values, PF-429242 had SI values > 16.26 and >1.43 for rCl13 and rCl13-RRRR, respectively, whereas BOS-318 had SI values of >1 and >12.42 for rCl13 and rCl13-RRRR, respectively. The actual SI value of BOS-318 for rCl13-RRRR is expected to be much higher than the estimated >12.42, as no effects on cell viability was observed at the highest tested concentration (2 µM). We observed decreased NP expression levels in rCl13-RRRR-infected cells treated with the higher doses of PF-429242, which likely reflected compound-associated cytotoxicity. Likewise, in multi-step growth kinetic assays, PF-429242 inhibited production of rCl13 but not rCl13-RRRR infectious progeny (Fig. 1h).
a–c Multi-step growth kinetics of rCl13, rCl13-RRRR, and rCl13-RRLL in WT (a), S1P-KO (b), and SP2-KO (c) HAP1 cells. Cells were seeded at 3 × 105 cells/well in a 24-well plate and 24 h later infected (MOI = 0.01) with the indicated virus. Cell culture supernatants (CCSs) were collected at indicated hours post infection (hpi), and infectious virus titers were determined by focus-forming assay (FFA). d Processing of GPC in WT, S1P-KO, and SP2-KO HAP1 cells infected (MOI = 1) with rCl13, rCl13-RRRR, or rCl13-RRLL. At 24 hpi, cell lysates were prepared and analyzed by western blotting using the mouse monoclonal G204 to GPC. e Multi-step growth kinetics of rCl13, rCl13-RRRR, and rCl13-RRLL in furin-deficient LoVo cells and LoVo-furin cells with reconstituted furin expression. Cells were seeded at 3 × 105 cells/well in a 24-well plate and 24 h later infected (MOI = 0.01) with the indicated virus. CCSs were collected at the indicated hpi, and infectious virus titers were determined by FFA. f, g Susceptibility of rCl13 and rCl13-RRRR to furin inhibitor BOS-318 (f) and S1P inhibitor PF-429242 (g). A549 cells were seeded in 96-well plates at 2 × 104 cells/well and 24 h later infected (MOI = 0.01) with the indicated virus and treated with serial dilutions (1:2, 6 replicates per dilution) of either BOS-318 (starting at 2 µM), or PF-429242 (starting at 40 µM). At 72 hpi, cell viability was determined by CellTiter 96 Aqueous One Solution (purple), then cells were fixed with 4% PFA. The level of infection was assessed by IF using the rat monoclonal antibody VL4 to NP (green), and cell viability was also alternatively determined by DAPI staining (blue). Raw values of CellTiter, NP, and DAPI staining were normalized to vehicle-treated and infected control cells present in each plate. Dose-response curves for BOS-318 and PF-429242 were generated using a nonlinear regression model with log(inhibitor) vs response – variable response (four parameters) equation in GraphPad Prism. The dashed line represents 50% of vehicle control. h Effect of PF-429242 on multi-step growth kinetics of rCl13 and rCl13-RRRR in A549 cells. A549 cells were seeded at 2.5 × 105 cells/well in a 24-well plate and 24 h later infected (MOI = 0.01) with the indicated virus. After 90 min adsorption, the virus inoculum was removed, and medium containing the indicated concentration of PF-429242 was added to the cells. At the indicated hpi, CCSs were collected and virus titers determined by FFA. Cells lacking S1P or treated with PF-429242 were grown in media supplemented with lipids (1 mM sodium mevalonate, 20 μM sodium oleate, 5 μg/ml cholesterol).
Cell-based fusion assays, where GPC was delivered via transfection, revealed no differences among GPC-WT, GPC-RRRR and GPC-RRLL in their ability to trigger the pH-dependent membrane fusion (Fig. 2a). Treatment with the furin inhibitor BOS-318 inhibited the fusion activity only of GPC-RRRR (Fig. 2b). Staining with an antibody to GPC displayed similar levels of GFP expression under all experimental conditions, indicating that BOS-318 inhibition of fusion mediated by GPC-RRRR was not due to reduced levels of GPC expression.
a GPC-RRRR had similar fusion activity compared to GPC-WT. HEK293T cells were seeded at 2.5 × 105 cells/well in a 24-well plate and 24 h later transfected with pCAGGS plasmids (1 µg) expressing LCMV GPC-WT, GPC-RRRR, GPC-RRLL, or an empty plasmid, together with 50 ng of pCAGGS-GFP. At 48 h post transfection, cell monolayers were treated with citrate buffer (pH 5.0) or PBS (pH 7.0) for 15 min, then returned to neutral pH normal growth media (DMEM containing 10% FBS) and fusion monitored over time. Once fusion activity was observed (~20 min) cells were allowed 10 min to complete fusion then fixed (4% PFA) and stained with a human monoclonal antibody to GPC (red). Fusion was visualized based on the distinctive appearance of GFP signal. b GPC-RRRR mediated fusion is inhibited by treatment with the furin inhibitor BOS-318. HEK293T cells were prepared and transfected as in (a) but also treated with BOS-318 (2 µM) or DMSO at the time of transfection, or at 46 h post transfection. Fusion assay was done at 48 h post transfection. Representative images from each condition are shown. All images were taken on a Zeiss LSM 780 Confocal laser scanning microscope.
S1P-mediated GPC cleavage is required for persistence of rCl13 in immunocompetent mice
Adult immunocompetent B6 mice infected with LCMV Cl13 (2 × 106 FFU/mouse, IV) have been shown to develop a long-term persistent infection with sustained high viremia49,50,51,52,54,55,56,57. Consistent with published findings, B6 mice infected (2x106 FFU/mouse, IV) with either rCl13 or rCl13-RRLL, both dependent on S1P for GPC processing, persisted for more than 30 days with rCl13-RRLL consistently showing lower levels of viremia compared to rCl13 (Fig. 3a). In contrast, B6 mice infected with rCl13-RRRR (2 × 106 FFU/mouse, IV) had viremia at or below the limit of detection (LoD) at 3 and 7 days after infection and undetectable thereafter throughout the duration of the experiment (Fig. 3a). We detected LCMV-specific antibody titers in serum samples collected from mice at 14 days post infection (dpi) with rCl13-RRRR, confirming positive infection and seroconversion (Fig. 3b). We also determined viral load in multiple organs collected from mice at different times after infection. Mice infected with rCl13-RRRR had consistently lower tissue viral load compared to mice infected with rCl13 (Fig. 3c). Tissue viral load in rCl13-infected mice started to increase from 1 dpi to at least 6 dpi, while in rCl13-RRRR-infected mice, tissue viral load was higher in earlier timepoints, but lower compared to rCl13, and started to decrease after 4 dpi. At 15 dpi, organs collected from rCl13-RRRR infected mice had undetectable levels of infectious virus, while all tested tissues from rCl13-infected mice had high virus titers, including significant viral titer in brain tissues, which had low or no detectable virus in prior timepoints (Fig. 3c).
