ABSTRACT
Several host and viral processes contribute to forming infectious virions. Polyamines are small host molecules that play diverse roles in viral replication. We previously demonstrated that polyamines are crucial for RNA viruses; however, the mechanisms by which polyamines function remain unknown. Here, we investigated the role of polyamines in the replication of the bunyaviruses Rift Valley fever virus (vaccine strain MP-12) and La Crosse virus (LACV). We found that polyamine depletion did not impact viral RNA or protein accumulation, despite significant decreases in titer. Viral particles demonstrated no change in morphology, size, or density. Thus, polyamine depletion promotes the formation of noninfectious particles. These particles interfere with virus replication and stimulate innate immune responses. We extended this phenotype to Zika virus; however, coxsackievirus did not similarly produce noninfectious particles. In sum, polyamine depletion results in the accumulation of noninfectious particles that interfere with replication and stimulate immune signaling, with important implications for targeting polyamines therapeutically, as well as for vaccine strategies.
IMPORTANCE Bunyaviruses are emerging viral pathogens that cause encephalitis, hemorrhagic fevers, and meningitis. We have uncovered that diverse bunyaviruses require polyamines for productive infection. Polyamines are small, positively charged host-derived molecules that play diverse roles in human cells and in infection. In polyamine-depleted cells, bunyaviruses produce an overabundance of noninfectious particles that are indistinguishable from infectious particles. However, these particles interfere with productive infection and stimulate antiviral signaling pathways. We further find that additional enveloped viruses are similarly sensitive to polyamine depletion but that a nonenveloped enterovirus is not. We posit that polyamines are required to maintain bunyavirus infectivity and that polyamine depletion results in the accumulation of interfering noninfectious particles that limit infectivity. These results highlight a novel means by which bunyaviruses use polyamines for replication and suggest promising means to target host polyamines to reduce virus replication.
INTRODUCTION
Bunyaviruses such as Rift Valley fever virus (RVFV) and La Crosse virus (LACV) are emerging arthropod-borne pathogens that cause significant disease. RVFV, which is currently restricted to Africa and the Middle East, can cause encephalitis and hemorrhagic fever in humans, as well as spontaneous abortion in domesticated animals (1). Thus, RVFV infection has significant economic and human health costs. Distantly related to RVFV, LACV can cause encephalitis following infection transmitted by mosquito bite and is localized primarily to midwestern and mid-Atlantic states of the United States (2, 3). These as well as other bunyaviruses pose significant threats to human health. Further, the rapid dissemination of several other arthropod-borne viruses (arboviruses), such as chikungunya virus (4) and Zika virus (ZIKV) (5), demonstrates the epidemic potential of arboviruses, including bunyaviruses.
To remain pathogenic, viruses must maintain infectivity following successful replication of their viral genomes. Several factors contribute to the infectivity of progeny bunyaviruses, including genome integrity (6), inclusion of replicative proteins (polymerase L and nucleoprotein N), and successful envelopment. The proportion of infectious virions versus noninfectious particles in RNA virus infection is low, with noninfectious particles outnumbering infectious particles by 100- to 1,000-fold for Bunyamwera virus, a related bunyavirus (7). The factors contributing to this ratio are multiple but have important implications for infection and pathogenesis (8). The noninfectious particles can serve as decoys from immune system neutralization, enhancing infection by infectious particles (9, 10). In contrast, noninfectious particles can interfere with productive infection, by binding cellular receptors or usurping cellular and viral machinery from infectious viruses (11). Defective genomes play a role in pathogenesis; for example, paramyxoviruses persist in their hosts by maintaining a balance of replication and apoptosis, which is modulated by defective viruses (12). Additionally, arthropod-borne viruses persist in their hosts at least partially through defective viral genomes feeding into the RNA interference (RNAi) pathway (13). The precise mechanisms by which viruses produce defective genomes remains to be understood for several viral families. Importantly, the cellular contribution to this ratio of infectious virus to noninfectious virus is unclear.
Polyamines are small, positively charged molecules found abundantly in all cells. Mammalian cells exclusively synthesize the biogenic polyamines, putrescine, spermidine, and spermine, from an ornithine precursor through ornithine decarboxylase (ODC1). These polyamines are important for cellular processes, such as cell cycling, nucleic acid binding, and altering membrane fluidity (14–16). Importantly, an FDA-approved drug, difluormethylornithine (DFMO), is a specific, nontoxic inhibitor of ODC1, which reduces cellular or organismal polyamine levels. Interestingly, despite a reliance on polyamines for cellular growth and division, polyamine-depleted mammalian cells maintain viability without significant toxicity (17, 18).
We demonstrated that RNA viruses rely on polyamines for replication. Initially, we found that the positive-stranded RNA viruses chikungunya virus and Zika virus required polyamines for translation of the viral polyprotein as well as viral RNA-dependent RNA polymerase activity (19). Olsen et al. uncovered that Ebola virus relies on polyamines for translation of viral proteins through the spermidine metabolite hypusine (20, 21). We further showed that a range of RNA viruses rely on polyamines for replication, both in vitro and in vivo (22). Prior work has established a role for polyamines in virion packaging of nucleic acid in several DNA viruses (23, 24). RNA viruses, in contrast, were found to package negligible levels of polyamines (25). Currently, little is understood about polyamines in the viral life cycle of bunyaviruses such as RVFV or LACV.
To test whether polyamines play a role in the generation of infectious phlebovirus, we measured the ratio of infectious viruses to noninfectious viruses in RVFV infection under conditions of polyamine depletion. We observed that RVFV strain MP-12 viral titers were sensitive to polyamine depletion; however, the relative abundance of viral genomes and proteins was unchanged. We observed that upwards of 100-fold more genomes were required per infectious virus with polyamine depletion. Additionally, viral protein levels and virion abundance, size, and buoyancy properties were unchanged. These noninfectious viruses interfered with the replication of infectious virus and stimulated innate immune responses significantly more than infectious virus. We observed a similar phenotype for Zika virus and Keystone virus (KEYV), two enveloped viruses, but not for coxsackievirus B3 (CVB3), a nonenveloped enterovirus. These results suggest that polyamines contribute to the infectivity of these viruses, uncovering a novel proviral role of polyamines during infection.
