*Article* **Interferon-Induced HERC5 Inhibits Ebola Virus Particle Production and Is Antagonized by Ebola Glycoprotein**

**Ermela Paparisto <sup>1</sup> , Nina R. Hunt <sup>1</sup> , Daniel S. Labach <sup>1</sup> , Macon D. Coleman <sup>1</sup> , Eric J. Di Gravio <sup>1</sup> , Mackenzie J. Dodge <sup>1</sup> , Nicole J. Friesen <sup>1</sup> , Marceline Côté 2 , Andreas Müller <sup>3</sup> , Thomas Hoenen <sup>3</sup> and Stephen D. Barr 1,\***


**Abstract:** Survival following Ebola virus (EBOV) infection correlates with the ability to mount an early and robust interferon (IFN) response. The host IFN-induced proteins that contribute to controlling EBOV replication are not fully known. Among the top genes with the strongest early increases in expression after infection in vivo is IFN-induced HERC5. Using a transcription- and replication-competent VLP system, we showed that HERC5 inhibits EBOV virus-like particle (VLP) replication by depleting EBOV mRNAs. The HERC5 RCC1-like domain was necessary and sufficient for this inhibition and did not require zinc finger antiviral protein (ZAP). Moreover, we showed that EBOV (Zaire) glycoprotein (GP) but not Marburg virus GP antagonized HERC5 early during infection. Our data identify a novel 'protagonist–antagonistic' relationship between HERC5 and GP in the early stages of EBOV infection that could be exploited for the development of novel antiviral therapeutics.

**Keywords:** Ebola virus; Marburg virus; HERC5; antiviral; interferon

## **1. Introduction**

Ebola virus (EBOV) is a member of the *Filoviridae* family of single-stranded negativesense RNA viruses with a filamentous morphology. EBOV infection results in severe hemorrhagic fever and can lead to death 6-16 days after the onset of symptoms in up to 90% of cases, making EBOV one of the most virulent pathogens to infect humans [1]. Studies involving primate models, and human studies carried out during the 2013–2016 outbreak, showed that EBOV exposure results in an early and robust immune response, largely characterized by the up-regulation of IFN-stimulated genes [2–12]. A contributing factor to the pathophysiology of EBOV infection is the ability of the virus to evade the host IFN response [7,13–16]. Using in vitro models of infection, it was shown that EBOV is able to evade the innate immune response through various IFN antagonisms, notably involving VP24 and VP35 proteins [17–19]. The key mediators of this early cellular IFN response to EBOV and how EBOV withstands this early response are not fully characterized.

Restriction factors are key intrinsic mediators of the early IFN response and potently inhibit different steps in the life cycle of evolutionarily diverse viruses in the absence of viral antagonists [20]. Bone marrow stromal cell antigen 2 (BST-2)/tetherin is one such factor that potently inhibits the release of EBOV from cells by tethering virions to the surface of cells [21,22]. This inhibition is counteracted by EBOV GP [23–26]. IFNinducible trans-membrane proteins 1–3 (IFITM1–3) comprise another family of factors

**Citation:** Paparisto, E.; Hunt, N.R.; Labach, D.S.; Coleman, M.D.; Di Gravio, E.J.; Dodge, M.J.; Friesen, N.J.; Côté, M.; Müller, A.; Hoenen, T.; et al. Interferon-Induced HERC5 Inhibits Ebola Virus Particle Production and Is Antagonized by Ebola Glycoprotein. *Cells* **2021**, *10*, 2399. https://doi.org/10.3390/ cells10092399

Academic Editors: Reinhild Prange and Alexander E. Kalyuzhny

Received: 29 April 2021 Accepted: 31 August 2021 Published: 13 September 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

that restrict the cellular entry of EBOV, although an EBOV antagonist to these proteins has yet to be identified [27,28]. HECT and RCC1-like containing domain 5 (HERC5) are some of the genes with the strongest early increases in expression in multiple tissues after EBOV infection [3,5,6,29]. HERC5 is an evolutionarily ancient restriction factor that inhibits the replication of diverse viruses [30–36]. By virtue of its C-terminal HECT domain, HERC5 is the main cellular E3 ligase for conjugating ISG15 to substrates and localizes to polyribosomes to modify newly translated viral proteins, thereby disrupting key aspects of viral particle production [31,35,37,38]. E3 ligase-independent antiviral activity has also been demonstrated towards HIV-1, where it inhibits the nuclear export of incompletely-spliced viral RNAs by a mechanism requiring its N-terminal RCC1-like domain (RLD) [30].

Here, we examined the antiviral activity of HERC5 towards EBOV VLP production and replication. We identified a novel E3 ligase-independent mechanism by which HERC5 inhibits viral particle production involving the depletion of EBOV mRNAs. In addition, we demonstrated that EBOV GP antagonizes HERC5 activity and rescues EBOV VLP production and replication.

#### **2. Materials and Methods**

#### *2.1. Cell Lines*

293T and HeLa cells were obtained from American Type Culture Collection. 293T ZC3HAV1 (ZAP) knockout cells were obtained from Dr. Takaoka (Hokkaido University, Japan) via Dr. Li (University of California, Los Angeles, CA, USA) and Dr. MacDonald (The Rockefeller University, New York, NY, USA). Cells were maintained in standard growth medium (Dulbecco's Modified Eagle's Medium (DMEM)), supplemented with 10% heatinactivated Fetal Bovine Serum (FBS), 100 U/mL Penicillin and 100 µg/mL Streptomycin) at 37 ◦C with 5% CO2.

#### *2.2. Plasmids, Transfections, Antibodies and Quantitative Western Blotting*

Expression plasmids carrying FLAG-tagged HERC5, HERC5-∆RLD, HERC5-∆HECT and HERC5-C994A, and HERC4 have been described previously [36]. The plasmid carrying FLAG-tagged RLD only (pFLAG-RLDonly) was generated by standard restriction enzyme cloning of the HERC5 RLD (containing a 30 stop codon) into p3xFLAG-CMV-10 (Sigma). The promoterless empty vector plasmid pGL3, pEGFP-C1 (pEGFP) and pZAP (short isoform) were obtained from Promega, Clontech and Dharmacon, respectively. pLKO.1/scrambled shRNA and pLKO.1/HERC5 shRNA were previously described [30,31]. VP40 and GP were cloned into p3xFLAG-CMV-10 (Sigma) to generate pFLAG-VP40, pFLAG GP and pEGFP-C1 (containing a CMV promoter) (Clontech) to generate pVP40- EGFP using standard restriction enzyme cloning. EBOV expression plasmids: pCAGGS plasmids (containing a CMV enhancer, chicken beta-actin promoter and beta-actin intron sequence) carrying only EBOV (Zaire) VP40, VP30, VP35, L, NP, or GP were obtained from Dr. Kawaoka (University of Wisconsin) [39]. Plasmids for the trVLP assay were provided by Dr. Hoenen (Friedrich-Loeffler-Institut, Germany): Plasmids carrying NP, VP35, VP30, L, Tim-1, T7 and the tetracistronic minigenomes (p4cisvRNA-hrLuc, p4cis-vRNA-EGFP) have been previously described [40,41]. All EBOV gene sequences in the minigenomes and plasmids carrying NP, VP35, VP30, and L originated from the Zaire EBOV isolate *H. sapiens*tc/COD/1976/Yambuku-Mayinga. The EBOV GP and MARV GP expression plasmids were kind gifts of Dr. Cunningham (Brigham and Women's Hospital) [42,43]. Transfections were performed using Lipofectamine 2000 (Invitrogen) per manufacturer's instructions unless otherwise stated. Co-transfections of HERC5 plasmids with pVP40 were performed at a ratio of 10:1, respectively, unless otherwise noted. VP40 VLPs were purified from cell supernatants by centrifugation over a 20% sucrose cushion at 21,000× *g* for 2 h. Cell lysates and VP40 VLP pellets were subjected to quantitative Western blot analyses using LI-COR, as previously described [30]. Densitometric analysis was performed using ImageJ 1.53e 64-bit version software. Antibodies: Anti-FLAG was purchased from Sigma, anti-ZAP from AbCam (Cat. #ab154680), anti-VP40 from GeneTex (Cat. #GTX134034), anti-MARV GP

from Alpha Diagnostic International (Cat. #MVGP12-A), anti-EBOV GP from Bio-Techne (Cat. #MAB9016), anti-β-actin from Rockland, anti-EGFP from Clontech and anti-GAPDH (clone 6C5) from EMD/Millipore.

#### *2.3. Confocal Immunofluorescence Microscopy*

HeLa cells were cultured in 12-well plates on 18 mm coverslips and co-transfected with either pFLAG-HERC5 and pVP40-EGFP (10:1 ratio) or pGL3 and pVP40-EGFP (10:1 ratio). Twenty-four hours after transfection, the coverslips containing the cells were washed twice with PF buffer (1× PBS + 1% FBS), fixed for 10 min in 1× PBS containing 4% formaldehyde and 2% sucrose, permeabilized in 1× PBS containing 0.1% Triton X 100 (Sigma) and then washed twice more with PF buffer. Coverslips were incubated with primary antibody rabbit anti-FLAG (1:500 dilution) for 1 h, washed 3× with PF buffer and incubated with either secondary antibody anti-rabbit 594 (1:1000) for 1 h. Coverslips were washed 3×, incubated in Hoechst 33342 (1:10,000 dilution) (Life Technologies) for 5 min and washed 6× with PF buffer. Coverslips were then mounted on glass slides with 10 µL Vectashield mounting media (Vector Laboratories Inc., Burlingame, CA, USA) and sealed with nail polish. Confocal micrographs were obtained using a Leica TCS SP8 (Leica Microsystems) microscope, and Leica Application Software X was used for image acquisition.