a B6 mice (n = 5/group) were infected with rCl13, rCl13-RRR or rCl13-RRLL (2 × 106 FFU/mouse, IV). Blood samples were collected using the retro orbital (r.o.) route at the indicated times pi and viremia determined by FFA. b Serum from rCl13-RRRR infected mice at 14 days post infection (dpi) was tested for seroconversion against LCMV by ELISA. Mouse anti-LCMV antibodies were detected by HRP-conjugated goat anti-mouse secondary antibody. Plates were read using a CLARIOstar Plus microplate reader at 450 nm absorption. c B6 mice (n = 3/group for 1 dpi, n = 5/group for 2, 4, 6, and 15 dpi) infected with rCl13 or rCl13-RRRR (2 × 106 FFU/mouse, IV) were euthanized at the indicated days post infection and tissues (brain, lung, liver, spleen, and kidney) as well as serum were collected. Tissue homogenates were prepared and virus titers determined by FFA. d B6 mice were infected with rCl13 (n = 3/group, 2 × 106 FFU/mouse, IV) or rCl13-RRRR (n = 4/group, 2 × 106 FFU/mouse, IV), or co-infected at the same time with both viruses (n = 5/group, 2 × 106 FFU/mouse of each virus, IV). Blood was collected r.o. at indicated times pi, and viremia determined by FFA. e Serum samples from co-infected mice in (d) were used to infect (15 µL serum/well in 24-well plate) WT or S1P-KO HAP1 cells. CCSs were collected at 72 hpi and virus titers determined by FFUA. All statistical tests were done with two-way ANOVA in GraphPad Prism. Number of asterisks indicates the p value (*p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; ****p ≤ 0.0001).
We reasoned that rCl13-RRRR infection could trigger at early times of infection a shift in the host immune response that favored efficient viral clearance and therefore expect a faster clearance of rCl13 in mice co-infected with rCl13 and rCl13-RRRR compared to mice infected with rCl13 alone. However, mice infected only with rCl13 or co-infected with both rCl13 and rCl13-RRRR exhibited similar viremia levels at 5, 10, 20, 30, and 40 dpi, although co-infection with rCl13 and rCl13-RRRR did slightly accelerate clearance of rCl13 (~50 dpi) (Fig. 3d). We characterized viruses present in sera from co-infected mice with respect to their ability to multiply in WT and S1P-KO HAP1 cells (Fig. 3e), as WT HAP1 cells allowed propagation of both rCl13 and rCl13-RRRR whereas S1P-KO HAP1 cells prevented propagation of rCl13. We infected WT and S1P-KO HAP1 cells with serum samples (15 µL, MOI ~5 × 10−4) from 5, 10, 20, 30, and 40 dpi, and at 72 h post infection (hpi) determined virus titers in CCSs. For all timepoints titrated (5, 10, and 20 dpi), virus titers in CCS of WT HAP1 cells were 10–100-fold higher than in CCS from S1P-KO HAP1 cells (Fig. 3e), indicating that viral populations present in serum samples from co-infected mice were dominated by rCl13, whose GPC processing is mediated by S1P. We also used nanopore sequencing to genetically characterize the infectious progenies from WT and S1P-KO HAP1 cells infected with the serum samples. All viral GPC sequences from WT HAP1 cells contained the S1P cleavage site (RRLA). In contrast, viral GPC sequences derived from S1P-KO HAP1 cells infected with serum samples from 5, 10 and 20 dpi showed a mix of both S1P (RRLA) and furin (RRRR) cleavage sites. At 30 dpi, two out of five serum samples still consisted of a mix of both cleavage sites, whereas at 40 dpi, all serum samples only contained the S1P site sequence. As control, we also infected WT and S1P-KO HAP1 cells with serum samples from rCL13-RRRR infected mice collected at 5, 10, and 20 dpi. However, due to the very low serum titers of the 5 dpi samples (Fig. 3d), no infectious viral progeny was detected in CCS samples of WT and S1P-KO HAP1 cells at 72 hpi. As expected, infection of WT and S1P-KO HAP1 cells with 10 and 20 dpi serum samples did not produce detectable infectious virus, as these serum samples had undetectable viremia levels. These results suggested that co-infection with rCl13 can rescue the persistence phenotype of rCl13-RRRR for up to 30 days, but rCl13-RRRR was still cleared more efficiently than rCl13.
rCl13-RRRR and rCl13 exhibit different splenic cellular tropism and distribution
Published evidence supports that early events after infection can dictate the ability of Cl13 to establish persistence in adult immunocompetent mice68. We hypothesized that differences at early stages of infection resulted in rCl13, but not rCl13-RRRR, ability to persist. To examine this question, we assessed the splenic cellular tropism and distribution of rCl13 and rCl13-RRRR by flow cytometry using staining of intracellular LCMV NP to identify infected cells and validated cell surface markers to identify the different cell populations. At 1 dpi, fewer pDC, Ly6- monocytes and neutrophiles were NP+ in rCl13-RRRR compared to rCl13-infected mice. However, the numbers of NP+ macrophages were much higher in rCl13-RRRR than in rCl13-infected mice (Fig. 4a, b and Fig. S4). Segregation of total macrophages into different subsets revealed that compared to rCl13, rCl13-RRRR infected a higher percentage of red pulp macrophages (RPMs) but exhibited reduced efficiency of infection of CD169+ metallophilic marginal-zone macrophages (MMMs) (Fig. 4c, d and Fig. S5). To visualize the differences in viral distribution within the spleen, we prepared frozen sections from spleens collected at 1 dpi and stained them for CD169, a marker for MMMs located on the inner border of marginal zones adjacent to the white pulp, and an antibody to LCMV GPC to identify infected cells. Higher degree of colocalization (yellow) between CD169 and GPC was observed in spleens from rCl13 than rCl13-RRRR-infected mice (Fig. 4e). In rCl13-RRRR-infected mice, GP staining was limited to the red pulp area outside of the boundaries drawn by the MMMs, a finding consistent with the flow results. Staining for the viral antigen (GP) was in overall lower in spleens from rCl13-RRRR than rCl13-infected mice, a finding consistent with the overall lower viral load in rCl13-RRRR-infected mice compared to rCl13-infected mice (Fig. 4f). Together, these findings indicated that switching from S1P to furin-mediated processing of GPC, results in a virus that is impaired in its ability to infect MMMs that line the marginal sinus between the white and red pulp of the spleen, which may contribute to rCl13-RRRR being unable to infiltrate the white pulp.