RESULTS
Bunyaviruses are sensitive to polyamine depletion.Our previous data focused on the role of polyamines in alphavirus and flavivirus replication (19). We additionally demonstrated that polyamines are required for a number of distinct viral families (22). We understand little about potential roles for polyamines in these diverse viruses. To initially examine polyamines in bunyavirus replication, we sought to determine sensitivity to polyamine depletion. DFMO, an ODC1 inhibitor, at a concentration of 500 μM was sufficient to block alpha- and flavivirus replication; thus, we investigated whether these parameters would similarly reduce MP-12 infection. We treated Huh7 cells with 500 μM DFMO for 4 days prior to infection at a multiplicity of infection (MOI) of 0.01 PFU per cell. Samples were collected every 8 h, and titers were determined on Vero-E6 cells. We observed characteristic sigmoidal growth kinetics of untreated cells; however, DFMO-treated cells failed to produce virus (Fig. 1A). In fact, viral titers remained flat over 64 h of infection, suggesting that MP-12 failed to replicate without polyamines.
Rift Valley fever virus MP-12 and La Crosse virus are sensitive to polyamine depletion. (A) Huh7 cells were left untreated or were treated with 500 μM DFMO for 4 days prior to infection at a multiplicity of infection (MOI) of 0.01 PFU of RVFV MP-12 strain per cell. Samples were collected every 8 h for 64 h, and the titer was subsequently determined via plaque assay. (B) Huh7 cells were treated with escalating doses of DFMO as indicated and subsequently infected with MP-12 at an MOI of 0.1 PFU per cell. Viral titers were determined at 48 h postinfection. (C) Intracellular polyamine levels were measured using a dual-luciferase reporter assay of an OAZ1 transcript construct following DFMO treatment for 4 days. Relative luciferase activity was normalized to untreated samples. (D) Thin-layer chromatography on cells treated as for panel B to measure biogenic polyamine levels following DFMO treatment. (E) Huh7 cells were treated with 100 μM DENSpm for 16 h prior to infection with MP-12 at an MOI of 0.01. Samples were collected every 8 h for 32 h, and the titer was determined via plaque assay. (F) Huh7 cells were treated with escalating doses of DENSpm as indicated and infected with MP-12 at an MOI of 0.01 PFU per cell. Viral titers were determined at 48 hpi. (G and H) Relative polyamine levels (G) as measured by thin-layer chromatography (H) were quantified via ImageJ analysis and normalized to untreated controls. (I) Huh7 cells were treated with escalating doses of exogenous polyamines by adding a polyamine mixture to cellular supernatant. Cells were infected at an MOI of 0.1 with MP-12, and viral titers were determined at 24 hpi via plaque assay. (J and K) Huh7 cells treated with DFMO (J) or DENSpm (K) were treated as described above before infection with LACV at an MOI of 0.1. Titers were determined by plaque assay at 24 hpi. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001 (using Student’s t test [n ≥ 3], comparing untreated cells to DFMO treatment). Error bars represent ±1 standard error of the mean (SEM). Statistical comparisons were performed between treated and untreated conditions. NT, not treated.
To measure sensitivity to DFMO and, thereby, polyamine depletion, we treated Huh7 cells with increasing doses of DFMO from 100 μM to 5 mM and measured virus replication at 48 h postinfection (hpi). We observed a significant decrease in titer when cells were treated with a minimum of 500 μM DFMO (Fig. 1B). Both 1 mM and 5 mM treatments significantly reduced titers as well. We verified that DFMO was depleting polyamines by using a quantitative polyamine-sensitive luciferase assay (26), which demonstrated significant reductions in polyamine levels at all tested concentrations (Fig. 1C). To confirm that these doses of DFMO were, in fact, depleting polyamines, we performed thin-layer chromatography (TLC) on polyamines. We found that, compared to untreated samples, DFMO treatment as low as 100 μM reduced polyamine levels (Fig. 1D).
Several molecules have been developed to target the polyamine pathway. We next investigated whether MP-12 was sensitive to another polyamine depleting pharmaceutical, N1,N11-diethylnorspermine (DENSpm). First, we determined a growth curve of MP-12 using 100 μM DENSpm, an activator of SAT1 which acetylates polyamines, leading to their neutralization and export from the cell (27, 28). We observed significant decreases in viral titers over a course of infection (Fig. 1E). Using a range of DENSpm concentrations, we observed a decrease in MP-12 titers at concentrations greater than 50 μM (Fig. 1F), similar to our data with other viruses (22). We confirmed via TLC and ImageJ quantitation of chromatograms that DENSpm reduced polyamine levels (Fig. 1G and H).
Because DFMO depletes polyamines and reduces viral titers, we investigated whether polyamines could stimulate bunyavirus titers. Huh7 cells were treated with increasing doses of a polyamine mixture, from 1× to 10×, for 2 h prior to infection with MP-12. Viral titers were measured at 48 hpi, and we observed no significant change, even at the highest dose of polyamines (Fig. 1I). These data suggest that exogenous polyamines cannot stimulate virus replication above basal levels, perhaps because virus replication requires a threshold level of polyamines and does not proportionately respond to additional polyamines.
Finally, we extended this phenotype to La Crosse virus (LACV), a distantly related bunyavirus. We similarly treated cells with increasing doses of DFMO and measured titers at 24 hpi. We noted a significant decrease in viral titers at doses greater than 100 μM DFMO (Fig. 1J). LACV was also sensitive to DENSpm (Fig. 1K). These data suggest that sensitivity to polyamine depletion is shared between these two bunyaviruses, as well as a wide array of RNA viruses (22).