#### *2.4. Transmission Electron Microscopy*

Cells were co-transfected with empty vector or pFLAG-HERC5 and pVP40-EGFP at a 10:1 ratio. After 48 h, cells were resuspended in media, fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate (pH 7.4) for 2 h, and washed 3× in 0.1 M sodium cacodylate. Cells were pelleted and fixed with 2% osmium tetroxide in sodium cacodylate. After ~1 h in the dark, cells were washed 3× in ddH2O. Water was discarded, and samples were left at 4 ◦C overnight. Samples were dehydrated by adding 1 mL 20% acetone in ddH2O, mixed and incubated for 10 min at room temperature. Cells were pelleted, acetone removed, and the procedure was repeated with 50%, 70%, 90%, 100%, 100% and 100% acetone. Cells were embedded in resin by adding 1 mL of a 2:1 mix of acetone:resin (Epon) and incubated for ~4 h at room temperature in a rotating tube shaker. Cells were pelleted, acetone:resin mix was discarded and repeated with a 1:1 mix overnight, 1:2 mix overnight, and finally, resin only overnight. Samples were cut in 70 nm slices using a Sorval Ultracut ultramicrotome and placed onto 400 mesh nickel grids (Embra). Grids were placed on drops of 2% uranyl acetate in ddH2O to stain for 20 min in the dark and washed 5–6× in ddH2O for 1 min. Samples were then stained in drops of Sato's lead citrate (5 mM calcined lead citrate, 11 mM lead nitrate, 11 mM lead acetate, 95 mM sodium citrate) for 1 min and washed using ddH2O. Samples were imaged using a Phillips CM10 Transmission Electron Microscope. The AMT Advantage digital imaging system was used for image acquisition.

#### *2.5. Quantitative PCR*

The total RNA was extracted using the PureLink RNA mini kit (Ambion, Life Technologies). Using the M-MLV reverse transcriptase and Oligo(dT) primers (Eurofins), 500 ng of RNA was reverse transcribed to cDNA. Prior to qPCR, cDNA samples were diluted 1:5 with water. Each PCR reaction consisted of 10 µL of SYBR Green Master Mix, 1.6 µL of gene-specific primers (0.8 µL of 10 µM forward primer and 0.8 µL of 10 µM reverse primer), 4 µL of diluted cDNA, and water to a total volume of 20 µL. Quantification of endogenous mRNA was run on the QuantStudio5 qPCR machine (Applied Biosystems) under the following cycling conditions: 2 min at 95 ◦C and 40 cycles of 5 sec at 95 ◦C, 10 s at 60 ◦C, and 20 s at 72 ◦C. The QuantStudio Design and Analysis Desktop Software (version 1.4) was used to determine the C<sup>T</sup> for each PCR reaction. Primer pairs were as follows: HERC5- (fwd: 50 ATG AGC TAA GAC CCT GTT TGG 30 ; rev: 50 CCC AAA TCA GAA ACA TAG GCA AG 30 ); ZAP- (fwd: 50 CGCTTAATGGTAGCTGCAGC 30 ; rev: 50 CCTACAGAACA-GAGGTGGATTCC 30 ); GAPDH- (fwd: 50 CAT GTT CGT CAT GGG TGT GAA CCA 3 0 ; rev: 50 AGT GAT GGC ATG GAC TGT GGT CAT 30 ); EGFP- (fwd: 50 GACAAC-

CACTACCTGAGCAC 30 ; rev: 50 CAGGACCATGTGATCGCG30 ); EBOV VP40- (fwd: 5 0GCTTCCTCTAGGTGTCGCTG30 ; rev 50GGTTGCCTTGCCGAAATGG30 ); EBOV GP- (fwd: 50GTGAATGGGCTGAAAACTGC30 ; rev 50CCGTTCCTGATACTTTGTGC30 ); EBOV VP30- (fwd: 50CCAGACAGCATTCAAGGG30 ; rev 50GCTGGAGGAACTGTTAATGG30 ); EBOV VP35- (fwd: 50CGACTCAAAACGACAGAATGC30 ; rev 50GGTTTGGCTTCGTTTGT TGC30 ); EBOV NP- (fwd: 50GCCAACTTATCATACAGGCC30 ; rev 50CCAAATACTTGACT GCGCC30 ); EBOV L- (fwd: 50CCTAGTCACTAGAGCTTGCG30 ; rev 50GGCTCAACAGGA CAGAATCC30 ). To ensure no carry-over of DNA into each total purified RNA sample, 100 ng of RNA was used directly as a template without reverse transcription for qPCR using the primer sets described above.

#### *2.6. trVLP Assay*

Expression plasmids carrying tim-1, T7, NP, VP35, VP30, L, and the tetracistronic minigenome (p4cis-vRNA-hrLuc) carrying luciferase, VP40, GP and VP24 have been previously described [40,44]. trVLP assays were performed as previously described, with the following changes [40,41]. Passage zero (p0) cells were seeded in 12-well plates and transfected at 50% confluency using Transit LT-1 (Mirus Bio LLC, Madison, WI, USA) with expression plasmids carrying T7-polymerase (125 ng; all amounts per well), the viral proteins NP (62.5 ng), VP35 (62.5 ng), VP30 (37.5 ng), L (500 ng), a tetracistronic minigenome (125 ng), and Firefly luciferase (100 ng) following the manufacturer's instructions. Twentyfour hours prior to infection of p1, p2, p3 and p4 cells, target cells were pre-transfected with expression plasmids carrying NP (62.5 ng), VP35 (62.5 ng), VP30 (37.5 ng), L (500 ng), Tim-1 (125 ng) and either HERC5 (125 ng) or empty vector (125 ng).

#### *2.7. Cell Viability Assay*

293T cells were co-transfected with pFLAG-VP40, GFP-VP40 or GFP alone, as well as increasing concentrations of pFLAG-HERC5 or empty vector control plasmid. Forty-eight hours post-transfection Cell Counting Kit-8 (CCK-8) (GLPBIO) was used to measure cell viability as per the manufacturer's instructions.

#### *2.8. Statistical Analyses*

GraphPad Prism v9 was used for all statistical analyses stated in the text. *p* values and statistical tests used are stated in the text where appropriate. *p* values less than 0.05 were deemed significant. Quantification of immunogold labelling for statistical analysis was performed as described [31].

#### **3. Results**

#### *3.1. HERC5 Inhibits EBOV trVLP Replication*

Previous studies have identified HERC5 as a potent inhibitor of diverse viruses [30–36]. To determine if HERC5 restricts EBOV particle production and replication, we used an EBOV (Zaire) transcription- and replication-competent VLP (trVLP) system. This system utilizes a tetracistronic minigenome ('4cis') carrying a *luciferase* reporter gene together with *VP40*, *VP24*, and *GP* (Figure 1A) [40,45]. The advantage of this system over conventional VLP assays is that the viral proteins VP40, GP and VP24 are encoded by the minigenome and expressed from the EBOV promoter in a more natively regulated fashion [40]. The co-expression of this minigenome with NP, VP35, VP30, and L drive genome replication and transcription, synthesis of the minigenome-encoded proteins, and formation of infectious trVLPs. These trVLPs incorporate minigenomes and are capable of undergoing multiple rounds of replication and infection in target cells that express NP, VP35, VP30, L and Tim-1 (Figure 1B). The replication of these trVLPs was quantified over multiple passages (every three days) by measuring the luciferase reporter activity within cells. As a negative control, the plasmid carrying the Ebola *L* gene was omitted from the transfections, which abrogated the trVLP formation. Compared to the control cells transfected with an empty vector plasmid, cells expressing HERC5 exhibited a significant

*Cells* **2021**, *10*, x 5 of 20

reduction in trVLP replication over four passages (Figure 1C). The reduction in luciferase reporter activity also correlated with a reduction in GP and VP40 mRNA levels (Figure 1D). with an empty vector plasmid, cells expressing HERC5 exhibited a significant reduction in trVLP replication over four passages (Figure 1C). The reduction in luciferase reporter activity also correlated with a reduction in GP and VP40 mRNA levels (Figure 1D).

infectious trVLPs. These trVLPs incorporate minigenomes and are capable of undergoing multiple rounds of replication and infection in target cells that express NP, VP35, VP30, L and Tim-1 (Figure 1B). The replication of these trVLPs was quantified over multiple passages (every three days) by measuring the luciferase reporter activity within cells. As a negative control, the plasmid carrying the Ebola *L* gene was omitted from the transfections, which abrogated the trVLP formation. Compared to the control cells transfected

**Figure 1.** HERC5 inhibits EBOV trVLP replication. (**A**) Schematic depicting EBOV full-length genome and the derived tetracistronic minigenome. (**B**) The trVLP propagation assay. A tetracistronic EBOV minigenome (4cis) is expressed in cells together with the viral ribonucleoprotein complex (RNP) proteins (NP, VP35, VP30 and L). After the initial transcription by a co-expressed T7 polymerase, the minigenome is replicated and transcribed by the RNP proteins. Expression of **Figure 1.** HERC5 inhibits EBOV trVLP replication. (**A**) Schematic depicting EBOV full-length genome and the derived tetracistronic minigenome. (**B**) The trVLP propagation assay. A tetracistronic EBOV minigenome (4cis) is expressed in cells together with the viral ribonucleoprotein complex (RNP) proteins (NP, VP35, VP30 and L). After the initial transcription by a co-expressed T7 polymerase, the minigenome is replicated and transcribed by the RNP proteins. Expression of VP40, GP and VP24 from the minigenome leads to the formation of infectious trVLPs containing minigenomes, which can infect target cells. Multiple infectious cycles can be modeled in cells expressing NP, VP35, L, VP30 and Tim-1 without the need for additional transfections of plasmids carrying VP40, GP and VP24. The figure was adapted from (Watt et al., 2014), copyright © American Society for Microbiology, J. Virol. 88, 2014, 10,511–10,524, doi:10.1128/JVI.01272-14. (**C**) Quantification of trVLP

propagation in the presence and absence of HERC5. The trVLP propagation assay was performed using tetracistronic minigenomes carrying a luciferase reporter, EBOV VP40, VP24 and EBOV GP over four passages (spanning 12 days). All EBOV minigenomes and plasmids carrying the EBOV proteins are based on EBOV *H. sapiens*-tc/COD/1976/Yambuku-Mayinga. Luciferase reporter activity relative to the control (trVLPs propagated in the absence of HERC5) is shown. The data shown represent the average (+/− S.E.M.) of four independent experiments. Linear regression analysis, F = 39.14. DFn = 1, DFd = 36; *p* < 0.0001. (**D**) The mRNA of GP and VP40 was measured using qRT-PCR at each passage. The data shown represent the average (+/− S.E.M.) of the four independent experiments represented in part C. \* *p* < 0.05; One-way ANOVA with Dunnet's multiple comparisons test compared to the control.