a Eight-week-old B6 mice were infected with rCl13, rCl13-RRRR (n = 5/group, 2 × 106 FFU/mouse, IV), or mock-infected (n = 2, 200 µL PBS, IV). Spleens were harvested at 24 hpi and processed to generate single-cell suspensions. Specific cell types were identified by flow cytometry through surface marker staining: B cells: B220+, CD3−, CD4−, Siglec-H−; CD4 T cells: B220−, CD3+, CD4+; CD8 T cells: B220−, CD3+, CD8+; plasmacytoid dendritic cells (pDCs): B220+, CD3−, CD4+, Siglec-H+; natural killer (NK) cells: B220−, CD3−, NK1.1+; CD8+ dendritic cells (DCs): B220−, CD3−, NK1.1−, Ly6G−, CD11chigh, MHC-II+, CD11blow, CD8+; CD11b+ cDCs: B220−, CD3−, NK1.1−, Ly6G−, CD11chigh, MHC-II+, CD11b+, CD8−; macrophages: B220−, CD3−, NK1.1−, Ly6G−, CD11clow, CD11b−, F4/80+; Ly6C+ monocytes: B220−, CD3−, NK1.1−, Ly6G−, CD11clow, CD11bhigh, Ly6Chigh; Ly6C- monocytes: B220−, CD3−, NK1.1−, Ly6G−, CD11clow, CD11bhigh, Ly6Clow. Intracellular NP was stained with AF488-conjugated VL4 rat monoclonal antibody. b shows the representative contour plots of NP+ gates from the macrophage populations. c, d Infection of B6 mice and processing of spleen samples were done as in (a). Specific cell types were identified by flow cytometry through surface marker staining: metallophilic marginal-zone macrophages (MMM): lin-(CD19−, CD3−, NK1.1−), Ly6G−, Ly6C−, F4/80+, CD11bhigh, CD169high; red pulp macrophages (RPM): lin-(CD19−, CD3−, NK1.1−), Ly6G−, Ly6C−, F4/80+, CD11blow, CD169−. Intracellular NP was stained with AF488-conjugated VL4 rat monoclonal antibody. Representative contour plots of NP+ gates from the parental gates are shown. A statistical test (unpaired t-test) was done between rCl13 and rCl13-RRRR groups in GraphPad Prism. P values are shown or represented by asterisks (ns not significant; *p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; ****p ≤ 0.0001). e, f Eight-week-old B6 mice were infected as in (a). Spleens were collected at 24 hpi and split in half. One half was flash frozen in OCT and 8 µm tissue sections were stained with human anti-GPC antibody (green), rat anti-CD169 antibody (red) and DAPI (blue). Sections were prepared for four mice per group, and representative images from each group are shown. The other half of the spleen was homogenized and virus titers determined by FFA. A statistical test (unpaired t-test) was done between rCl13 and rCl13-RRRR groups in GraphPad Prism. P values are represented by asterisks (ns not significant; *p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; ****p ≤ 0.0001).
Interferon (IFN) and CD8+ T cells are required for the clearance of rCl13-RRRR
We observed that at 24 hpi, B6 mice infected with rCl13-RRRR had lower systemic IFNα in serum compared to mice infected with rCl13 (Fig. 5a). To investigate the role of IFN and identify other host factors that may contribute to the clearance of rCl13-RRRR, we examined the ability of rCl13-RRRR to persist in B6 mice deficient for different immune components. In IFNαR-KO mice, rCl13-RRRR reached and sustained similar viremia levels as rCl13 (Fig. 5b), demonstrating that rCl13-RRRR can replicate to high titers in an immunocompromised host. IFNαR blockade with MAR1 antibody69,70,71 also rescued high serum titers (>104 FFU/mL) of rCl13-RRRR in WT B6 mice at 3 and 7 dpi, followed by reduced viremia levels after 14 dpi (Fig. 5c). Early and efficient clearance of rCL13-RRRR was not affected in IL27αR-KO mice or following CD4+ T cell depletion (Fig. S1). However, CD8+ T cell depletion by treatment with the rat monoclonal antibody anti-CD8 YTS 169.472,73,74,75 prevented clearance of rCl13-RRRR with serum virus titers of ~103 FFU/mL detected at 4 and 7 dpi (Fig. 5d), whereas in mice treated with a rat IgG2b control, serum titers of rCL13-RRRR were at or lower than the limit of detection (LoD). We used control Armstrong strain of LCMV (rARM) known to cause an acute infection that is controlled and cleared by CD8+ T cells74 to verify positive depletion of CD8+ T cells. These results revealed that, similarly to rARM as previously reported74, clearance of rCl13-RRRR depends on CD8+ rather than CD4+ T cells.
a Eight-week-old WT B6 mice (n = 4/group) were infected (2 × 106 FFU/mouse, IV) with rCl13 or rCl13-RRRR. At the indicated hpi, blood samples were collected r.o. and levels of IFNα in serum were measured by ELISA. b, c IFNαR KO mice (n = 5/group) (b), and WT B6 mice (n = 5/group) treated with IFNαR blockade antibody (MAR1) (c) were infected with rCl13 or rCl13-RRRR. Blood samples were collected r.o. at the indicated dpi and serum virus titers determined by FFA. d WT B6 mice (n = 5/group) were treated with CD8 depletion antibody (YTS 169.4) or rat IgG2b isotype control, then infected (2 × 106 FFU/mouse, IV) with rCl13-RRRR or a recombinant Armstrong strain of LCMV (rARM). Blood samples were collected r.o. at 4 and 7 dpi and serum virus titers determined by FFA. e, f WT B6 mice (n = 5/group) were infected with rCl13, rCl13-RRRR, or rARM. Spleens were harvested at 7 dpi and processed to generate single-cell suspensions. Total splenocytes were stimulated with LCMV GP33 peptides for 5 h, and the activated CD8+ T cells (B220−, CD4−, CD8+, CD44+) were stained to detect intracellular levels of IFNγ and TNFα and analyzed by flow cytometry. Representative contour plots of IFNγ+ and IFNγ+ TNFα+ double-positive gates from the parental gates are shown. All statistical tests were done with one-way or two-way ANOVA in GraphPad Prism. Number of asterisks indicates the p value (ns not significant; *p ≤ 0.05; **p ≤ 0.01; ***p ≤ 0.001; ****p ≤ 0.0001).
Infection of B6 mice with rCl13 is known to cause T cell exhaustion and decreased CD8+ T cell function58,76,77,78,79,80,81,82. Since immunocompetent mice can efficiently clear rCl13-RRRR, we hypothesized that CD8+ T cells retain their function in mice that cleared rCl13-RRRR infection. To confirm this prediction, we collected spleens from B6 mice at 7 dpi with rCl13 or rCl13-RRRR and prepared total splenocyte suspensions that were tested for their responses to stimulation with LCMV GP33 peptide as determined by intracellular cytokine staining (ICS) by flow cytometry. The percentage of IFNγ+ activated CD8+ T cells was higher in splenocytes derived from rCl13-RRRR-infected mice than in splenocytes derived from mice infected with either rCl13 or rARM (Fig. 5e and Fig. S6). However, rARM-infected mice consistently had larger spleen sizes (data not shown), reflected in higher absolute numbers of IFNγ+ cells. Percentages of double positive IFNγ+ TNFα+ cells (Fig. 5f and S6) were higher in splenocytes derived from rCl13-RRRR than rCl13-infected mice, indicating that infection with rCl13-RRRR did not result in functional exhaustion of CD8+ T cells however did significantly reduce the magnitude of functional CD8 T cells compared to rARM infection.