The genome-to-PFU ratio is increased with polyamine depletion.Reports have suggested that polyamines are present in the virions of diverse viruses and promote genome packaging (23, 24). We hypothesized that polyamines may function similarly for bunyavirus infection. To test this hypothesis, we aimed to quantify the ratio of the number of genomes per virus in infected cell supernatant. To this end, we treated Huh7 cells with DFMO and subsequently infected them with MP-12 at an MOI of 0.1 for 48 h. We determined the titer of the virus and measured the number of genomes (small [S], medium [M], and large [L] segments) via reverse transcription-quantitative PCR (qRT-PCR) with genome-specific primers. As in Fig. 1B, we observed reduced MP-12 titers with DFMO treatment (Fig. 2A). We anticipated that if polyamines were involved in packaging, we would observe significantly fewer viral genomes in the infected-cell supernatant. Surprisingly, we observed equivalent numbers of genomes in the supernatant with escalating doses of DFMO (Fig. 2B). We next calculated the genome-to-PFU ratio by dividing the number of viral genomes by the titers (values in Fig. 2B divided by those in Fig. 2A). We found that there was a dramatic increase in the number of viral genomes per infectious virus in a DFMO dose-dependent manner (Fig. 2C). We confirmed these phenotypes by depleting cells with DENSpm. We treated cells with 10 or 100 μM DENSpm for 16 h prior to infection at an MOI of 0.1. We collected the virus, determined its titer (Fig. 2D), and measured genomes (Fig. 2E) at 48 hpi. As with DFMO, MP-12 titers were decreased but viral genome levels were unchanged, resulting in a high genome-to-PFU ratio (Fig. 2F).
Specific infectivity of RVFV MP-12 is diminished with polyamine depletion. (A to C) Huh7 cells were treated with either 500 μM or 1 mM DFMO for 4 days prior to infection with MP-12 at an MOI of 0.1 PFU per cell. (A and B) Viral titers were determined via plaque assay (A) and viral genomes in culture supernatant were quantified via qRT-PCR (B) at 48 hpi. (C) The relative number of viral genomes from panel B was divided by the titer from panel A to determine the genome-to-PFU ratio, normalized to untreated controls. (D to F) Huh7 cells were treated with 10 or 100 μM DENSpm for 16 h prior to infection with MP-12 at an MOI of 0.1 PFU per cell. (D and E) Viral titers were determined via plaque assay (D) and viral genomes quantified via qRT-PCR (E) at 48 hpi. (F) The ratio of viral genomes (E) was divided by the titer (D) to determine the genome-to-PFU ratio, normalized to untreated controls. (G) Huh7 cells were treated and infected as for panel A, and the number of cell-associated viral genomes was determined via qRT-PCR, normalizing to cellular GAPDH. (H and I) Samples from panels B and E were treated with RNase prior to genome quantification via qRT-PCR and divided by viral titers from panels A and D to determine the genome-to-PFU ratio. Values above data bars represent the fold change compared to untreated conditions. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001 (using Student’s t test [n ≥ 3], comparing DFMO treatment to untreated controls). Error bars represent ±1 SEM. Statistical comparisons were performed between treated and untreated conditions.
We next measured cell-associated viral RNA by extracting RNA from attached, infected cells. As with supernatant RNA, we observed no change in the number of genomes produced (Fig. 2G). However, the MP-12 genomic RNA measurements could be free RNA or virion-associated RNA. To distinguish between the two, we treated our infected-cell supernatant with RNase A prior to genome quantification. We observed no change in the genome-to-PFU ratio (Fig. 2H), suggesting that a portion of supernatant genomic RNA was protected from RNase degradation. We presume that these RNase-protected genomes are virion associated; however, these data do not exclude the possibility that viral RNA could be contained in a vesicle or other layer to protect from degradation. This phenotype was confirmed again with DENSpm treatment (Fig. 2I). Together, these data suggest that polyamine-depleted cells have no defect in viral genome production or export.
Finally, we investigated these results with LACV. DFMO-treated cells were infected at an MOI of 0.1 and supernatant collected at 24 hpi for titration and genome quantitation. We observed the same phenotype for LACV as for MP-12: viral titers were reduced, but genome levels were not significantly decreased, with either DFMO (Fig. 3A) or DENSpm (Fig. 3B). Thus, the genome-to-PFU ratio was escalated with polyamine depletion for an additional bunyavirus.
The LACV genome/PFU ratio is altered with polyamine depletion. Huh7 cells were treated with increasing doses of DFMO for 4 days prior to infection at an MOI of 0.1 PFU/cell with LACV. Viral titers and genomes were quantified via plaque assay and qRT-PCR, respectively, at 24 hpi. The genome/titer ratio was calculated by dividing the number of genomes by the viral titer without (A) and with (B) RNase treatment. Error bars represent ±1 SEM (n ≥ 3). Statistical comparisons were performed between treated and untreated conditions.
Viral protein levels are unchanged in infected-cell supernatant.We observed that viral titers are reduced greater than 100-fold with DFMO treatment (Fig. 1A and B). However, genome levels are unchanged relative to those in the control (Fig. 2B). We hypothesized that this discrepancy could be due to aberrant virion protein content. To test this hypothesis, we measured the levels of nucleoprotein (N) and glycoprotein N (GN) in both cells and infected-cell supernatant. We observed no difference between the levels of proteins in the supernatant (Fig. 4A), though there was a slight decrease in viral protein content when DFMO-treated intact cells were collected (Fig. 4B).
Viral protein levels are unchanged in released virus. Huh7 cells were treated with escalating doses of DFMO or left untreated for 4 days and subsequently infected at an MOI of 0.1 PFU per cell or left uninfected. At 48 hpi, supernatant (A) and cells (B) were collected for Western blot analysis for viral proteins GN and N, as well as cellular GAPDH. (C) Huh7 cells were left untreated or treated with DFMO and infected with MP-12 for 48 h prior to indirect immunofluorescence staining for nucleoprotein (N) (green), double-stranded RNA (dsRNA) (red), actin (purple), and nucleic acid (blue). (D and E) Samples from panels A and B were run on an acrylamide gel and analyzed for total protein content by silver staining for supernatant (D) and cell-associated (E) proteins. **, P ≤ 0.01 using two-way analysis of variance (ANOVA) (n = 2) (untreated to DFMO-treated samples).
Because we observed significant changes in N protein levels in cell-associated protein but not in supernatant (Fig. 4A and B), we investigated whether the N protein distribution changed with DFMO treatment. Toward this end, we imaged the N protein as well as double-stranded RNA (dsRNA) (J2) in infected cells that were either left untreated or treated with 500 μM DFMO. We noted that the N protein localized to perinuclear speckles that colocalized with dsRNA under both conditions (Fig. 4C). No significant detectable difference between treatment conditions was noted, though signal for the N protein was slightly reduced under DFMO treatment conditions, fitting with our observed Western blot results.