#### *3.2. HERC5 Inhibits EBOV VP40 Particle Production*

Previous studies showed that HERC5 interferes with the function of key viral structural proteins [30,31,35,36]. The EBOV structural protein VP40 is necessary and sufficient for the assembly and budding of virus particles. When expressed in the absence of any other viral protein, VP40 can form VLPs that bud and are released from cells similar to wild-type EBOV [46–48]. To determine if HERC5 targets VP40, we co-transfected 293T cells with a plasmid carrying VP40 and increasing concentrations of plasmids carrying either empty vector control or FLAG-tagged HERC5. VP40 protein levels within cells and in VLPs were measured using quantitative Western blotting. HERC5 transfection did not alter cell viability (Figure S1A). As shown in Figure 2A and Figure S1B,C, HERC5 inhibited the production of VP40 VLPs in a dose-dependent manner when VP40 is tagged with either GFP or with FLAG but had no effect on intracellular GFP levels. As a control, transfection with *HERC4*, a closely related member of the small *HERC* family, did not significantly alter cell viability, VP40 or GFP levels (Figure S1D–F). In contrast, when *HERC5* mRNA levels were reduced using RNA interference, an increase in intracellular VP40 protein levels and an increase in the production of VP40 VLPs were observed compared to the control cells (Figure 2B,C).

We also assessed the impact of HERC5 expression on VLPs using confocal microscopy and transmission electron microscopy (TEM). As expected, cells expressing VP40 with enhanced green fluorescent protein fused at its amino-terminus (VP40-EGFP) exhibited punctate fluorescence at the cell surface (Figure 2D). In contrast, cells co-expressing VP40- EGFP and HERC5 exhibited substantially less punctate fluorescence at the cell surface compared to the control cells. The presence of VP40 protein at the cell surface was also confirmed using TEM and immunogold TEM (Figure 2E,F). In cells expressing VP40-EGFP alone, an accumulation of immunogold particles was observed in budding structures at the cell surface, which was significantly different from a random distribution (Tables S1 and S2). Cells expressing HERC5 exhibited markedly fewer VP40-EGFP-containing structures at the cell surface compared to the control cells. In addition, cells expressing HERC5 exhibited on average eight-fold fewer immunogold particles per cell compared to the control cells (Figure 2G). Notably, the few VP40-EGFP-containing structures that were observed in cells expressing HERC5 were located predominantly in a region under the plasma membrane.

We then asked whether the reduced VP40 protein levels correlated with reduced intracellular VP40 mRNA levels. The quantitative PCR showed that 293T cells co-expressing HERC5 and FLAG-tagged VP40 exhibited reduced intracellular levels of VP40 mRNA (nine-fold) compared to the control cells not expressing HERC5 (Figure 2H). Similar results were obtained when HERC5 was co-expressed with a VP40-EGFP fusion protein (Figure S2). As a control, HERC5 expression had no significant effect on EGFP mRNA levels when EGFP was expressed alone (Figure 2H and Figure S2). To determine if the effect of HERC5 is specific for VP40 mRNA, we assessed the impact of HERC5 expression on the level of other EBOV mRNAs. Cells co-expressing HERC5 and either VP30, VP35, L or NP exhibited a two- to five-fold reduction in mRNA levels compared to the control cells (Figure 2H). Together, these data show that HERC5 inhibits EBOV VP40 particle production by a mechanism involving the depletion of EBOV mRNAs.

*Cells* **2021**, *10*, x 8 of 20

**Figure 2.** HERC5 inhibits EBOV VP40 particle production. (**A**) 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40 (pFLAG-VP40) and increasing concentrations of FLAG-tagged HERC5 (pFLAG-HERC5). Empty vector

plasmid was transfected in the condition with no HERC5 and used to ensure equal amounts of DNA were transfected in each condition. Forty-eight hours post-transfection, purified VLPs released into the cell supernatant and intracellular protein were subjected to quantitative Western blot analysis using anti-FLAG, anti-VP40 and anti-GAPDH. The average densitometric quantification of VP40 protein bands is shown to the right after normalization to GAPDH levels (+/− S.E.M.). A representative Western blot of four independent experiments is shown. (**B**) 293T cells were co-transfected with pFLAG-VP40 and either scrambled short-hairpin RNA (shRNA) (scram) or HERC5shRNA (shHERC5). Forty-eight hours after transfection, intracellular levels of HERC5 mRNA were quantified via qPCR. Data shown is the average (+/− S.E.M.) of three independent experiments. (**C**) 293T cells were transfected with either scrambled short-hairpin RNA (shRNA) (scram) or HERC5shRNA (shHERC5) for 24 h and then with pFLAG-HERC5 and pFLAG-VP40 for forty-eight hours. Purified VLPs released into the cell supernatant and intracellular protein were subjected to quantitative Western blot analysis using anti-FLAG and anti-GAPDH. The average densitometric quantification of VP40 protein bands is shown to the right after normalization to GAPDH levels (+/− S.E.M.). A representative Western blot of four independent experiments is shown. (**D**) HeLa cells were co-transfected with pVP40-EGFP and either empty vector (control) or pFLAG-HERC5 and visualized using confocal microscopy 48 h post-transfection. (**E**) 293T cells were "mock" transfected (control), transfected with empty vector and pVP40-EGFP, or transfected with pFLAG-HERC5 and pVP40-EGFP and analyzed via transmission electron microscopy (TEM) after 48 h. Virus particles beneath the plasma membrane are indicated with arrows. (**F**) Representative immunogold TEM images of 293T cells transfected as in (**E**) and labelled with 5 (+/− 2) nm anti-GFP immunogold particles. Immunogold-labelled VLPs are indicated with arrows. Scale bars = 500 nm. (**G**) The number of gold particles per positive cell was counted and presented as the average number of particles per cell (+/− S.E.M). (**H**) 293T cells were co-transfected with plasmids carrying FLAG-HERC5 (or empty vector) and either EBOV VP40, VP30, VP35, L, NP, GP or GFP at a ratio of 10:1 (HERC5: EBOV plasmids). Forty-eight hours post-transfection viral mRNA was measured using qPCR after normalization to GAPDH mRNA levels. Data shown are representative of three independent experiments (+/− S.E.M.). \*\*\*\* *p* < 0.0001, \*\*\* *p* < 0.001, \*\* *p* < 0.01, \* *p* < 0.05; One-way ANOVA with Dunnet's multiple comparisons test compared to the control (**A**,**G**); Student's paired t-test (**B**,**C**,**H**).

#### *3.3. HERC5 RLD Is Necessary and Sufficient for Inhibition of VP40 Particle Production*

To determine if the RLD or HECT domains of HERC5 are required for inhibition, we tested the ability of several HERC5 mutants to inhibit VP40 particle production. 293T cells were co-transfected with plasmids carrying VP40 and either empty vector (control), wild type HERC5 or HERC5 mutants lacking the RCC1-like domain (HERC5-∆RLD), spacer region (HERC5-∆spacer) or HECT domain (HERC5-∆HECT). We also tested the HERC5 RLD alone (HERC5-RLDonly) or HERC5 containing a cysteine to an alanine point mutation of residue 994 (HERC5-C994A), which specifically inactivates its E3 ligase activity (Figure 3A). Each of the FLAG-tagged mutant proteins was expressed at similar levels in 293T cells (Figure 3B).

As shown in Figure 3C, cells expressing wild type HERC5, HERC5-∆HECT or HERC5- C994A reduced VP40 protein levels, which also correlated with reduced VP40 VLP production. In contrast, cells expressing HERC5-∆RLD, and to a lesser extent HERC5-∆spacer, exhibited a diminished capacity to reduce VP40 protein levels and VP40 VLP production. Notably, expression of the HERC5 RLD alone (HERC5-RLDonly) reduced VP40 protein levels and VP40 VLP production similar to wild-type HERC5 (Figure 3D). We also examined the ability of the different HERC5 mutants to reduce VP40 mRNA levels. All HERC5 mutants except for HERC5-∆RLD significantly reduced VP40 mRNA levels (Figure 3E). Taken together, these data show that the HERC5 RLD is necessary and sufficient to reduce VP40 mRNA levels and VP40 particle production.

**Figure 3.** The RLD is necessary and sufficient for HERC5-mediated restriction. (**A**) Schematic of the different HERC5 mutant constructs. (**B**) Representative Western blot showing consistent expression of wild-type HERC5 and mutant forms of HERC5.