rCl13-RRRR is attenuated in mouse models of LCMV lethal infection and can induce protective immunity against a subsequent lethal challenge
Consistent with published data58,59, we found that FVB mice infected IV with a high dose (2 × 106 FFU/mouse) of rCl13 or rCl13-RRLL developed a lethal disease characterized by large (15–20%) body weight loss and severe CD8 + T cell-mediated lung pathology, with 100% of rCl13 and 80% of rCl13-RRLL infected mice succumbing to infection by 14 dpi (Fig. 6a, b). In contrast, FVB mice infected with rCl13-RRRR (2 × 106 FFU/mouse, IV) exhibited minimal clinical signs, including slightly ruffled coat around 4 dpi and modest loss of body weight, with all (100%) recovering from infection (Fig. 6a, b). Seroconversion was confirmed in all five rCl13-RRRR-infected mice at 14 dpi (Fig. S2). At 5 dpi viremia was drastically reduced in rCl13-RRRR-infected compared to rCl13-infected mice, and by 14 dpi, all rCl13-RRRR-infected mice had undetectable levels of viremia (Fig. 6c). The one surviving mouse infected with rCl13-RRLL still had detectable viremia (~350 FFU/mL) at 14 dpi.
a–c rCl13-RRRR is attenuated in the FVB mouse model of Cl13 infection. Seven-week-old FVB mice were infected with rCl13, rCl13-RRRR, rCl13-RRLL (n = 5/group, 2 × 106 FFU/mouse, IV) or mock-infected (n = 3, PBS 200 µL, IV), and monitored daily for clinical symptoms, including changes in body weight (a), and survival (b). Blood samples were collected r.o. at 5 dpi, and from surviving mice at the experimental endpoint (14 dpi), and serum virus titers determined by FFA (c). d rCl13-RRRR is highly attenuated in the fatal LCM model of LCMV infection. Eight-week-old B6 mice were challenged intracranially with rCl13 or rCl13-RRRR at the indicated doses and monitored daily for clinical symptoms and survival. e–g Infection with rCl13-RRRR induces a protective immune response in FVB mice against a lethal challenge with rCl13. Surviving FVB mice from the rCl13-RRRR group (n = 5/5) and PBS group (n = 3/3) were challenged with rCl13 at 40 dpi and monitored daily for changes in body weight (e) and survival (f). Blood samples were collected r.o. at 10 dpi of the secondary challenge and serum virus titers determined by FFA (g).
To further characterize the in vivo virulence of rCl13-RRRR, we tested the virus in a lethal LCM model of LCMV infection, where naïve B6 mice succumb to death within 2 weeks following an intracranial (IC) challenge with rCl13, using a viral dose as low as 1 FFU as a result of CD8+ T cell-mediated pathology83,84,85. We infected (IC) mice with 1000, 100, or 10 FFU of rCl13 or rCl13-RRRR. All mice inoculated with rCl13 succumbed to death 8-11 dpi, depending on the dosage, while rCl13-RRRR caused a lethal infection only at the highest dose of 1000 FFU (Fig. 6d), indicating that rCl13-RRRR has an LD50 about 100-fold higher than that of rCl13.
Since all rCl13-RRRR-infected FVB mice seroconverted and survived without noticeable clinical manifestations, we hypothesized that these mice may have developed a protective immune response against a subsequent lethal challenge with rCl13. To test this hypothesis, we infected (IV) surviving rCl13-RRRR-infected mice (Fig. 4a) with 2 × 106 FFU/mouse of rCl13 at 40 days post initial infection. None of the mice challenged with the second rCl13 infection lost weight or developed any other clinical symptoms (Fig. 6e), whereas 100% of initially PBS-inoculated control mice developed clinical symptoms and succumbed to the rCl13 challenge by 10 dpi (Fig. 6f). Consistent with the lack of clinical symptoms, we did not detect viremia in rCl13-RRRR immunized mice at day 10 post rechallenge with rCl13 (Fig. 6g). We could not collect blood from mice in the PBS group because all animals had succumbed to death by 10 dpi. We also found that all (100%, n = 5) rCl13-RRRR immunized (2 × 106 FFU/mouse, IV) B6 mice survived a subsequent lethal IC challenge (1000 FFU) with rCl13 at 35 days post initial rCl13-RRRR infection (Fig. S3). These findings indicated that rCl13-RRRR is highly attenuated in vivo but able to induce a protective immune response against a subsequent viral lethal challenge.
Discussion
In this study, we present evidence that replacement of the S1P with the furin recognition cleavage site in the GPC of the immunosuppressive variant Cl13 of LCMV resulted in a virus, rCl13-RRRR, that exhibited rCl13 WT-like fitness in cell culture, demonstrating that rCl13, similarly to rARM48, can efficiently use a host cell protease other than S1P for the processing of its GPC. However, we found that rCl13-RRRR is greatly attenuated in immunocompetent mice both in terms of its ability to establish a persistent infection and pathogenicity. The Cl13 variant of the ARM strain of LCMV is able, following IV infection with 2 × 106 FFU, to cause a persistent infection in adult immunocompetent B6 mice that is characterized by high and sustained (>40 days) viremia levels. Cl13 only has three amino acid mutations compared to the parental ARM strain: N176D and F260L in GPC, and K1079Q in L, with mutations GPC (F260L) and L (K1079Q) being strictly required for Cl13 persistence in vivo49,54,55. The K1079Q mutation in the viral polymerase is linked to higher replicative potential of Cl13 in macrophages compared to ARM49,86, whereas mutation F260L in GPC is associated with high binding affinity to the cellular receptor of LCMV, alpha dystroglycan (αDG)51,87,88. Our results suggest that S1P-mediated cleavage of GPC represents a third requirement for Cl13 persistence, as rCl13-RRRR could not persist despite having all three mutations (GPC: N176D; F260L and L: K1079Q) previously identified as being associated with Cl13 persistence. The S1P cleavage site (262-265) is near L260, and it is plausible that the change 262RRLA265 (S1P) to 262RRRR265 (furin) could affect the conformation of the receptor binding site in GP1, lowering its affinity to αDG. This possibility seems unlikely as rCl13-RRRR did not exhibit, compared to rCl13, reduced fitness in cultured cells or in immunocompromised mice. A detailed assessment, beyond the scope of the present work, of the potential effect of mutations LA to RR on the structure of GPC, and its biological implications, will be required to address this issue.