As an additional control, we measured global viral and cellular protein levels in infected cells and supernatants via silver-stained gels to determine if a DFMO-modulated nonviral protein may contribute to infectivity. Again, we observed no gross differences in protein profiles on our silver-stained gels, either in cells or in the supernatant (Fig. 4D and E). These data suggest that DFMO does not alter viral protein levels in the infected-cell supernatants.
Viral particles from polyamine-depleted cells show no significant physical differences.We next sought to determine if polyamine depletion had any physical effect on the virions produced from infected cells. First, we generated viruses from untreated and DFMO-treated cells, concentrated and purified virions, and examined the particles by electron microscopy. We observed virions of a size and shape previously described (Fig. 5A) (29), with an average diameter of 90 to 110 nm. Intriguingly, we observed a similar number and size for virions derived from DFMO-treated cells (Fig. 5B), despite a lack of infectivity. These data suggest that fully formed viral particles are being produced but are not infectious.
Physical properties of secreted viral particles are unchanged with polyamine depletion. (A) Huh7 cells were treated with 500 μM DFMO or left untreated and subsequently infected with MP-12 at an MOI of 0.1 PFU per cell for 48 h. Secreted viruses were collected, concentrated, and purified via ultracentrifugation prior to electron microscopy. Representative images are shown for each treatment condition. (B) Diameters of viral particles derived from untreated or DFMO-treated cells as for panel A were measured with ImageJ. NS, not significant. (C to F) MP-12 particles derived as for panel A were subjected to discontinuous sucrose gradient ultracentrifugation, from 20% to 45% sucrose. Fractions were collected and analyzed by silver staining (C), Western blotting for viral N protein (D), qRT-PCR to detect viral genomes (E), and plaque assay to determine viral titers (F). Error bars represent ±1 SEM (n ≥ 3).
To determine whether polyamine depletion altered the buoyancy properties of the MP-12 virions, we employed differential sucrose gradient ultracentrifugation. Using a 20% to 45% discontinuous gradient, we considered total protein content by silver staining the same fractions and observed viral proteins to similar extents under both untreated and DFMO-treated conditions (Fig. 5C), with peaks for these proteins in fractions from 22.5% to 30% sucrose. We observed MP-12 N protein in gradients ranging from 22.5% to 30% sucrose when analyzed by Western blotting, as previously described (30), for virions from both untreated and DFMO-treated cells. (Fig. 5D). Viral genomes measured via qRT-PCR were also not statistically significantly different with polyamine depletion (Fig. 5E). We observed a peak in viral titers in these same fractions, though titers under DFMO treatment conditions were 10- to 100-fold lower (Fig. 5F). In total, these results suggest that DFMO-mediated polyamine depletion does not alter the buoyancy properties or virion physical size, despite significant reductions in infectivity.
Viral particles from polyamine-depleted cells interfere with productive viral infection.Given the similar observed physical properties of virions from untreated and polyamine-depleted cells, we hypothesized that these noninfectious particles could interfere with productive virus infection, akin to defective interfering viral particles. To examine this, we combined 104 PFU of our stock virus, produced on Huh7 cells without any drug treatment, with equal numbers of infectious virus (104 PFU) from untreated and DFMO-treated cells (virus passaged once). At 48 hpi, we collected and determined the titer of virus in the supernatant via plaque assay. We detected a modest yet repeatable and statistically significant decrease in virus production when stock was combined with virus from DFMO-treated cells (Fig. 6A). Interestingly, this phenotype was dose dependent and correlated with our data in Fig. 2C. As confirmation of this phenotype, we performed the same assay using LACV and observed similar results (Fig. 6B): viral titers were significantly reduced when stock virus was combined with virus produced from polyamine-depleted cells, suggesting that the noninfectious particles interfere with infectious-virus infection.
Viruses derived from polyamine-depleted cells interfere with productive viral infection. (A) Virus derived from untreated or DFMO-treated cells was added to stock MP-12 by combining equal numbers of PFU of stock virus and passaged virus. These virus combinations were used to infect Huh7 cells. At 48 hpi, the titer of the virus was determined via plaque assay and percent replication calculated by comparing the titer to that for Huh7 cells infected with stock virus alone. (B) Viruses were combined and analyzed as for panel A but with LACV. (C) Vero cells were infected and analyzed as for panel A, and titers were determined after 48 h of infection. (D and E) Huh7 cells were infected with MP-12 (D) and LACV (E) derived from untreated and DFMO-treated cells, and cell-associated RNA was collected at 30 min and 4 h postinfection. RNA was reverse transcribed, and cell-associated viral genomes were analyzed via qPCR. Error bars represent ±1 SEM (n ≥ 3). *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001 (using Student’s t test [n ≥ 3], comparing combination treatment to stock-alone infection).
In addition to Huh7 cells, we investigated whether our MP-12 derived from DFMO-treated cells could interfere with replication of virus stocks on an interferon-null cell type. Using Vero-E6 cells, we combined virus from our stock with virus passaged in untreated or DFMO-treated Huh7 cells, infected Vero-E6 cells for 48 h, and measured titers by plaque assay. Similar to our results with Huh7 cells, we observed a small yet significant reduction in viral titers when stock virus was combined with virus derived from DFMO-treated cells (Fig. 6C), suggesting that the noninfectious particles interfere with infectious virus infection in multiple cell types and, namely, interferon-null Vero cells.
To further characterize that virus derived from untreated and DFMO-treated cells was competent to bind and enter cells, we performed binding and entry assays. After inoculating cells and washing away unbound virus, we measured cell-associated virus at 30 min (bound) and 4 h (entered). We observed that approximately 1% of our inoculum genomes bound to cells after 30 min, with no difference between virus from untreated or DFMO-treated cells. Similarly, no difference was observed at 4 h (Fig. 6D). These data were also obtained for LACV (Fig. 6E). Together, these data suggest that noninfectious virus derived from DFMO-treated cells is able to bind to and enter cells, at a level similar to that for infectious virus.