293T cells were transfected with either empty vector or plasmids carrying FLAG-tagged HERC5, HERC5-∆RLD, HERC5- RLDonly, HERC5-∆Spacer, HERC5-∆HECT or HERC5-C994A. Forty-eight hours after transfection, cell lysate was subjected to Western blot analysis using anti-FLAG and anti-GAPDH. (**C**) 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector, wild-type HERC5 or one of the HERC5 mutants listed in (**A**). Forty-eight hours post-transfection, purified VLPs released into the supernatant and intracellular protein were examined by Western blotting using anti-FLAG and anti-GAPDH. VP40 protein levels were quantified densitometrically after normalization to GAPDH levels (graphs on the right). (**D**) 293T cells were co-transfected with plasmids carrying VP40-EGFP and either empty vector, HERC5 or HERC5-RLDonly. Cell lysates and VLPs were analyzed via Western blotting using anti-GFP and anti-GAPDH. VP40-EGFP protein levels were quantified densitometrically (graphs on the right). (**E**) 293T cells were cotransfected with plasmids carrying FLAG-tagged VP40 and either empty vector, HERC5, HERC5-∆RLD, HERC5-RLDonly, HERC5-∆Spacer, HERC5-∆HECT or HERC5-C994A. Forty-eight hours post-transfection, mRNA was isolated and used to measure intracellular VP40 mRNA levels using qPCR. All data shown are representative of three independent experiments (+/− S.E.M.). \*\*\*\* *p* < 0.0001, \*\*\* *p* < 0.001, \*\* *p* < 0.01, \* *p* ≤ 0.05, ns (not significant) *p* > 0.05; One-way ANOVA with Dunnet's multiple comparisons test compared to the control. **Figure 3.** The RLD is necessary and sufficient for HERC5-mediated restriction. (**A**) Schematic of the different HERC5 mutant constructs. (**B**) Representative Western blot showing consistent expression of wild-type HERC5 and mutant forms of HERC5. 293T cells were transfected with either empty vector or plasmids carrying FLAG-tagged HERC5, HERC5-ΔRLD, HERC5-RLDonly, HERC5-ΔSpacer, HERC5-ΔHECT or HERC5-C994A. Forty-eight hours after transfection, cell lysate was subjected to Western blot analysis using anti-FLAG and anti-GAPDH. (**C**) 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector, wild-type HERC5 or one of the HERC5 mutants listed in (**A**). Fortyeight hours post-transfection, purified VLPs released into the supernatant and intracellular protein were examined by Western blotting using anti-FLAG and anti-GAPDH. VP40 protein levels were quantified densitometrically after normalization to GAPDH levels (graphs on the right). (**D**) 293T cells were co-transfected with plasmids carrying VP40-EGFP and either empty vector, HERC5 or HERC5-RLDonly. Cell lysates and VLPs were analyzed via Western blotting using anti-GFP and anti-GAPDH. VP40-EGFP protein levels were quantified densitometrically (graphs on the right). (**E**) 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector, HERC5, HERC5-ΔRLD, HERC5- RLDonly, HERC5-ΔSpacer, HERC5-ΔHECT or HERC5-C994A. Forty-eight hours post-transfection, mRNA was isolated and used to measure intracellular VP40 mRNA levels using qPCR. All data shown are representative of three independent experiments (+/- S.E.M.). \*\*\*\* *p* < 0.0001, \*\*\* *p* < 0.001, \*\* *p* < 0.01, \* *p* ≤ 0.05, ns (not significant) *p* > 0.05; One-way ANOVA with Dunnet's multiple comparisons test compared to the control.

#### *3.4. HERC5 Depletes VP40 mRNA Independently of ZAP 3.4. HERC5 Depletes VP40 mRNA Independently of ZAP*

ZAP (also called Zinc finger CCCH-type, antiviral 1, ZC3HAV1, and Poly (ADPribose) polymerase 13, PARP13) is an antiviral protein that causes significant loss of viral mRNAs from evolutionarily diverse RNA viruses, including *Filoviridae, Retroviridae, Togaviridae* and *Hepadnaviridae* [49–55]. We, therefore, asked if ZAP was required for HERC5-mediated depletion of EBOV mRNA. We co-expressed VP40 and HERC5 in 293T cells that were knocked out for all ZAP isoforms and measured VP40 mRNA and protein levels using qPCR and Western blotting [56,57]. Cells expressing HERC5 in the absence of ZAP significantly reduced VP40 mRNA levels (Figure 4A). Exogenous expression of ZAP (short isoform) in the ZAP knockout cells reduced VP40 mRNA levels as previously shown [52,56]. Co-expression of HERC5 and ZAP together resulted in an enhanced loss of VP40 mRNA (Figure 4A). In support of this observation, cells expressing HERC5 in the absence of ZAP significantly reduced intracellular VP40 protein and VP40 VLPs the cell supernatant (Figure 4B,C). Together, these data show that ZAP is not required for HERC5-mediated reduction of VP40 mRNA. ZAP (also called Zinc finger CCCH-type, antiviral 1, ZC3HAV1, and Poly (ADP-ribose) polymerase 13, PARP13) is an antiviral protein that causes significant loss of viral mRNAs from evolutionarily diverse RNA viruses, including *Filoviridae, Retroviridae, Togaviridae* and *Hepadnaviridae* [49–55]. We, therefore, asked if ZAP was required for HERC5 mediated depletion of EBOV mRNA. We co-expressed VP40 and HERC5 in 293T cells that were knocked out for all ZAP isoforms and measured VP40 mRNA and protein levels using qPCR and Western blotting [56,57]. Cells expressing HERC5 in the absence of ZAP significantly reduced VP40 mRNA levels (Figure 4A). Exogenous expression of ZAP (short isoform) in the ZAP knockout cells reduced VP40 mRNA levels as previously shown [52,56]. Co-expression of HERC5 and ZAP together resulted in an enhanced loss of VP40 mRNA (Figure 4A). In support of this observation, cells expressing HERC5 in the absence of ZAP significantly reduced intracellular VP40 protein and VP40 VLPs the cell supernatant (Figure 4B,C). Together, these data show that ZAP is not required for HERC5 mediated reduction of VP40 mRNA.

**Figure 4.** HERC5 restricts VP40 independently of ZAP. 293T ZAP knockout cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector control, HERC5, ZAP **Figure 4.** HERC5 restricts VP40 independently of ZAP. 293T ZAP knockout cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector control, HERC5, ZAP (short isoform),

or HERC5 and ZAP (short isoform). Twenty-four hours post-transfection, cell lysates and VLPcontaining supernatants were harvested. (**A**) Intracellular VP40 mRNA levels were measured using qPCR and normalized to GAPDH. The data shown are representative of four independent experiments. (**B**) Purified VLPs released into the cell supernatant and intracellular proteins were subjected to Western blot analysis using anti-FLAG and anti-GAPDH. Representative Western blot of three independent experiments is shown. (**C**) The average densitometric quantification of VP40 protein bands from B is shown after normalization to GAPDH levels. Results are presented as mean (± SEM) fold changes in VP40 protein or mRNA. \*\*\*\* *p* < 0.001, \*\*\* *p* < 0.001, One-way ANOVA with Tukey's multiple comparisons test.

#### *3.5. EBOV GP and L Proteins Antagonize HERC5*

Despite an early and robust IFN-signaling response to EBOV infection, EBOV proteins ultimately suppress this response leading to pathogenesis [2–12]. Given the potent antiviral activity of HERC5 towards EBOV mRNAs, we asked if any of the EBOV proteins could antagonize this activity. VP40 mRNA levels in cells co-expressing HERC5 and various EBOV proteins were measured by qPCR. As shown in Figure 5A, VP40 mRNA levels were rescued in cells co-expressing GP or L protein, but not VP30, VP35, NP or the non-EBOV protein vesicular stomatitis virus-G (VSV-G) protein. Western blot analysis of cell lysates correlated with the qPCR data where only L and GP proteins rescued intracellular VP40 protein levels (Figure 5B). Western blot analysis of VP40 VLPs in the supernatant revealed that GP but not L protein rescued VLP production, indicating that only GP was able to fully rescue VLP production.

To determine if the ability of EBOV GP to antagonize HERC5 is specific to the *Ebolavirus* genus, we tested the ability of Marburg virus (MARV) GP, which belongs to the *Marburgvirus* genus, to antagonize HERC5. In contrast with EBOV GP, co-expression of MARV GP failed to rescue VP40 VLP production (Figure 5C). Together these data show that EBOV GP antagonizes HERC5 activity and that this antagonism does not appear to be conserved between filovirus genera.

**Figure 5.** EBOV GP and L antagonize HERC5. 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector or HERC5 and one plasmid carrying either EBOV VP30, VP35, L NP, GP or VSV-G. Forty-eight hours

post-transfection, VP40 mRNA was measured using qPCR (**A**) and VP40 protein levels in cell lysates and VLPs released into supernatant were analyzed by quantitative Western blotting and quantified densitometrically after normalization to GAPDH levels (**B**). (**C**) 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector or HERC5, and one of EBOV GP (eGP) or MARV GP (mGP). Forty-eight hours post-transfection, VP40 protein levels in cell lysates and VLPs released into the supernatant were analyzed via Western blotting using anti-FLAG and anti-GAPDH. The data shown represent the average (+/− S.E.M.) of three independent experiments. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001, \*\*\*\* *p* < 0.0001, ns (not significant) *p* > 0.05; One-way ANOVA with Dunnett's multiple comparisons test compared to the control (**A**); Student's paired t-test (**B**,**C**). and either empty vector or HERC5 and one plasmid carrying either EBOV VP30, VP35, L NP, GP or VSV-G. Forty-eight hours post-transfection, VP40 mRNA was measured using qPCR (**A**) and VP40 protein levels in cell lysates and VLPs released into supernatant were analyzed by quantitative Western blotting and quantified densitometrically after normalization to GAPDH levels (**B**). (**C**) 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40 and either empty vector or HERC5, and one of EBOV GP (eGP) or MARV GP (mGP). Forty-eight hours post-transfection, VP40 protein levels in cell lysates and VLPs released into the supernatant were analyzed via Western blotting using anti-FLAG and anti-GAPDH. The data shown represent the average (+/- S.E.M.) of three independent experiments. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001, \*\*\*\* *p* < 0.0001, ns (not significant) *p* > 0.05; One-way ANOVA with Dunnett's multiple comparisons test compared to the control (**A**); Student's paired t-test (**B**,**C**).