We found that rCl13 and rCl13-RRRR exhibited differences in their splenic cellular tropism at early (1 dpi) times of infection, with spleens from rCl13-RRRR-infected mice containing about a two-fold increase in NP+ macrophages compared to spleens from rCl13-infected mice (Fig. 4a). However, spleens from rCl13-RRRR-infected mice showed only about a 5% increase in NP+ red pulp macrophages (RPMs) compared to spleens from rCl13-infected mice, and fewer MMMs were NP+ in rCl13-RRRR than rCl13-infected mice (Fig. 4b). These findings suggest that other macrophage subtypes that we did not capture in our analysis exhibit differential susceptibility to infection with rCl13 and rCl13-RRRR. We also observed higher numbers of CD8+ T and NK cells but lower numbers of pDC, Ly6C- monocytes and neutrophils being infected by rCl13-RRRR than by rCl13. Future studies will evaluate the biological implications of these findings. Intriguingly, analysis of virus splenic cellular tropism at 2 and 3 dpi revealed, for all cell types examined, higher numbers of NP+ cells in rCl13 than rCl13-RRRR-infected mice, suggesting that early events during the first 24 h of infection may dictate the ability of rCl13 to persist, whereas rCl13-RRRR is controlled and cleared.
Treatment of rCl13-RRRR-infected B6 mice with the IFNαR blockade antibody mostly rescued high and sustained (>30 days) viremia (Fig. 5bii), followed by reduced rCl13-RRRR viremia and its clearance at later timepoints, which might reflect the waning of IFNαR blockade antibody and inability of rCl13-RRRR to persist in an immunocompetent host environment. However, IFNαR blockade resulted also in control and faster clearance of rCl13, as previously reported71, and the same underlying mechanisms could have also contributed to the control and clearance of rCl13-RRRR.
To our knowledge, this is the first study where the role of S1P-mediated processing of GPC was investigated in vivo. Mammarenaviruses are zoonotic viruses that establish life-long persistent infection in their natural rodent hosts7,89. As human-to-human transmissions are infrequent events, the main driver for mammarenavirus evolution are the selective pressures in its rodent natural host89. Thus, the strict S1P dependency may be the consequence of evolution within their natural hosts, as choosing S1P could provide mammarenaviruses with survival advantages, for example, being able to persist while not causing noticeable clinical manifestations, so it is likely to be transmitted to other uninfected individuals. However, the exact evolutionary pressure that drove the dependence towards S1P rather than any other proteases remains to be determined. As our work has demonstrated, the furin-dependent rCl13-RRRR was only attenuated in vivo, and it seemed that early viral-host interactions immediately after infection were crucial for the infection outcome. Follow-up studies are required to uncover the key differences in these interactions between the rCl13-RRRR and rCl13.
The discoveries presented in this paper provide valuable insights regarding antiviral and vaccine designs to combat human pathogenic mammarenaviruses. Small molecules targeting S1P can be developed as host-directed antivirals against human pathogenic mammarenaviruses, an approach that is supported by existing evidence48,90,91,92,93,94,95. Switching from S1P to furin-mediated processing of GPC resulted in a high degree of virus attenuation in vivo, supporting the minimal risk posed by an S1P inhibitor-mediated selection of viruses capable of using a host cell protease other than S1P for GPC processing, which has been observed in vitro previously as well48. The fact that rCl13-RRRR was controlled and rapidly cleared in immunocompetent mice in the absence of noticeable clinical symptoms but able to trigger a protective immune response against a subsequent lethal challenge, suggests that replacement of the S1P by the furin recognition cleavage site represents a virus genetic determinant of attenuation that should be considered for its incorporation into the design of mammarenavirus live-attenuated vaccines (LAV). Currently there is no FDA-licensed vaccine against human pathogenic mammarenaviruses, although multiple strategies have been proposed2,96,97,98,99,100,101,102,103,104,105.
It will need to be determined whether a recombinant LASV whose GPC is processed by furin instead of S1P exhibits the same attenuated in vivo phenotype as rCl13-RRRR. Moreover, the fact that rCl13-RRRR did persist in immunocompromised mice would raise concerns about a LASV LAV solely based on its GPC being processed by furin instead of S1P, as the target population for a LASV vaccine is likely to include people with different degrees of immunosuppression due to the high prevalence of malaria and HIV in Western Africa. However, replacement of the S1P by the furin cleavage recognition site in GPC of LASV could be an additional safety determinant that could be incorporated into recombinant forms of LASV expressing a codon deoptimized GPC99 or containing reorganized L-IGR96 that have already been shown to exhibit excellent features as LASV LAV candidates.
Data availability
The datasets generated and/or analyzed during the current study are available from the corresponding author on reasonable request.
Code availability
The authors did not generate or use any custom code in this manuscript.
References
Asogun, D. A., Gunther, S., Akpede, G. O., Ihekweazu, C. & Zumla, A. Lassa fever: epidemiology, clinical features, diagnosis, management and prevention. Infect. Dis. Clin. North Am. 33, 933–951 (2019).
Garry, R. F. Lassa fever - the road ahead. Nat. Rev. Microbiol. 21, 87–96 (2023).
Frame, J. D., Baldwin, J. M. Jr., Gocke, D. J. & Troup, J. M. Lassa fever, a new virus disease of man from West Africa. I. Clinical description and pathological findings. Am. J. Trop. Med. Hyg. 19, 670–676 (1970).
Basinski, A. J. et al. Bridging the gap: Using reservoir ecology and human serosurveys to estimate Lassa virus spillover in West Africa. PLoS Comput. Biol. 17, e1008811 (2021).
Sogoba, N. et al. Lassa virus seroprevalence in Sibirilia Commune, Bougouni District, Southern Mali. Emerg. Infect. Dis. 22, 657–663 (2016).
Grant, D. S. et al. Seroprevalence of anti-Lassa Virus IgG antibodies in three districts of Sierra Leone: a cross-sectional, population-based study. PLoS Negl. Trop. Dis. 17, e0010938 (2023).
Gunther, S. & Lenz, O. Lassa virus. Crit. Rev. Clin. Lab. Sci. 41, 339–390 (2004).
Grant, A. et al. Junin virus pathogenesis and virus replication. Viruses 4, 2317–2339 (2012).
Lendino, A., Castellanos, A. A., Pigott, D. M. & Han, B. A. A review of emerging health threats from zoonotic New World mammarenaviruses. BMC Microbiol. 24, 115 (2024).
Bonthius, D. J. Lymphocytic choriomeningitis virus: a prenatal and postnatal threat. Adv. Pediatr. 56, 75–86 (2009).
Charrel, R. N. et al. Acquired hydrocephalus caused by a variant lymphocytic choriomeningitis virus. Arch. Intern. Med. 166, 2044–2046 (2006).
Fischer, S. A. et al. Transmission of lymphocytic choriomeningitis virus by organ transplantation. N. Engl. J. Med. 354, 2235–2249 (2006).
Kinori, M., Schwartzstein, H., Zeid, J. L., Kurup, S. P. & Mets, M. B. Congenital lymphocytic choriomeningitis virus-an underdiagnosed fetal teratogen. J. AAPOS 22, 79–81.e71 (2018).
Larsen, P. D., Chartrand, S. A., Tomashek, K. M., Hauser, L. G. & Ksiazek, T. G. Hydrocephalus complicating lymphocytic choriomeningitis virus infection. Pediatr. Infect. Dis. J. 12, 528–531 (1993).