Viral particles from polyamine-depleted cells activate interferon signaling.We have observed that polyamine depletion, through either DFMO or DENSpm, reduces bunyavirus titers, alters the genome-to-PFU ratio, and generates interfering noninfectious particles. Given that bunyavirus infection initiates innate immune signaling (31–33), we hypothesized that the particles generated during infection of polyamine-depleted cells would stimulate type I interferon signaling and, due to their noninfectious nature, would not be able to counteract interferon signaling. To examine this, we generated phorbol 12-myristate 13-acetate (PMA)-differentiated THP-1 human macrophages, added equal numbers (104 PFU) of infectious viruses from each treatment condition (untreated or DFMO treated), and left cells unstimulated or treated with 1 μg/ml lipopolysaccharide as a positive control. We stimulated for 16 h, collected and purified cellular RNA, reverse transcribed, and measured interferon-stimulated gene (ISG) expression by qRT-PCR, normalizing to GAPDH (glyceraldehyde-3-phosphate dehydrogenase). We observed that MP-12 infection of the differentiated THP-1 cells induced a robust type I interferon response, as measured by Stat1 (Fig. 7A), viperin (Fig. 7B), and IFIT1 (Fig. 7C) expression. When we stimulated the cells with virus derived from DFMO-treated cells, we observed a significantly higher ISG response for all three ISGs. When we infected cells and normalized the number of genomes (consisting of both infectious and noninfectious virus), we observed similar interferon-stimulated gene responses (Fig. 7D), suggesting that the noninfectious-virus components are responsible for the elevated interferon response observed in Fig. 7A to C. Importantly, we observed that DFMO itself did not stimulate or alter an interferon response, as differentiated THP-1 cells treated with DFMO did not exhibit enhanced ISG expression, nor did DFMO treatment change ISG expression when THP-1 cells were treated with interferon beta and DFMO concurrently (Fig. 7E). Finally, we further observed this phenotype in mouse RAW264 macrophages when we measured viperin expression (Fig. 7D). Taken together, these results suggest that the viral particles derived from DFMO-treated cells stimulate innate immune signaling more robustly than virus from cells with polyamines.
Viruses derived from polyamine-depleted cells activate interferon signaling. (A to C) Virus derived from untreated or DFMO-treated Huh7 cells was added to differentiated THP-1 monocytes and left for 16 h; 1 μg/ml lipopolysaccharide (LPS) was added as a control. After 16 h of infection, cells were collected for determination of interferon-stimulated gene expression via qRT-PCR. Cells were probed for Stat1 (A), viperin (B), and IFIT1 (C) expression, normalized to cellular GAPDH. (D) Differentiated THP-1 monocytes were stimulated as for panels A to C but with equal numbers of viral genomes, as quantified by qPCR. IFIT1 expression was measured 16 h later via qRT-PCR. (E) Differentiated THP-1 monocytes were treated with DFMO in combination with interferon beta (IFNβ) and analyzed for ISG15 expression 16 h later via qRT-PCR. (F) RAW264.7 cells were similarly stimulated with virus or LPS and analyzed for viperin expression 16 h later. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001 (using Student’s t test [n ≥ 3], comparing treated samples to unstimulated controls). Error bars represent ±1 SEM.
Polyamines promote infectivity for ZIKV but not an enterovirus.We observed that both the MP-12 strain of RVFV and LACV generate noninfectious viral particles that interfere with productive infection and stimulate innate immune signaling. To determine the breadth of this phenotype, we measured the genome-to-PFU ratios of additional viruses from distinct viral families. Both MP-12 and LACV are negative-sense (partially ambisense) enveloped viruses. As a model positive-sense RNA virus, we used Zika virus (ZIKV) (African strain MR766), and we measured viral titers (Fig. 8A), supernatant genomes (Fig. 8B), and the genome-to-PFU ratio (Fig. 8C) in DFMO-treated cells. Similar to our results with bunyaviruses, we observed sensitivity to DFMO at the level of viral titer but not supernatant genome, and the genome-to-PFU ratio was significantly increased in DFMO-treated cells in a dose-dependent manner. These results suggest that this phenotype is shared with these distinct viral families.
Polyamines enhance specific infectivity of enveloped viruses. (A to C) Huh7 cells were treated with 500 μM or 1 mM DFMO for 4 days prior to infection with ZIKV (strain MR766) at an MOI of 0.1 PFU per cell. At 48 hpi, viral titers were determined via plaque assay (A) and relative viral genomes were determined by qRT-PCR (B), and the ratio of genomes to PFU was calculated (C). (D to F) Vero cells were treated with 500 μM or 1 mM DFMO for 4 days prior to infection with CVB3 (Nancy strain) at an MOI of 0.1 PFU per cell. At 24 hpi, viral titers were determined via plaque assay (D) and relative viral genomes were determined by qRT-PCR (E), and the ratio of genomes to PFU was calculated (F). (G to I) Identical infection and analysis methods were performed for KEYV in Huh7 cells to determine viral titers (G), viral genomes (H), and genome-to-PFU ratio (I) at 48 hpi. (J to L) These methods were also used for VSV (strain Indiana) infected at an MOI of 0.01 PFU per cell for 24 h. Viral titers (J), genomes (K), and genome-to-PFU ratio (L) were calculated. *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001 (using Student’s t test [n ≥ 3]). Error bars represent ±1 SEM. Statistical comparisons were performed between treated and untreated conditions.
We employed coxsackievirus B3 (CVB3) (Nancy strain) as a positive-sense nonenveloped virus. As with ZIKV and the bunyaviruses, we measured viral titers (Fig. 8D), supernatant genomes (Fig. 8E), and genome-to-PFU ratio (Fig. 8F) in DFMO-treated cells. Interestingly, we observed a reduction in titer and a concomitant decrease in viral genome quantity that did not significantly alter the genome-to-PFU ratio. While we had observed a >100-fold increase in ratio for the bunyaviruses and an 18-fold increase for ZIKV, the genome-to-PFU ratio for CVB3 was at most 2.4-fold elevated with DFMO treatment compared to that for the wild type.