*Cells* **2021**, *10*, x 14 of 20

**Figure 5.** EBOV GP and L antagonize HERC5. 293T cells were co-transfected with plasmids carrying FLAG-tagged VP40

#### *3.6. EBOV and MARV GP Differentially Antagonize HERC5 Inhibition of EBOV trVLP Replication 3.6. EBOV and MARV GP Differentially Antagonize HERC5 Inhibition of EBOV trVLP Replication*

We utilized the EBOV trVLP system described in Figure 1 to determine if genusspecific GP (EBOV or MARV) could antagonize the ability of HERC5 to inhibit trVLP replication. To test the effect of different GPs on trVLP replication, two different sets of trVLP particles were generated at P0. One set contained EBOV GP (trVLPEBOV GP) and was generated as described in Figure 1A. The second set was generated in an identical way except that the EBOV *GP* gene in the '4cis' plasmid minigenome was substituted with the MARV *GP* gene (trVLPMARV GP). This allowed us to test the impact of different GPs in the VLPs while maintaining the same background of EBOV proteins. As a negative control, the plasmid carrying the Ebola *L* gene was omitted from the transfections, which abrogates trVLP formation. Compared to the control cells not expressing HERC5, cells expressing HERC5 exhibited significantly reduced levels of trVLPMARV GP and trVLPEBOV GP replication over four passages (spanning 12 days) (Figure 6). Notably, HERC5 inhibited trVLPMARV GP replication significantly more than trVLPEBOV GP replication over two passages (*p* < 0.01, Two-way ANOVA). Together, these data show that EBOV GP and MARV GP differentially antagonize HERC5 inhibition of EBOV trVLP replication. We utilized the EBOV trVLP system described in Figure 1 to determine if genus-specific GP (EBOV or MARV) could antagonize the ability of HERC5 to inhibit trVLP replication. To test the effect of different GPs on trVLP replication, two different sets of trVLP particles were generated at P0. One set contained EBOV GP (trVLPEBOV GP) and was generated as described in Figure 1A. The second set was generated in an identical way except that the EBOV *GP* gene in the '4cis' plasmid minigenome was substituted with the MARV *GP* gene (trVLPMARV GP). This allowed us to test the impact of different GPs in the VLPs while maintaining the same background of EBOV proteins. As a negative control, the plasmid carrying the Ebola *L* gene was omitted from the transfections, which abrogates trVLP formation. Compared to the control cells not expressing HERC5, cells expressing HERC5 exhibited significantly reduced levels of trVLPMARV GP and trVLPEBOV GP replication over four passages (spanning 12 days) (Figure 6). Notably, HERC5 inhibited trVLPMARV GP replication significantly more than trVLPEBOV GP replication over two passages (*p* < 0.01, Twoway ANOVA). Together, these data show that EBOV GP and MARV GP differentially antagonize HERC5 inhibition of EBOV trVLP replication.

**Figure 6.** EBOV GP and MARV GP differentially antagonize HERC5. Quantification of trVLP propagation in the presence and absence of HERC5. The trVLP propagation assay was performed using tetracistronic minigenomes carrying a luciferase reporter, EBOV VP40, VP24 and either EBOV GP or MARV GP over four passages (spanning 12 days). All EBOV minigenomes and plasmids carrying the EBOV proteins are based on EBOV *H. sapiens*-tc/COD/1976/Yambuku-Mayinga. As a negative control ('No L'), the plasmid carrying the Ebola *L* gene was omitted from the transfections. Luciferase reporter activity relative to the control (trVLPs propagated in the absence of HERC5) is shown. The data shown represent the average (+/− S.E.M.) of at least six independent experiments. \*\*\*\* *p* < 0.0001, \* *p* ≤ 0.05, ns (not significant) *p* > 0.05; Twoway ANOVA with Dunnett's multiple comparisons test compared to the no HERC5 control. **Figure 6.** EBOV GP and MARV GP differentially antagonize HERC5. Quantification of trVLP propagation in the presence and absence of HERC5. The trVLP propagation assay was performed using tetracistronic minigenomes carrying a luciferase reporter, EBOV VP40, VP24 and either EBOV GP or MARV GP over four passages (spanning 12 days). All EBOV minigenomes and plasmids carrying the EBOV proteins are based on EBOV *H. sapiens*-tc/COD/1976/Yambuku-Mayinga. As a negative control ('No L'), the plasmid carrying the Ebola *L* gene was omitted from the transfections. Luciferase reporter activity relative to the control (trVLPs propagated in the absence of HERC5) is shown. The data shown represent the average (+/− S.E.M.) of at least six independent experiments. \*\*\*\* *p* < 0.0001, \* *p* ≤ 0.05, ns (not significant) *p* > 0.05; Two-way ANOVA with Dunnett's multiple comparisons test compared to the no HERC5 control.

#### **4. Discussion**

Hundreds of IFN-induced proteins are part of the early and robust immune response to EBOV infection in primates [2–12]. Characterization of the key effector proteins of this defense and how EBOV overcomes them will provide a better understanding of the virus–host interactions that occur early in infection. HERC5 is one of the most up-regulated antiviral proteins in the early response to EBOV infection in vivo; however, its role in EBOV replication was previously unknown [3,5,6,29].

In this study, we showed that HERC5 inhibits EBOV VLP replication via a novel E3 ligase-independent mechanism. This mechanism involves the depletion of viral mRNAs and requires the RLD domain of HERC5. We previously showed that HERC5 inhibits the nuclear export of HIV-1 RNA genomes by a different E3 ligase-independent mechanism, one that also requires the RLD domain of HERC5 [30]. These E3 ligase-independent antiviral activities, together with its well-documented E3 ligase-dependent antiviral activities [58], identifies HERC5 as a multifunctional antiviral protein. It is perhaps not surprising that HERC5 has evolved multiple mechanisms of restriction of viruses. The ancestral *HERC* gene is believed to have arisen from a gene fusion event between an *RCC1* like gene and a *HECT* gene [59,60]. This fusion event gave rise to a family of small HERC proteins containing an amino-terminal RLD and a carboxyl-terminal HECT domain that is highly conserved among vertebrates spanning >595 million years of evolution [36,59,60]. Moreover, HERC5 has been evolving under strong positive selection, which is characteristic of many host restriction factors involved in an evolutionary struggle with viruses [30,60,61]. The ability of HERC5 to inhibit viruses via both E3 ligase-dependent and -independent mechanisms would confer a strong evolutionary advantage to its host, making it more difficult for viruses to evolve countermeasures to HERC5.

Like HERC5, ZAP is present in evolutionarily diverse vertebrates and has evolved under strong positive selection [30,36,62]. ZAP targets diverse viruses such as HIV-1, MoLV and XMRV (*Retroviridae*), Ebola and Marburg viruses (*Filoviridae*), alphavirus, Sindbis, Semliki Forest and Ross River viruses (*Togaviridae*), hepatitis B virus (*Hepadnaviridae*) and double-stranded DNA murine gamma herpesvirus (*Herpesviridae*) [49–55,63]. ZAP is known to inhibit a wide range of antiviral activities, including recruiting the exosome complex to target viral RNAs for degradation [49,51,53,55,57,64–68]. ZAP also exhibits virus specificity since it has no antiviral effect on vesicular stomatitis, poliovirus, yellow fever and herpes simplex I viruses [49]. We showed here that HERC5 depletes EBOV mRNAs in a ZAP-independent manner. Our finding that the HERC5 RLD is necessary and sufficient for EBOV mRNA depletion further supports an E3 ligase-independent mechanism of restriction. It was previously shown that the RLD is required for the association of HERC5 with polyribosomes [35]. It is possible that HERC5 exploits this interaction to recruit other RNA degradation machinery to EBOV mRNAs.

Although we showed that the RLD alone was necessary and sufficient to inhibit particle production, HERC5 lacking the RLD failed to completely inhibit VP40 VLP particle production. Since the RLD is important but not essential for its E3 ligase activity, it is possible that the E3 ligase activity of HERC5 also confers some antiviral activity towards VLP production via ISGylation of viral and/or host proteins involved in particle production [31,35,38]. It was previously shown that over-expression of ISG15 alone inhibited budding of EBOV VP40 VLPs by disrupting Nedd4 function and subsequent ubiquitination of VP40, which is necessary for viral egress [69]. It is unknown whether HERC5 was involved in this activity since it was not investigated. Although our data show that the predominant mechanism by which HERC5 inhibits EBOV VLP production involves the depletion of EBOV mRNAs, visual inspection of cells co-expressing EBOV VP40 and HERC5 by TEM and confocal microscopy revealed an accumulation of the VP40 protein at the localized regions in the plasma membrane in some cells, consistent with the idea of a second mechanism of inhibition acting later in particle production. HERC5-induced trapping of virus particles at the plasma membrane has also been observed with HIV-1 [31]. However, it is also possible that these accumulations represent particles in the process of

budding that have escaped HERC5 restriction. HERC5 reduced intracellular mRNA levels of viral protein expressed both from a plasmid system (Figure 2H) and of viral mRNA expressed from a tetracistronic minigenome. It is unknown how HERC5 can target viral RNAs but not non-viral RNA such as GFP. Perhaps virus-specific RNA sequences recruit HERC5 and/or RNA depletion machinery similar to how ZAP selectively recognizes high CpG-containing viral RNA. Further studies are needed to decipher this novel antiviral function of HERC5.

Animal model studies have suggested that the Type I IFN response plays an important role in restricting EBOV replication and that the ability of EBOV to overcome this response may be a requirement for lethal infection [70,71]. Although EBOV VP24 and VP35 can act broadly to dampen the IFN response, several IFN-induced antiviral proteins, including HERC5, are also highly upregulated early in response to other stimuli associated with infection, such as pro-inflammatory cytokines [72–74]. As such, it is likely that EBOV evolved additional antagonists of such antiviral proteins. Indeed, EBOV GP can directly antagonize the restriction factor BST-2/tetherin without altering BST-2/tetherin expression levels or cellular localization [24,75–82]. As shown herein, EBOV GP also antagonizes HERC5 without altering HERC5 expression levels. Although controversial, GP sequence diversity has been shown to affect EBOV transmission and virulence, as demonstrated in the 2013-2016 EBOV epidemic [83,84]. We showed here that variations in GP sequence, such as those found between different filovirus genera (e.g., EBOV and MARV), also influence the potency of antagonism of HERC5 during the early stages of EBOV trVLP replication. It is unclear how GP, which is predominantly localized to the plasma membrane, can rescue EBOV mRNA levels. GP expression is known to alter the expression and trafficking of select cellular proteins; therefore, it is possible that proteins involved in viral RNA stability are affected by GP expression [85–87]. Important next steps will be to characterize the mechanism of GP antagonism and to test the importance of this HERC5-GP axis early in infection using animal models.

It is interesting that EBOV L protein was also able to rescue HERC5-induced VP40 mRNA depletion but unable to antagonize the release of VP40 VLPs into the cell supernatant. The mechanism underlying this antagonism is not fully understood; however, it was previously shown that L protein antagonizes ZAP [52]. It is possible that L protein also specifically antagonizes HERC5-induced depletion of mRNAs. However, we speculate that the E3 ligase activity of HERC5 remains functional, leading to the ISGylation of viral and/or host proteins and subsequent arrest of later steps in viral particle production.