Yu, J. T., Culican, S. M. & Tychsen, L. Aicardi-like chorioretinitis and maldevelopment of the corpus callosum in congenital lymphocytic choriomeningitis virus. J. AAPOS 10, 58–60 (2006).
Gass, J. T. et al. Discovery of a novel lymphocytic choriomeningitis virus strain associated with severe human disease in immunocompetent patient, New Mexico. Emerg. Microbes Infect. 14, 2542250 (2025).
Bausch, D. G., Hadi, C. M., Khan, S. H. & Lertora, J. J. Review of the literature and proposed guidelines for the use of oral ribavirin as postexposure prophylaxis for Lassa fever. Clin. Infect. Dis. 51, 1435–1441 (2010).
Cheng, H. Y. et al. Lack of evidence for ribavirin treatment of Lassa fever in systematic review of published and unpublished studies(1). Emerg. Infect. Dis. 28, 1559–1568 (2022).
Eberhardt, K. A. et al. Ribavirin for the treatment of Lassa fever: a systematic review and meta-analysis. Int. J. Infect. Dis. 87, 15–20 (2019).
Raabe, V. N. et al. Favipiravir and ribavirin treatment of epidemiologically linked cases of Lassa fever. Clin. Infect. Dis. 65, 855–859 (2017).
Salam, A. P. et al. Ribavirin for treating Lassa fever: a systematic review of pre-clinical studies and implications for human dosing. PLoS Negl. Trop. Dis. 16, e0010289 (2022).
Enria, D. A., Briggiler, A. M. & Sanchez, Z. Treatment of Argentine hemorrhagic fever. Antivir. Res. 78, 132–139 (2008).
Southern, P. J. et al. Molecular characterization of the genomic S RNA segment from lymphocytic choriomeningitis virus. Virology 157, 145–155 (1987).
Riviere, Y. et al. The S RNA segment of lymphocytic choriomeningitis virus codes for the nucleoprotein and glycoproteins 1 and 2. J. Virol. 53, 966–968 (1985).
Pinschewer, D. D., Perez, M. & de la Torre, J. C. Role of the virus nucleoprotein in the regulation of lymphocytic choriomeningitis virus transcription and RNA replication. J. Virol. 77, 3882–3887 (2003).
Lee, K. J., Novella, I. S., Teng, M. N., Oldstone, M. B. & de La Torre, J. C. NP and L proteins of lymphocytic choriomeningitis virus (LCMV) are sufficient for efficient transcription and replication of LCMV genomic RNA analogs. J. Virol. 74, 3470–3477 (2000).
Salvato, M. S. & Shimomaye, E. M. The completed sequence of lymphocytic choriomeningitis virus reveals a unique RNA structure and a gene for a zinc finger protein. Virology 173, 1–10 (1989).
Strecker, T. et al. Lassa virus Z protein is a matrix protein and sufficient for the release of virus-like particles. J. Virol. 77, 10700–10705 (2003).
Perez, M., Craven, R. C. & de la Torre, J. C. The small RING finger protein Z drives arenavirus budding: implications for antiviral strategies. Proc. Natl. Acad. Sci. USA 100, 12978–12983 (2003).
Urata, S. & Yasuda, J. Molecular mechanism of arenavirus assembly and budding. Viruses 4, 2049–2079 (2012).
Garten, W. in Activation of Viruses by Host Proteases Ch. 9 (Böttcher-Friebertshäuser, E., Garten, W., Klenk, H-D). (Springer, 2018).
Pasquato, A., Cendron, L. & Kunz, S. in Activation of Viruses by Host Proteases Ch. 3 (Böttcher-Friebertshäuser, E., Garten, W., Klenk, H-D). (Springer, 2018).
Beyer, W. R., Popplau, D., Garten, W., von Laer, D. & Lenz, O. Endoproteolytic processing of the lymphocytic choriomeningitis virus glycoprotein by the subtilase SKI-1/S1P. J. Virol. 77, 2866–2872 (2003).
Lenz, O., ter Meulen, J., Klenk, H. D., Seidah, N. G. & Garten, W. The Lassa virus glycoprotein precursor GP-C is proteolytically processed by subtilase SKI-1/S1P. Proc. Natl. Acad. Sci. USA 98, 12701–12705 (2001).
Kunz, S., Edelmann, K. H., de la Torre, J.-C., Gorney, R. & Oldstone, M. B. A. Mechanisms for lymphocytic choriomeningitis virus glycoprotein cleavage, transport, and incorporation into virions. Virology 314, 168–178 (2003).
Wright, K. E., Spiro, R. C., Burns, J. W. & Buchmeier, M. J. Post-translational processing of the glycoproteins of lymphocytic choriomeningitis virus. Virology 177, 175–183 (1990).
Rojek, J. M., Lee, A. M., Nguyen, N., Spiropoulou, C. F. & Kunz, S. Site 1 protease is required for proteolytic processing of the glycoproteins of the South American hemorrhagic fever viruses Junin, Machupo, and Guanarito. J. Virol. 82, 6045–6051 (2008).
Hastie, K. M. et al. Crystal structure of the prefusion surface glycoprotein of the prototypic arenavirus LCMV. Nat. Struct. Mol. Biol. 23, 513–521 (2016).
Richardson, C. et al. The nucleotide sequence of the mRNA encoding the fusion protein of measles virus (Edmonston strain): a comparison of fusion proteins from several different paramyxoviruses. Virology 155, 508–523 (1986).
Volchkov, V. E., Feldmann, H., Volchkova, V. A. & Klenk, H. D. Processing of the Ebola virus glycoprotein by the proprotein convertase furin. Proc. Natl. Acad. Sci. USA 95, 5762–5767 (1998).
Keelapang, P. et al. Alterations of pr-M cleavage and virus export in pr-M junction chimeric dengue viruses. J. Virol. 78, 2367–2381 (2004).
McCune, J. M. et al. Endoproteolytic cleavage of gp160 is required for the activation of human immunodeficiency virus. Cell 53, 55–67 (1988).
Follis, K. E., York, J. & Nunberg, J. H. Furin cleavage of the SARS coronavirus spike glycoprotein enhances cell-cell fusion but does not affect virion entry. Virology 350, 358–369 (2006).
Peacock, T. P. et al. The furin cleavage site in the SARS-CoV-2 spike protein is required for transmission in ferrets. Nat. Microbiol. 6, 899–909 (2021).
Klenk, H. D., Rott, R., Orlich, M. & Blodorn, J. Activation of influenza A viruses by trypsin treatment. Virology 68, 426–439 (1975).
Urata, S. et al. Analysis of assembly and budding of Lujo virus. J. Virol. 90, 3257–3261 (2015).
Altamura, L. A. et al. Identification of a novel C-terminal cleavage of Crimean-Congo hemorrhagic fever virus PreGN that leads to generation of an NSM protein. J. Virol. 81, 6632–6642 (2007).