Finally, we explored additional negative-sense viruses, including Keystone virus (KEYV), which gained recent notoriety for the first human infection (34). As with MP-12 and LACV, KEYV titers were sensitive to DFMO (Fig. 8G), but genomes were not (Fig. 8H). Thus, the genome-to-PFU ratio (Fig. 8I) for KEYV was elevated in a fashion similar to that for MP-12, LACV, and ZIKV. An unrelated negative-sense RNA virus, vesicular stomatitis virus (VSV) (Indiana strain), was also considered by measuring virus replication and normalizing to viral genomes. We observed that VSV replication was sensitive to polyamine depletion (Fig. 8J) but that the number of viral genomes (Fig. 8K) was unchanged, in a fashion similar to that for ZIKV and bunyaviruses. Thus, the virus-to-genome ratio (Fig. 8L) for VSV was elevated with polyamine depletion. Together, these results suggest that polyamines are crucial for maintaining a genome-to-PFU ratio for a subset of RNA viruses. Altogether, our data suggest that polyamine depletion results in the accumulation of noninfectious particles for several enveloped RNA viruses.
DISCUSSION
Our data suggest that polyamines are crucial for the replication of bunyaviruses, specifically RVFV, LACV, and KEYV, and that polyamines promote infectivity of progeny virions via a to-be-defined mechanism(s). While we previously uncovered a role for polyamines in enzymatic activity of viral enzymes (19), this study is the first to find that polyamines are required for infectivity. Both alphaviruses and flaviviruses were sensitive to polyamine depletion at the level of genome translation and RNA-dependent RNA polymerase activity; however, in those prior studies, we had not investigated whether noninfectious particles were still produced from polyamine-depleted cells. Here, we have generated a working model whereby polyamines facilitate productive bunyavirus infection, and in the absence of polyamines, noninfectious particles accumulate that interfere with replication and stimulate innate immune signaling to quell infection (Fig. 9).
Working model. Cells containing polyamines support robust virus replication. In contrast, infection of polyamine-depleted cells results in the accumulation of noninfectious viral particles that stimulate innate immune responses and directly interfere with productive virus replication. As a result, polyamine depletion limits bunyavirus replication.
Based on these data, as well as our and others’ previous data, polyamines play pleiotropic roles during viral infection (35). Owing to their nature as small, abundant, positively charged molecules, it is not surprising that they affect virus infection at distinct stages. Here, we find that for bunyaviruses, polyamines are important for infectivity: with polyamine depletion, infected cells produce and export viral proteins and genomes, but the produced virions are noninfectious. Interestingly, this phenotype did not hold for the nonenveloped CVB3, a positive-sense RNA virus. Enteroviruses like CVB3 are known to recombine to rapidly evolve (36), and this propensity to recombine may preclude changes in the ratio of infectious to noninfectious virus. Whether polyamines affect viral membranes to promote infectivity remains unclear, but this defining feature of these distinct phenotypes merits investigation.
Previous work detected polyamines in several viruses (23, 24), while several RNA viruses investigated contained small amounts of polyamines (37, 38). In the case of herpesviruses, the viral genomes associate with histones in the nucleus (39–41) but are neutralized by polyamines in the capsid. It is unclear whether this phenotype is a function of genome size and the need to neutralize a large amount of negatively charged nucleic acid, versus the case for a relatively smaller single-stranded RNA virus genome. Additionally, RNA viruses have evolved distinct packaging mechanisms that are not dependent on polyamines. The presence of polyamines in bunyavirus virions (or flavivirus or alphavirus virions) remains unexplored. Whether virion-bound polyamines affect infectivity or virion stability is an important area of future research.
Defective viral particles have received increasing attention, and their roles in viral infection are becoming more appreciated. Due to their inability to replicate independently, defective viral particles usurp infectious viral (and cellular) machinery, reducing the capacity of the infectious virus to replicate (42). Several mechanisms have been described whereby RNA virus genomes become defective, via error-prone polymerases, unique RNA structures, or mistakes during viral genome replication (11). While attention has focused on how defective genomes attenuate the infectivity of viral particles, how cellular factors contribute to defective virus production is less clear. Our results suggest that polyamine depletion results in noninfectious virus generation but with fully intact virions.
We previously uncovered a novel role for polyamines in the innate immune response via the polyamine-acetylating enzyme SAT1 (19). Upon interferon stimulation, SAT1 is induced and reduces polyamine pools within cells. Whether interferon-mediated polyamine depletion plays a role in generating defective virions from infection is an important hypothesis for future testing. A tantalizing model could be that innate immune responses trigger polyamine depletion to limit virus infection and generate defective viruses that could amplify innate immunity and initiate adaptive immunity. Further, the ability to target polyamines pharmacologically suggests that we may be able to tip this balance between virus infection and host response to enhance noninfectious virion production and, consequently, innate immune signaling. Pharmacologically, targeting polyamines is achievable, as DFMO is approved for treatment of trypanosomiasis (43, 44), and several clinical trials are exploring the potential of DFMO as an anticancer therapeutic. DFMO is also used topically to reduce facial hair growth (45). Interestingly, humans can tolerate large doses of DFMO over long periods of time. Major side effects, including thrombocytopenia and ototoxicity, reverse after treatment completion. Whether DFMO (or other polyamine-targeting molecules) could be used clinically to quell RNA virus replication remains to be understood. However, our data suggest that the polyamine pathway is central to virus replication and that reducing polyamine pools may effectively limit infection. Our results also suggest novel strategies to attenuate viruses: polyamine-depleted cells produce fully assembled viruses that are noninfectious. Such an attenuation strategy may serve well to stimulate immune responses, which may function to enhance vaccine efficacy. While significant work remains, understanding how polyamines contribute to virus infectivity can provide insight into basic mechanisms of virus infectivity and highlight novel therapeutic routes.
MATERIALS AND METHODS
Cell culture.Cells were maintained at 37°C and 5% CO2 in Dulbecco’s modified Eagle’s medium (DMEM) (Life Technologies) with bovine serum and penicillin-streptomycin. Vero cells (BEI Resources, NR-10385) were supplemented with 10% newborn calf serum (NBCS) (Thermo Fisher), and Huh7 cells, kindly provided by Susan Uprichard, were supplemented with 10% fetal bovine serum (FBS) (Thermo Fisher). THP-1 cells were donated by Makio Iwashima and maintained in RPMI (Thermo Fisher) supplemented with 2% FBS and beta-mercaptoethanol (BME) (50 mM; Thermo Fisher). THP-1 cells were differentiated with phorbol 12-myristate 13-acetate (PMA) (100 pg/ml; Thermo Fisher). RAW264.1 macrophages were provided by Katherine Knight and maintained in 2% FBS.