In conclusion, we showed that HERC5 inhibits EBOV virus particle production by a mechanism involving the depletion of EBOV mRNAs. Our data also identifies a novel 'protagonist–antagonistic' relationship between HERC5 and GP early in EBOV infection. With the ability to inhibit HERC5 and other restriction factors, GP is an attractive target for the development of small molecule compounds that interfere with this antagonism.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/cells10092399/s1, Figure S1: HERC5 depletes GFP- and FLAG-tagged VP40 mRNA but not GFP mRNA, Figure S2: HERC5 depletes GFP- and FLAG-tagged VP40 mRNA but not GFP mRNA, Table S1: Quantification of 5nm gold particle-labeled anti-GFP in cells expressing empty vector and VP40-EGFP, Table S2: Quantification of 5nm gold particle-labeled anti-GFP in cells expressing HERC5 and VP40-EGFP.

**Author Contributions:** Conceptualization, S.D.B., E.P. and N.R.H.; methodology, E.P., N.R.H., D.S.L., M.D.C., E.J.D.G., M.J.D. and N.J.F.; formal analysis, E.P., N.R.H., D.S.L., M.D.C., E.J.D.G., M.J.D., N.J.F. and S.D.B.; investigation, E.P., N.R.H., D.S.L., M.D.C., E.J.D.G., M.J.D. and N.J.F.; resources, E.P., N.R.H., D.S.L., M.D.C., E.J.D.G., M.J.D., M.C., A.M., T.H. and S.D.B.; writing—original draft preparation, S.D.B., E.P. and N.R.H.; writing—review and editing, S.D.B., E.P., N.R.H., D.S.L., M.D.C., M.C. and T.H.; supervision, S.D.B.; project administration, S.D.B.; funding acquisition, S.D.B. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Canadian Institutes of Health Research (CIHR) to S.D.B., grant numbers HBF134179 and HBF137693.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** We thank Melody Li (University of California, Los Angeles, CA, USA), Margaret MacDonald (The Rockefeller University, New York, NY, USA) and Akinori Takaoka (Hokkaido University, Hokkaido, Japan) for kindly providing the ZAP knockout cell line. We thank Tirthankar Ray for their help in constructing the VP40 plasmid. We thank Karen Nygard and Richard Gardiner for their help with electron microscopy at the Western University Biotron Facility, and Claudia Seah and Flavio Beraldo at the London Regional Microscopy Facility, Western University) for their help with confocal microscopy.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


**Abenaya Muralidharan and St Patrick Reid \***

Department of Pathology and Microbiology, University of Nebraska Medical Center, Omaha, NE 68198-5900, USA; abenaya.muralidharan@unmc.edu

**\*** Correspondence: patrick.reid@unmc.edu; Tel.: +402-559-3644

**Abstract:** Arboviruses are known to cause large-scale epidemics in many parts of the world. These arthropod-borne viruses are a large group consisting of viruses from a wide range of families. The ability of their vector to enhance viral pathogenesis and transmission makes the development of treatments against these viruses challenging. Neutrophils are generally the first leukocytes to be recruited to a site of infection, playing a major role in regulating inflammation and, as a result, viral replication and dissemination. However, the underlying mechanisms through which neutrophils control the progression of inflammation and disease remain to be fully understood. In this review, we highlight the major findings from recent years regarding the role of neutrophils during arboviral infections. We discuss the complex nature of neutrophils in mediating not only protection, but also augmenting disease pathology. Better understanding of neutrophil pathways involved in effective protection against arboviral infections can help identify potential targets for therapeutics.

**Keywords:** neutrophils; arboviruses; mosquito; inflammation; pathology

**Citation:** Muralidharan, A.; Reid, S.P. Complex Roles of Neutrophils during Arboviral Infections. *Cells* **2021**, *10*, 1324. https://doi.org/10.3390/cells 10061324

Academic Editors: Thomas Hoenen and Allison Groseth

Received: 30 March 2021 Accepted: 21 May 2021 Published: 26 May 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

## **1. Introduction**

Neutrophils are the most abundant leukocytes in the blood. They serve as the first line of defense against incoming pathogens, quickly mobilizing to the site of infection [1]. While neutrophils can have protective immunostimulatory activities, they can also have debilitating immunosuppressive activities by inhibiting T cell functions [1–3]. In addition, some viruses such as influenza, specifically H5N1, and West Nile virus are known to infect and replicate within neutrophils, using these cells as reservoirs for dissemination, although mechanisms involved in this phenomenon remain unclear [4,5].

Arthropod-borne viruses or arboviruses have a unique effect on neutrophil function since viral factors as well as vector factors can affect the activity of these cells. Arboviruses are a diverse group of viruses [6–8]. They are transmitted through blood-feeding insects such as mosquitoes and ticks, and include viruses such as chikungunya virus (CHIKV), dengue virus (DENV), West Nile virus (WNV), Zika virus (ZIKV), yellow fever virus (YFV), and eastern equine encephalitis virus (EEEV). This group of viruses have been known to cause significant morbidity and mortality around the world, with the potential to spread quickly and expand their geographical range due to their distinct mode of transmission using arthropods. Furthermore, the changes in climate and increase in urbanization help augment transmission and infectivity of these viruses [9–15].

The heterogeneity among this exceptionally large group of viruses makes developing therapeutics challenging. Neutrophils are generally the first to infiltrate infected sites. However, the role of these cells during viral infection is not yet fully understood [16]. Targeting neutrophil pathways or proteins that activate or suppress neutrophils can serve as a useful strategy for drug and vaccine development. In this review, we highlight the recent advancements in understanding the beneficial and detrimental nature of neutrophils during arboviral infections.

#### **2. Mosquitoes**

Ticks and mosquitoes enhance disease severity as well as transmission of many viruses [17–21]. Mosquitoes are the most common type of arthropods that spread diseases including malaria, dengue, West Nile, Zika, and chikungunya fevers. Mosquito-transmitted diseases affect hundreds of millions of people each year, resulting in about 750,000 deaths every year [22–25]. During a blood meal, an adult female mosquito punctures the skin of a vertebrate and ingests the fluid. If the vertebrate is infected, the virus is also ingested along with the blood. The virus can then replicate, cross the midgut barrier, and reside in the salivary gland of the mosquito at high titers [26–29]. During the subsequent blood meal, the contents of the salivary gland are released below the skin to counteract the host's hemostasis and inflammatory responses, allowing the virus to enter the epidermis and dermis [30–32].

Mosquitoes' innate immune responses allow them to survive infections, making them effective carriers. They use the RNA interference pathway for protection against viral infections, including arboviruses [33,34]. In addition, *Aedes aegypti* mosquitoes have been shown to use the JAK/STAT pathway in response to WNV, DENV, and YFV [35]. Some species of mosquitoes also utilize the immediate response of apoptosis in the salivary glands and midgut to control viral load [36,37].

The mosquito plays a major role in creating an ideal environment for virus entry and replication. Indeed, the vector secretes anti-hemostatic, angiogenic, and vasodilatory molecules through its saliva to maintain optimum blood flow during feeding [31,32,38,39]. This microenvironment allows for enhanced infection and pathogenicity of the virus by controlling the initial replication of the virus and the potential for the infection to become systemic [38,40–45]. While some groups have hypothesized suppression of anti-viral immune responses by mosquito saliva during infection, Pingen et al. (2016) showed that mosquito bites facilitate infection by triggering a cellular influx that is inadvertently beneficial for the virus [31,32,38,39,46].

The saliva of a mosquito has been shown to contain highly active molecules involved in modulating early viral infection. Indeed, while infecting mice with arboviruses through a mosquito bite or a needle accompanied by an uninfected mosquito bite, the former resulted in more severe disease [17–20,47]. Furthermore, in chickens infected with West Nile virus (WNV) via mosquito bite, significantly high viral titers were observed in the serum compared to the group infected using a needle [48]. Similar augmentation of viremia was seen in mice infected with WNV via bite accompanied by faster neuro-invasion compared to needle-inoculated animals [42].

There are many effects a mosquito bite has on immune cells [49]. Some species of mosquitoes, namely *Anopheles stephensi* and *Anopheles gambiae*, secrete saliva that can result in chemotactic activity. The vascular permeabilization and mast cell degranulation in the skin caused by the saliva were shown to recruit dendritic cells to the feeding site and neutrophils to the draining lymph node [50–52]. Another study using humanized mice showed that seven days post-mosquito bite, there was a decrease in IL-8, a neutrophil chemoattractant, in the serum correlating to a decrease in circulating neutrophils. This corresponded to increased neutrophils in the skin [49].

Pingen et al. used mice infected with aedine mosquito-borne Semliki Forest virus (SFV), an alphavirus shown to replicate efficiently in immune-competent mice, and Bunyamwera virus (BUNV), a genetically unrelated RNA virus [6,46,53,54]. In their study, mosquito bites induced an influx of inflammatory neutrophils, which, in turn, promoted myeloid cell entry into the bite site in a CCR2-dependent manner. This augmented viral infection since myeloid cells are permissive to the virus. Interestingly, viral infection via bite synergistically enhanced CXCL2 and IL-1β expression, and neutrophil influx compared to bite alone [46]. The researchers further confirmed the role of neutrophils by depleting them and blocking inflammasome activity. This resulted in decreased inflammation and a suppressive environment for the viral infection. Depleting neutrophils also significantly reduced edema by further enhancing the vascular leakage caused by the bite. In addition, neutrophil influx into the bite site at earlier stages of infection was required for the induction of vital bite-associated genes such as IL-1β, CCL2, CCL7, and CCL12. Importantly, neutrophil depletion did not affect virus-induced genes, while neutrophils expressing IL-1β were necessary for establishing cutaneous inflammatory responses to mosquito bites [46]. Therefore, factors secreted by the mosquito augment infection by increasing neutrophil-mediated inflammation at the bite site during early stages of infection, which later determines the systemic course of the infection in mice. At later stages, however, neutrophils were required to effectively resolve the infection and decrease mortality in mice. Indeed, higher number of the neutrophil-depleted mice infected with the more virulent SFV6 succumbed to infection compared to neutrophil-sufficient mice [46].