Rojek, J. M. et al. Targeting the proteolytic processing of the viral glycoprotein precursor is a promising novel antiviral strategy against arenaviruses. J. Virol. 84, 573–584 (2010).
Matloubian, M., Kolhekar, S. R., Somasundaram, T. & Ahmed, R. Molecular determinants of macrophage tropism and viral persistence: importance of single amino acid changes in the polymerase and glycoprotein of lymphocytic choriomeningitis virus. J. Virol. 67, 7340–7349 (1993).
Mueller, S. N. et al. Viral targeting of fibroblastic reticular cells contributes to immunosuppression and persistence during chronic infection. Proc. Natl. Acad. Sci. USA 104, 15430–15435 (2007).
Sevilla, N. et al. Immunosuppression and resultant viral persistence by specific viral targeting of dendritic cells. J. Exp. Med. 192, 1249–1260 (2000).
Ahmed, R., Salmi, A., Butler, L. D., Chiller, J. M. & Oldstone, M. B. Selection of genetic variants of lymphocytic choriomeningitis virus in spleens of persistently infected mice. Role in suppression of cytotoxic T lymphocyte response and viral persistence. J. Exp. Med. 160, 521–540 (1984).
Borrow, P., Evans, C. F. & Oldstone, M. B. Virus-induced immunosuppression: immune system-mediated destruction of virus-infected dendritic cells results in generalized immune suppression. J. Virol. 69, 1059–1070 (1995).
Matloubian, M., Somasundaram, T., Kolhekar, S. R., Selvakumar, R. & Ahmed, R. Genetic basis of viral persistence: single amino acid change in the viral glycoprotein affects ability of lymphocytic choriomeningitis virus to persist in adult mice. J. Exp. Med. 172, 1043–1048 (1990).
Salvato, M., Borrow, P., Shimomaye, E. & Oldstone, M. B. Molecular basis of viral persistence: a single amino acid change in the glycoprotein of lymphocytic choriomeningitis virus is associated with suppression of the antiviral cytotoxic T-lymphocyte response and establishment of persistence. J. Virol. 65, 1863–1869 (1991).
Traub, E. Persistence of lymphocytic choriomeningitis virus in immune animals and its relation to immunity. J. Exp. Med. 63, 847–861 (1936).
Zajac, A. J. et al. Viral immune evasion due to persistence of activated T cells without effector function. J. Exp. Med. 188, 2205–2213 (1998).
Oldstone, M. B. A. et al. Lymphocytic choriomeningitis virus Clone 13 infection causes either persistence or acute death dependent on IFN-1, cytotoxic T lymphocytes (CTLs), and host genetics. Proc. Natl. Acad. Sci. USA 115, E7814–e7823 (2018).
Schnell, F. J., Sundholm, S., Crumley, S., Iversen, P. L. & Mourich, D. V. Lymphocytic choriomeningitis virus infection in FVB mouse produces hemorrhagic disease. PLoS Pathog. 8, e1003073 (2012).
Sanchez, A. B. & de la Torre, J. C. Rescue of the prototypic Arenavirus LCMV entirely from plasmid. Virology 350, 370–380 (2006).
Robinson, J. E. et al. Most neutralizing human monoclonal antibodies target novel epitopes requiring both Lassa virus glycoprotein subunits. Nat. Commun. 7, 11544 (2016).
Brooks, D. G. et al. Interleukin-10 determines viral clearance or persistence in vivo. Nat. Med. 12, 1301–1309 (2006).
Danyukova, T., Schöneck, K. & Pohl, S. Site-1 and site-2 proteases: a team of two in regulated proteolysis. Biochim. Biophys. Acta Mol. Cell Res. 1869, 119138 (2022).
Velho, R. V. et al. Site-1 protease and lysosomal homeostasis. Biochim. Biophys. Acta Mol. Cell Res. 1864, 2162–2168 (2017).
Ohnishi, Y. et al. A furin-defective cell line is able to process correctly the gp160 of human immunodeficiency virus type 1. J. Virol. 68, 4075–4079 (1994).
Chiron, M. F., Fryling, C. M. & FitzGerald, D. Furin-mediated cleavage of Pseudomonas exotoxin-derived chimeric toxins. J. Biol. Chem. 272, 31707–31711 (1997).
Takahashi, S. et al. A second mutant allele of furin in the processing-incompetent cell line, LoVo. Evidence for involvement of the homo B domain in autocatalytic activation. J. Biol. Chem. 270, 26565–26569 (1995).
Sullivan, B. M., Teijaro, J. R., de la Torre, J. C. & Oldstone, M. B. Early virus-host interactions dictate the course of a persistent infection. PLoS Pathog. 11, e1004588 (2015).
Ng, C. T. et al. Blockade of interferon Beta, but not interferon alpha, signaling controls persistent viral infection. Cell Host Microbe 17, 653–661 (2015).
Sheehan, K. C. et al. Blocking monoclonal antibodies specific for mouse IFN-alpha/beta receptor subunit 1 (IFNAR-1) from mice immunized by in vivo hydrodynamic transfection. J. Interferon Cytokine Res. 26, 804–819 (2006).
Teijaro, J. R. et al. Persistent LCMV infection is controlled by blockade of type I interferon signaling. Science 340, 207–211 (2013).
Mody, C. H., Chen, G. H., Jackson, C., Curtis, J. L. & Toews, G. B. Depletion of murine CD8+ T cells in vivo decreases pulmonary clearance of a moderately virulent strain of Cryptococcus neoformans. J. Lab Clin. Med. 121, 765–773 (1993).
Mody, C. H., Chen, G.-H., Jackson, C., Curtis, J. L. & Toews, G. B. Un vivo depletion of murine CD8 positive T cells impairs survival during infection with a highly virulent strain ofCryptococcus neoformans. Mycopathologia 125, 7–17 (1994).
Matloubian, M., Concepcion, R. J. & Ahmed, R. CD4+ T cells are required to sustain CD8+ cytotoxic T-cell responses during chronic viral infection. J. Virol. 68, 8056–8063 (1994).
Zak, J. et al. JAK inhibition enhances checkpoint blockade immunotherapy in patients with Hodgkin lymphoma. Science 384, eade8520 (2024).
Wherry, E. J., Blattman, J. N., Murali-Krishna, K., van der Most, R. & Ahmed, R. Viral persistence alters CD8 T-cell immunodominance and tissue distribution and results in distinct stages of functional impairment. J. Virol. 77, 4911–4927 (2003).
Wherry, E. J. et al. Molecular signature of CD8+ T cell exhaustion during chronic viral infection. Immunity 27, 670–684 (2007).
Mueller, S. N. & Ahmed, R. High antigen levels are the cause of T cell exhaustion during chronic viral infection. Proc. Natl. Acad. Sci. USA 106, 8623–8628 (2009).
Zhou, X., Ramachandran, S., Mann, M. & Popkin, D. L. Role of lymphocytic choriomeningitis virus (LCMV) in understanding viral immunology: past, present and future. Viruses 4, 2650–2669 (2012).