Drug treatment.Difluoromethylornithine (DFMO) (TargetMol) and N1,N11-diethylnorspermine (DENSpm) (Santa Cruz Biotechnology) were diluted to 100× solutions (100 mM and 10 mM, respectively) in sterile water. For DFMO treatments, cells were trypsinized (VWR) and reseeded with fresh medium supplemented with 2% serum. Following overnight attachment, cells were treated with 100 μM, 500 μM, 1 mM, or 5 mM DFMO. Cells were incubated with DFMO for 96 h to allow for depletion of polyamines in Huh7 cells. For DENSpm treatment, cells were treated with 100 nM, 1 μM, 10 μM, 100 μM, and 1 mM DENSpm 16 h prior to infection. During infection, medium was cleared and saved from the cells. The same medium containing DFMO and DENSpm was then used to replenish the cells following infection. Cells were incubated at the appropriate temperature for the duration of the infection.
Infection and enumeration of viral titers.MP-12 (30), LACV, and KEYV were derived from the first passage of virus in Huh7 cells. ZIKV (MR766) was derived from the first passage of virus in Vero cells. CVB3 (Nancy strain) was derived from the first passage of virus in HeLa cells. ZIKV, LACV, and, KEYV were obtained from Biodefense and Emerging Infections (BEI) Research Resources. For all infections, DFMO and DENSpm were maintained throughout infection as designated. Viral stocks were maintained at −80°C. For infection, virus was diluted in serum-free DMEM for a multiplicity of infection (MOI) of 0.1 on Huh7 cells, unless otherwise indicated. Viral inoculum was overlain on cells for 10 to 30 min, and the cells were washed with phosphate-buffered saline (PBS) before replenishment of medium. Supernatants were collected from MP-12, LACV, KEYV, ZIKV, and CVB3 as indicated. Dilutions of cell supernatant were prepared in serum-free DMEM and used to inoculate confluent monolayers of Vero cells for 10 to 15 min at 37°C. Cells were overlain with 0.8% agarose in DMEM containing 2% NBCS. CVB3 samples incubated for 2 days, MP-12, ZIKV, and LACV samples incubated for 3 days, and KEYV samples incubated for 5 days at 37°C. Following appropriate incubation, cells were fixed with 4% formalin and revealed with crystal violet solution (10% crystal violet; Sigma-Aldrich). Plaques were enumerated and used to back-calculate the number of PFU per milliliter of collected volume.
TLC determination of polyamines.Polyamines were separated by thin-layer chromatography (TLC) as previously described (46). For all samples, cells were treated as described prior to being trypsinized and centrifuged. Pellets were washed with PBS and then resuspended in 200 μl 2% perchloric acid. Samples were then incubated overnight at 4°C, and then 200 μl of supernatant was combined with 200 μl 5-mg/ml dansyl chloride (Sigma-Aldrich) in acetone and 100 μl saturated sodium bicarbonate. Samples were incubated in the dark overnight at room temperature. Excess dansyl chloride was cleared by incubating the reaction mixture with 100 μl 150-mg/ml proline (Sigma-Aldrich). Dansylated polyamines were extracted with 50 μl toluene (Sigma-Aldrich) and centrifuged. Five microliters of sample was added in small spots to the TLC plate (silica gel matrix; Sigma-Aldrich) and exposed to ascending chromatography with 1:1 cyclohexane-ethylacetate. The plate was dried and visualized via exposure to UV.
Polyamine luciferase reporter assay.To measure free polyamine levels in cells, a dual-luciferase vector containing the wild-type −1 frameshift antizyme OAZ1 (pC5730), kindly sent to us by Tom Dever from the National Institutes of Health, was transfected into cells with LipoD293 (SignaGen). Free polyamines modulate OAZ1 mRNA frameshifting, and these constructs can measure relative endogenous polyamine concentrations via a dual-luciferase reporter as previously described (26). Huh7 cells were seeded with 2% medium and drug treated as described above. Cells were transfected with 62.5 ng of reporter plasmid, and after 24 h of incubation, the luminescent signal was measured using the dual-luciferase reporter assay system (Promega) by measuring both firefly and Renilla luciferases with the Veritas microplate luminometer (Turner Biosystems). Firefly luciferase was normalized to Renilla luciferase and the wild-type values and subsequently normalized to untreated controls.
Virus preparation and concentration.MP-12 was inoculated onto confluent Huh7 cells at an MOI of 0.001 and incubated for 48 to 72 h, until full cytopathic effect (CPE). The supernatant was clarified via centrifugation and then concentrated using VivaSpin 20 centrifugal concentrators (GE Lifesciences). Virus was concentrated 1,000-fold by resuspending in an appropriate amount of serum-free DMEM.
RNA purification and cDNA synthesis.Medium was cleared from cells and TRIzol reagent (Zymo Research) directly added. Lysate was then collected, and RNA was purified according to the manufacturer’s protocol utilizing the Direct-zol RNA Miniprep Plus kit (Zymo Research). Purified RNA was subsequently used for cDNA synthesis using high-capacity cDNA reverse transcription kits (Thermo Fisher), according to the manufacturer’s protocol, with 10 to 100 ng of RNA and random hexamer primers.
Viral genome quantification.Following cDNA synthesis, qRT-PCR was performed using the QuantStudio3 (Applied Biosystems by Thermo Fisher) and SYBR green master mix (DotScientific). Samples were held at 95°C for 2 min prior to 40 cycles of 95°C for 1 s and 60°C for 30 s. Primers were verified for linearity using 8-fold serially diluted cDNA and checked for specificity via melt curve analysis followed by agarose gel electrophoresis. All samples were normalized to total RNA using the ΔCT method. Primer sequences are shown in Table 1.
Primers used in these studies
Genome-to-PFU ratio calculations.The number of viral genomes, quantified as described above, was divided by the viral titer, as determined by plaque assay, to measure the genome-to-PFU ratio. Values obtained were normalized to untreated conditions to obtain the relative genome-to-PFU ratio.
Western blotting.Samples were collected with Bolt LDS buffer and Bolt reducing agent (Invitrogen) and run on polyacrylamide gels. Gels were transferred using the iBlot 2 Gel Transfer Device (Invitrogen). Membranes were probed with primary antibodies for Rift Valley fever virus nucleoprotein (N) and glycoprotein N (GN) (1:500; BEI Resources) and β-actin (1:5,000; Santa Cruz Biotechnology). Membranes were treated with SuperSignal West Pico Plus chemiluminescent substrate (Thermo Fisher Scientific) and visualized on a ProteinSimple FluorChem E imager.