#### **3. Zika Virus**

Zika virus (ZIKV) is a flavivirus transmitted mainly by *Aedes* species mosquitoes. ZIKV generally causes fever, cutaneous rash, headache, and malaise [55,56]. However, in the most recent 2015–2016 epidemic in Latin America and the Caribbean that affected more than 1.5 million people [57], ZIKV caused severe congenital malformations in the fetus, commonly known as Congenital Zika Syndrome, [58,59] and Guillain-Barre syndrome [60,61].

Recently, Hastings et al. conducted a study to identify specific antigenic salivary gland proteins in the *Aedes aegypti* mosquito that promotes ZIKV pathogenesis [62]. They used yeast display to identify a molecule in the saliva of the mosquito that can activate neutrophils in the host. The authors named this previously undescribed protein as neutrophil-stimulating factor 1 (NeSt1). When mice were treated with NeSt1-blocking antibodies before being bitten by ZIKV-infected mosquitoes, they had a 50% higher survival rate compared to untreated mice. Furthermore, NeSt1 was shown to activate neutrophils inducing their expression of IL-1β, and monocyte/macrophage-attracting chemokines CXCL2 and CCL2. The recruited macrophages may then be infected by the virus increasing the viral load [62]. Overall, NeSt1 stimulated neutrophils at the bite site augmenting early viral infection and ZIKV pathogenesis (Figure 1).

In contrast, another study using adult AG129 interferon α/β receptor knockout mice infected with a recent strain of ZIKV showed the protective effects of neutrophils [63]. In this mouse model, ZIKV has been shown to infect astrocytes and neurons in the brain and spinal cord. Zukor et al. observed that this infection resulted in astrogliosis along with T cell and neutrophil infiltration. The neutrophil recruitment inversely correlated with the virus-induced paresis protecting infected mice from motor deficits, indicating that neutrophils may be required for controlling ZIKV-induced disease [63]. Mechanisms underlying this protection need to be further explored. It is important to note that the differences in neutrophil activity observed in this study compared to Hastings et al. may be attributed to the absence of a mosquito vector or mosquito salivary components during infection of mice.

**Figure 1.** Neutrophil-mediated viral replication and dissemination induced by mosquito saliva. During a blood meal, mosquito carrying an arbovirus injects the virus along with its salivary gland proteins below the skin of the host. There is a decrease in IL-8 levels in the serum correlating to lower number of circulating neutrophils and higher number in the skin. One of the proteins in the saliva, neutrophil-stimulating factor 1 (NeSt1), activates the neutrophils in the dermis, the deepest layer of the skin, which houses the immune cells. IL-1β is secreted by these inflammatory neutrophils to establish cutaneous response to the bite. Additionally, bite-associated monocyte/macrophage-attracting chemokines, CCL2, CCL7, CCL12, and CXCL2, are upregulated. The infiltrating monocytes and macrophages are permissive to infection enhancing viral replication and increasing the potential for systemic spread. The mosquito saliva also causes vascular permeabilization and mast cell degranulation in the skin recruiting dendritic cells to the bite site, contributing to the inflammation, and neutrophils to the draining lymph nodes.

#### **4. Dengue Virus**

Dengue virus (DENV) can cause clinical outcomes that range from mild febrile illness to dengue fever to dengue hemorrhagic fever to life-threatening dengue shock syndrome [64]. With approximately 2.5 billion people at risk globally, DENV is the most common arbovirus [65,66]. Clinical studies of adult dengue patients showed severe neutropenia with lowest levels occurring five days post-infection [67,68]. The neutropenia, however, was not predictive of severe virus-induced disease or associated with prolonged hospital stay or death [67]. Interestingly, the low level of neutrophils was not for a lack of activation signals. In fact, neutrophil-activating cytokines, IL-8 and TNF-α, were high during DENV infections [69], while neutrophil-associated genes such as DEF4A, CEA-CAM8, BPI, and ELA2 were upregulated in the blood during severe DENV infection [70]. Neutrophil elastase levels were also increased in DENV-infected patients compared to uninfected controls, with higher elastase activity in patients with dengue hemorrhagic fever compared to dengue fever patients [71]. This suggests that enhanced neutrophil activation can be associated with severe disease.

In another study, researchers observed the formation of neutrophil extracellular traps (NETs) in vitro induced by DENV [72]. NET formation or NETosis consists of nuclear decondensation and delobulation, plasma membrane rupture, and release of DNA fibers that have anti-microbial peptides [73]. Although NETs play a crucial role when fighting infections, excessive NETosis and/or ineffective NET clearance can contribute to development of autoimmune diseases and inflammatory disorders [74,75]. Indeed, several NET-associated molecules, such as double-stranded DNA, histones, etc., are known to be autoantigens in systemic autoimmune diseases [74]. For instance, autoantibodies against NET components have been seen in systemic lupus erythematosus patients as well as an imbalance between NET formation and clearance, making them more prone to NET-mediated tissue damage [76–79]. Furthermore, NETs have also been implicated in the pathogenesis of inflammatory conditions including, but not limited to, small vessel vasculitis, psoriasis, and gout [74].

Examining the phenotypic and functional responses of neutrophils in adult dengue patients, Opasawatchai et al. observed an upregulation of CD66b on neutrophils and early stages of NET formation, indicating an activated state, during acute DENV infection [80]. CD66b is a granulocyte activation marker involved in degranulation and production of reactive oxygen species (ROS), which is essential for antiviral activity [81,82]. Interestingly, higher levels of NET components, IL-8, and TNF-α were found in patients diagnosed with the more severe dengue hemorrhagic fever compared to patients with dengue fever or healthy controls [80]. A study by Lien et al. identified the viral factor crucial for inducing NETosis in vitro and in mice to be DENV envelope protein domain III (EIII). This NET formation was alleviated in neutrophils from NLRP3 inflammasome-deficient mice, decreasing inflammation. Blocking EIII-neutrophil interactions also suppressed the NETosis [83].

The most severe disease caused by DENV comprises of systemic inflammation and increased vascular permeability. Many studies have also shown the activation of macrophages and platelets leading to an increase in proinflammatory cytokines and extracellular vesicles (EVs) [84–87] that transport proteins, peptides, and nucleic acids from one cell to another to modulate cell functions [88]. Indeed, DENV-induced release of IL-1β-containing EVs by platelets increased vascular permeability [87].

In addition, DENV enhanced release of EVs by activated platelets, which further activated CLEC5A, a spleen tyrosine kinase (Syk)-coupled C-type lectin receptor, and toll-like receptor 2 (TLR2) on neutrophils and macrophages. This induced NET formation and proinflammatory cytokine release [89]. Activation of CLEC5A is known to trigger NALP3 inflammasome activation and proinflammatory cytokine response [85,86,90], which augments systemic vascular permeability and hemorrhagic shock [86,91]. While blocking CLEC5A did not fully protect mice infected with a lethal dose of DENV [91], simultaneous blockade of CLEC5A and TLR2 significantly alleviated virus-induced inflammation and improved survival [89]. Together, these studies highlight the complex ways in which neutrophils mediate disease during the different stages of DENV infection.

#### **5. West Nile Virus**

Belonging to the same Flaviviridae family as DENV, West Nile virus (WNV) is a neuroinvasive pathogen [92]. WNV infection is typically only symptomatic in the elderly and immunocompromised individuals causing life-threatening neurological disease such as meningitis and encephalitis [92–95]. Strikingly, high levels of neutrophils were found in the cerebrospinal fluid collected from patients with WNV-induced disease, suggesting a major role of neutrophils in viral pathogenesis [96,97].

In mice infected with WNV, a rapid influx of neutrophils was seen at the site of infection promoting viral replication. Indeed, the expression of CXCL1 and CXCL2, neutrophilattracting chemokines, was significantly upregulated in macrophages upon infection [5]. Interestingly, neutrophil-depletion studies revealed a dual role of these leukocytes during infection. Neutrophils were required for effective clearance of WNV and survival shown

by higher viremia and death rate in mice depleted of neutrophils after infection. However, these cells were detrimental to the mice during early stages of infection since neutrophil depletion before WNV infection reduced viral burden and enhanced survival [5]. Overall, neutrophils can serve as reservoirs for WNV replication and dissemination as well as help defend against the virus at different stages of infection.

#### **6. Alphaviruses**

The alphavirus genus consists of many arthropod-borne viruses that are typically divided into two main groups, New World and Old World alphaviruses. New World alphaviruses such as eastern equine encephalitis virus (EEEV), western equine encephalitis virus (WEEV), and Venezuelan equine encephalitis virus (VEEV) cause encephalomyelitis in humans and are found in North and South America [98]. Old World alphaviruses that include chikungunya virus (CHIKV), Ross River virus (RRV), Mayaro virus, and o'nyongnyong virus, are now found in Europe, Africa, Asia, and Oceania and generally induce fever, rash, and arthritis [8,99].

Although New World alphaviruses, such as EEEV, can have mortality rates as high as 70%, while Old World alphaviruses rarely cause death, the latter has caused many epidemics in the past, resulting in high infection rates [100]. A RRV epidemic in 1979–1980 in the South Pacific involved more than 60,000 patients [101] while the o'nyong nyong virus infected approximately 2 million people in Africa in the 1959–1962 epidemic [102]. CHIKV has caused reoccurring epidemics in numerous countries around the Indian Ocean since 2004 with millions of confirmed cases [103] and a surprising emergence in Europe and the Pacific Region for the first time in 2007 and 2011, respectively [104–107].

Humans and horses infected with New World alphaviruses show changes in the central nervous system characterized by high levels of neutrophil infiltration during early stages of disease, which is replaced by lymphocytes as the disease progresses [108,109]. Due to the lack of literature on the roles of neutrophils during New World alphavirus infection, we will focus on Old World alphaviruses in this section.