Sandu, I., Cerletti, D., Claassen, M. & Oxenius, A. Exhausted CD8(+) T cells exhibit low and strongly inhibited TCR signaling during chronic LCMV infection. Nat. Commun. 11, 4454 (2020).
Utzschneider, D. T. et al. High antigen levels induce an exhausted phenotype in a chronic infection without impairing T cell expansion and survival. J. Exp. Med. 213, 1819–1834 (2016).
Sandu, I. et al. Landscape of exhausted virus-specific CD8 T cells in chronic LCMV infection. Cell Rep. 32, 108078 (2020).
Wolfe, T. et al. Reduction of antiviral CD8 lymphocytes in vivo with dendritic cells expressing Fas ligand-increased survival of viral (lymphocytic choriomeningitis virus) central nervous system infection. J. Immunol. 169, 4867–4872 (2002).
Kang, S. S. & McGavern, D. B. Lymphocytic choriomeningitis infection of the central nervous system. Front. Biosci. 13, 4529–4543 (2008).
Matullo, C. M., O’Regan, K. J., Hensley, H., Curtis, M. & Rall, G. F. Lymphocytic choriomeningitis virus-induced mortality in mice is triggered by edema and brain herniation. J. Virol. 84, 312–320 (2010).
Bergthaler, A. et al. Viral replicative capacity is the primary determinant of lymphocytic choriomeningitis virus persistence and immunosuppression. Proc. Natl. Acad. Sci. USA 107, 21641–21646 (2010).
Kober, D. L. et al. Identification of a degradation signal at the carboxy terminus of SREBP2: a new role for this domain in cholesterol homeostasis. Proc. Natl. Acad. Sci. USA 117, 28080–28091 (2020).
Smelt, S. C. et al. Differences in affinity of binding of lymphocytic choriomeningitis virus strains to the cellular receptor alpha-dystroglycan correlate with viral tropism and disease kinetics. J. Virol. 75, 448–457 (2001).
Andersen, K. G. et al. Clinical sequencing uncovers origins and evolution of Lassa virus. Cell 162, 738–750 (2015).
Urata, S. et al. Antiviral activity of a small-molecule inhibitor of arenavirus glycoprotein processing by the cellular site 1 protease. J. Virol. 85, 795–803 (2011).
Pasquato, A. et al. Evaluation of the anti-arenaviral activity of the subtilisin kexin isozyme-1/site-1 protease inhibitor PF-429242. Virology 423, 14–22 (2012).
Raini, S. K. et al. The novel therapeutic target and inhibitory effects of PF-429242 against Zika virus infection. Antivir. Res. 192, 105121 (2021).
Blanchet, M., Sureau, C., Guévin, C., Seidah, N. G. & Labonté, P. SKI-1/S1P inhibitor PF-429242 impairs the onset of HCV infection. Antivir. Res. 115, 94–104 (2015).
Wang, T. B. et al. SREBP1 site 1 protease inhibitor PF-429242 suppresses renal cell carcinoma cell growth. Cell Death Dis. 12, 717 (2021).
Uchida, L. et al. Suppressive effects of the site 1 protease (S1P) inhibitor, PF-429242, on Dengue virus propagation. Viruses 8, 46 (2016).
Cai, Y. et al. A Lassa virus live-attenuated vaccine candidate based on rearrangement of the intergenic region. mBio https://doi.org/10.1128/mBio.00186-20 (2020).
Iwasaki, M., Cubitt, B., Sullivan, B. M. & de la Torre, J. C. The high degree of sequence plasticity of the arenavirus noncoding intergenic region (IGR) enables the use of a nonviral universal synthetic IGR to attenuate arenaviruses. J. Virol. 90, 3187–3197 (2016).
Iwasaki, M., Ngo, N., Cubitt, B., Teijaro, J. R. & de la Torre, J. C. General molecular strategy for development of arenavirus live-attenuated vaccines. J. Virol. 89, 12166–12177 (2015).
Cai, Y. et al. A Lassa fever live-attenuated vaccine based on codon deoptimization of the viral glycoprotein gene. mBio https://doi.org/10.1128/mBio.00039-20 (2020).
Cooper, C. L. et al. Preclinical development of a replication-competent vesicular stomatitis virus-based Lassa virus vaccine candidate advanced into human clinical trials. EBioMedicine 114, 105647 (2025).
Geisbert, T. W. et al. Development of a new vaccine for the prevention of Lassa fever. PLoS Med. 2, e183 (2005).
Garbutt, M. et al. Properties of replication-competent vesicular stomatitis virus vectors expressing glycoproteins of filoviruses and arenaviruses. J. Virol. 78, 5458–5465 (2004).
Cross, R. W. et al. A recombinant VSV-vectored vaccine rapidly protects nonhuman primates against heterologous lethal Lassa fever. Cell Rep. 40, 111094 (2022).
Marzi, A., Feldmann, F., Geisbert, T. W., Feldmann, H. & Safronetz, D. Vesicular stomatitis virus-based vaccines against Lassa and Ebola viruses. Emerg. Infect. Dis. 21, 305–307 (2015).
Safronetz, D. et al. A recombinant vesicular stomatitis virus-based Lassa fever vaccine protects guinea pigs and macaques against challenge with geographically and genetically distinct Lassa viruses. PLoS Negl. Trop. Dis. 9, e0003736 (2015).
Acknowledgements
This research was funded by NIH/NIAID grants RO1 AI125626 “A Virus-Free Cell Platform to Discover Inhibitors of Lassa Fever Virus” and R21 AI128556 “Modulation of Lassa Virus vRNP Activity by Host Cell Factors” to J.C.T., and NIH/NIAID grant RO1 AI164744 “The role of IL27 in sustaining the exhausted CD8 T cell response to persistent infection and cancer” to J.R.T.
Author information
Authors and Affiliations
Contributions
R.Z., J.R.T., and J.C.T. conceptualized the work and wrote the main manuscript text. R.Z. prepared all figures. R.Z., H.W., J.R.T. and J.C.T. designed the experiments. R.Z. and M.B. performed the in vivo experiments. R.Z. and T.A. performed tissue sectioning and staining. R.Z., H.W., R.Y.S., and B.C. performed in vitro experiments, data collection and analysis. R.Z. and A.S.K. produced human monoclonal antibodies. M.B. and K.L.M. maintained the mice colonies. J.R.T. and J.C.T. provided supervision, project administration, and funding acquisition. All authors edited and approved the manuscript.
Corresponding authors
Ethics declarations
Competing interests
The authors declare no competing interests.
Additional information
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.
About this article
Cite this article
Zhou, R., Witwit, H., Ai, T. et al. Site-1 protease mediated GPC processing is required for persistence of LCMV Clone 13. npj Viruses 4, 18 (2026). https://doi.org/10.1038/s44298-026-00184-7
Received:
Accepted:
Published:
Version of record:
DOI: https://doi.org/10.1038/s44298-026-00184-7