Silver staining.Samples were collected with Bolt LDS buffer and Bolt reducing agent (Invitrogen) and run on polyacrylamide gels. The gels were then washed with a fixative solution (50% methanol, 12% acetic acid, 0.05% formaldehyde) for 2 to 24 h. After fixing, the gels were washed (35% ethanol) for 20 min three times. Gels were placed in a sensitizer solution (0.02% sodium thiolsulfate) for 5 min, followed by 3 brief rinses with double-distilled water (ddH2O). Following the brief rinsing steps, the gels sat in a staining solution (0.2% silver nitrate, 0.076% formaldehyde) for 20 min. After the staining, the gels were washed with ddH2O and revealed with developing solution (6% sodium carbonate, 0.05% formaldehyde, and 0.0004% sodium thiol sulfate) for 1 to 5 min. The reaction was then stopped with a stop solution (12% acetic acid) for 5 min, and the gels were stored in H2O at 4°C.
Indirect immunofluorescence.Huh7 cells grown on coverslips were either treated with 500 μM DFMO or untreated. After 4 days, cells were infected with MP-12. Cells were fixed with 4% formalin for 15 min, washed with PBS, permeabilized, and blocked with 0.2% Triton X-100 and 2% bovine serum albumin (BSA) in PBS (blocking solution) for 30 min at room temperature. Cells were sequentially incubated with primary mouse antinucleoprotein antibody (1:100 in blocking solution, overnight at 4°C) and secondary goat anti-mouse 488 nm, (1:500 in PBS, 1 h, room temperature). Cells were then incubated in affinity-purified Fab donkey anti-mouse IgG fragment (Jackson ImmunoResearch) (1:100 in blocking solution, 1 h, room temperature), followed by primary mouse anti-RNA (J2) antibody (1:100) and 633-nm-conjugated phalloidin (1:500) in blocking solution (overnight at 4°C) and then secondary donkey anti-mouse 568 nm (1:500 1 h at room temperature). After washing with PBS, cells were mounted with Invitrogen ProLong Diamond Antifade mounting medium with DAPI (4′,6′-diamidino-2-phenylindole) for 30 min before imaging. To ensure that there was no binding of the second fluorescent primary antibody to the first primary mouse antibody, control cells were processed as described above but without the second primary antibody step. Samples were imaged with a Zeiss Axio Observer 7 with a Lumencor Spectra X LED light system and a Hamamatsu Flash 4 camera using appropriate filters and Zen Blue software with a 40× objective.
Electron microscopy.Concentrated Rift Valley fever virus (RVFV) samples were prepared for imaging by transmission electron microscopy (TEM) according to a published method (29) with minor modifications. Concentrated RVFV samples suspended in medium were fixed in 5% glutaraldehyde (Electron Microscopy Sciences, Hatfield, PA) for 24 h at room temperature. Fixed viral samples were subjected to ultracentrifugation and then resuspended in 50 μl deionized water. Next, 5 μl of sample was pipetted onto a Formvar- and carbon-coated 200-mesh copper grid (Electron Microscopy Sciences) held in place by negative-action tweezers. Five minutes later, the edge of the grid was blotted with a wedge of Whatman filter paper to remove excess sample. The sample was negatively stained by immersing the grid in a 1% solution of phosphotungstic acid (PTA) (pH 6.8) for 1 min. After the grid was removed from the 1% PTA solution, the edge of the grid was blotted with a wedge of Whatman filter paper, and the sample was allowed to air dry for 5 min. The sample was placed into a grid storage box and allowed to dry for 24 h prior to imaging with a Philips CM120 transmission electron microscope (TSS Microscopy, Beaverton, OR) equipped with a BioSprint 16 megapixel digital camera (Advanced Microscopy Techniques, Woburn, MA).
Particle size measurement.Images of viral particles were obtained via TEM and negative staining. ImageJ was used to measure the diameter of the particles by measuring the provided standard from the TEM imaging software and measuring the particles, which were then normalized to the standard.
Fractionation.Sucrose gradient ultracentrifugation was performed using an Optima L-90K ultracentrifuge with an SW60 Ti Beckman rotor and Ultra-Clear centrifuge tubes (11 by 60 mm; Beckman Coulter). Various densities of sucrose (20% to 45%) were added to the centrifuge tubes in 2.5% increments between 20% and 30% and in 5% increments between 30% and 45%. Tubes were weighed before addition of 50 μl to 200 μl of viral supernatant, unless otherwise indicated. The centrifuge tubes were then spun at 45,000 rpm for 18 h at 4°C. Samples were then collected based on fraction amount (450 μl) and analyzed by Western blotting, silver staining, and RT-qPCR.
Interference assay.Virus passed on untreated or DFMO-treated Huh7 cells as described above was combined with stock virus, with equivalent PFU from both stock and passaged virus. This combined virus was used to infect naive Huh7 or Vero-E6 cells for 48 h prior to enumeration of viral titers via plaque assay. As a control, stock virus alone was used to infect cells.
Statistical analysis.Prism 6 (GraphPad) was used to generate graphs and perform statistical analysis. For all analyses, the two-tailed Student t test was used to compare groups, unless otherwise noted, with α = 0.05. For tests of sample proportions, P values were derived from calculated Z scores with two tails and α = 0.05.
ACKNOWLEDGMENTS
We are grateful to Thomas Gallagher for critical discussions and helpful insights concerning this project, as well as to Tuli Mukhopadhyay for electron microscopy advice. We thank Susan Uprichard for Huh7 cells, Katherine Knight for RAW264.7 cells, and Makio Iwashima for THP-1 cells, as well as for helpful discussions. We also thank Shinji Makino and Kaori Terasaki for generously providing the MP-12 strain of RVFV. We appreciate immunofluorescent imaging help and input from the labs of Ivana Kuo and Jordan Beach.
FOOTNOTES
- Received 2 April 2019.
- Accepted 28 April 2019.
- Accepted manuscript posted online 1 May 2019.
- Copyright © 2019 American Society for Microbiology.