Old World alphaviruses can cause musculoskeletal inflammatory disease in humans that can be significantly debilitating. Infection with arthritis/myositis-associated alphaviruses can present with fever, joint pain, myalgia, and impaired movement [101,110]. Importantly, the musculoskeletal pain induced by arthritogenic alphaviruses can last for months to years in RRV- or CHIKV-infected individuals [111–117]. Many studies have been conducted to determine the cause of such chronic pain. In one study by Stoermer et al., RRV infection in mice with specific deletion of arginase 1 (Arg1) in neutrophils and macrophages was well controlled at later stages of infection enhancing viral clearance from musculoskeletal tissues and improving skeletal muscle tissue pathology [118]. Arg1 is expressed by monocytes/macrophages, neutrophils, and myeloid-derived suppressor cells (MDSCs) and plays an important role in regulating immune responses [119–121]. Although LysMCre Arg1f/f mice, with conditional deletion of Arg1 in macrophages and neutrophils, had no change in the disease outcomes during the acute phase of infection, significantly enhanced protection was observed in the late stages of RRV infection [118]. Furthermore, conditional knockout of Arg1 substantially reduced Arg1 expression in musculoskeletal tissues following CHIKV and RRV infection, suggesting that macrophages and neutrophils are the predominant cells at the inflammatory sites following arthritogenic alphavirus infection [118]. Overall, the study highlighted the crucial role of Arg1 in contributing to disease severity. Specific neutrophil depleting methods such as Ly6G antibody treatments could help further narrow down the responsible cell type.

CHIKV infection is primarily characterized by macrophage and monocyte infiltration into the primary sites of virus replication, which are typically the skin, muscle, and joints. However, an influx of neutrophils, dendritic cells, natural killer cells, and lymphocytes has also been observed [122]. Indeed, resident cells at the site of infection produce neutrophilattracting chemokines, CXCL1 and CXCL2, following other viral infections [123,124]. This chemokine production by resident cells remains to be seen during CHIKV infection. The recruited neutrophils produce ROS and other cytotoxic mediators to decrease viral replication [125]. In non-mammalian models of CHIKV infection such as zebrafish, the neutrophils also serve as an important source of type I interferon for eliminating the virus and alleviating disease [126]. Even in the absence of active viral replication during chronic phases of infection, CHIKV-induced arthritis may progress due to increased cytokine expression and immune cell infiltration [122,127].

A recent study found the role of CXCL10, a chemoattractant for monocytes/ macrophages and T cells, during alphaviral infections using CHIKV and o'nyong nyong mouse models. At the peak of arthritic disease, which occurs 6 to 8 days post infection in mice, CXCL10−/<sup>−</sup> mice had decreased levels of immune infiltration as well as viral loads at the site of viral inoculation, the footpad, compared to wild-type mice [128]. The predominant populations in the infiltrates were macrophages and neutrophils in the wild-type mice following infection but this influx was significantly reduced in the CXCL10−/<sup>−</sup> mice. Interestingly, viral RNA was detected in these immune cells in wild-type mice, which was also significantly decreased in the knockout mice [128].

In another study, the role of NETs during CHIKV infection was explored. Ex vivo stimulation and infection of mouse-isolated neutrophils induced the release of NETs in a TLR7- and ROS-dependent manner neutralizing CHIKV [129]. The researchers used TLR3/7/9 triple knockout mice with TLR3−/<sup>−</sup> and TLR9−/<sup>−</sup> mice as controls due to the unavailability of TLR7−/<sup>−</sup> mice. Although knockout of TLR3 and TLR9 did not affect NET production after CHIKV infection, there may be some synergistic effects of the triple knockout affecting virus-mediated NET release [129]. In vivo infection of IFNAR−/<sup>−</sup> mice following NET inhibition enhanced susceptibility of the mice to an acute CHIKV infection confirming a crucial antiviral role of NETs. Moreover, clinical data also showed a correlation between the level of NETs in the blood and systemic viral loads in CHIKV infected patients [129–131]. Even though the role of NETs has been established during an acute CHIKV infection, they may also play a part during chronic infection. Indeed, neutrophils infiltrate the synovium and release NETs leading to damage of the joint tissues in rheumatoid arthritis [132].

CCR2 has been implicated in playing a protective role during CHIKV infection by preventing neutrophil-mediated pathology. CCR2−/<sup>−</sup> mice infected with CHIKV in the hind feet showed decreased levels of monocyte/macrophage infiltration with substantial increase in neutrophil infiltration, followed by eosinophils, compared to wild-type mice [133]. This change in cellular influx was associated with increased levels of CXCL1, CXCL2, G-CSF, and IL-1β with a decrease in IL-10, promoting neutrophil recruitment and exacerbating inflammation [134–140]. The eosinophil infiltration may be promoted by neutrophil-induced tissue damage to help control the inflammation in infected CCR2−/<sup>−</sup> mice [141]. CCR2 deficiency also led to cartilage damage in mice following CHIKV infection, which is normally not a symptom of alphaviral arthritis [133]. In fact, elevated macrophage and neutrophil infiltrates in CCR2−/<sup>−</sup> mice with collagen-induced arthritis is accompanied by more severe disease [134,142]. Interestingly, Poo et al. attempted neutrophil depletion in CCR2−/<sup>−</sup> mice after CHIKV infection, which resulted in new pathology characterized by increased foot swelling along with widespread hemorrhage and edema [133,143]. This, once again, may be suggestive of a dual role of neutrophils, where they are detrimental during certain stages of infection while protective during others.

Another group delineated the role of neutrophils during a pathogenic CHIKV infection on B cell maturation and lymphocyte influx. McCarthy et al. found that mice infected with a wild-type, not acutely cleared, strain of CHIKV had recruitment of monocytes and neutrophils to the draining lymph node (dLN). This aberrantly affected lymphocyte accumulation, lymph node organization, and virus-specific B cell responses, which was reversed by blocking the influx [144]. Interestingly, only pathogenic CHIKV decreased germinal center formation in the dLN, resulting in lower neutralizing antibodies in the serum compared to infection with an attenuated strain [145,146]. These diminished B cell responses were improved upon depletion of monocytes and neutrophils during early stages of infection [144].

Depleting either monocytes or neutrophils did not restore lymphocyte counts in the dLN, indicating that one of the two cell types is sufficient to block lymphocyte infiltration [144]. Furthermore, mice lacking type I interferon signaling (IFNAR−/−) had higher percentage of neutrophils in the dLN compared to wild-type mice following pathogenic CHIKV infection. In contrast, MyD88-deficient mice and wild-type mice treated with IL-1 receptor (IL-1R) blocking antibody at the time of infection had reduced the percentage of neutrophils [144]. Together, MyD88-IL-1R signaling plays a crucial role in promoting the accumulation of neutrophils in the dLN while type I interferon signaling inhibits the recruitment during pathogenic CHIKV infection [144].

While IFN-α was observed to inhibit neutrophil influx into the dLN, IFN-β was found to inhibit neutrophil infiltration into the musculoskeletal tissues during CHIKV infection [147]. Following CHIKV inoculation in the foot of IFN-β <sup>−</sup>/<sup>−</sup> mice, although no change was observed in viral load in the foot or the blood compared to wild-type mice, there were increased levels of neutrophils in the foot [147]. Neutrophil depletion in IFN- β −/− mice alleviated musculoskeletal disease induced by CHIKV observed through significantly reduced foot swelling. On the other hand, IFN-α <sup>−</sup>/<sup>−</sup> mice had higher viral burdens at the site of infection and in circulation [147]. This indicates that IFN-α helps limit viral replication whereas IFN-β modulates neutrophil recruitment to the site of infection that is necessary for exacerbation of disease pathology (Figure 2). Curiously, neither neutrophilattracting chemokines nor proinflammatory cytokines were upregulated in the IFN-β −/− mice to accompany the neutrophil-mediated inflammation making the mechanism through which IFN-β regulates neutrophil infiltration during acute CHIKV infection unclear [147].

It is important to note that most studies deplete neutrophils in vivo to understand their function. All the studies involving neutrophil depletion referenced in this review use Ly6G antibody treatments in mice. While Ly6G may be transiently expressed on many myeloid cells in the bone marrow including monocytes and other granulocytes, neutrophils that are circulating and recruited to the site of inflammation typically have higher Ly6G expression [148]. Basophils and eosinophils are thought to be Ly6G- or Ly6Glow/intermediate [148]. Additionally, some studies showed that Ly6G-mediated neutrophil depletion reduced only the Ly6Cintermediate neutrophil population and not the Ly6Chigh monocyte population [144].

**Figure 2.** Alphavirus infection-induced neutrophil recruitment and inflammation. Following footpad injection of mice with alphavirus, levels of neutrophil-attracting chemokines, CXCL1 and CXCL2, increase. These chemokines and CXCL10 recruit neutrophils, which release reactive oxygen species (ROS), neutrophil extracellular traps (NETs), and other cytotoxic mediators through degranulation, promoting viral clearance. The infiltrating neutrophils can also be directly infected, triggering NET release in a ROS-dependent manner. Furthermore, monocytes/macrophages infiltrating the site of infection can be directly infected. On the other hand, alphaviral infections induce monocyte and neutrophil recruitment into the draining lymph node (dLN) that inhibit germinal center formation decreasing B cell maturation and neutralizing antibody (Ab) production. MyD88-IL-1R signaling promotes the accumulation of neutrophils in the dLN, while IFN-α inhibits this influx.

#### **7. Conclusions**

Neutrophils are key players in the immune system, being the most abundant leukocytes. They are one of the first responders to the site of infection. However, the heterogeneity of their roles and the variability from one infection to another makes it difficult to determine if the effects will be beneficial or detrimental to the host (Table 1). Arboviruses not only induce neutrophil-mediated inflammation using viral factors but also through factors in their vector, adding another level of complexity. Their mode of transmission through arthropods immensely increases the rate at which they spread, highlighting the need for better understanding of the underlying mechanisms involved in pathogenesis. During arboviral infections, the time and amount of neutrophil infiltration to the site of infection may have a significant effect on the outcome. Following infection, an early influx with a high number of hyperactivated neutrophils releasing high levels of IL-1β, ROS, and NETs may augment infection and disease. However, an influx at later stages of infection may be protective. Regularly causing epidemics in the vulnerable areas of the world, arboviral infections need to be controlled with unique therapeutics that can control the vector-mediated and virus-mediated symptoms. Neutrophils are implicated in disease pathology induced by arboviruses and their vectors, making them a potential therapeutic target. In-depth understanding of the neutrophil pathways involved may be crucial for successful treatment of arboviral infections.


*Cells* **2021**, *10*, 1324


*Cells* **2021**, *10*, 1324

**Author Contributions:** Conceptualization, A.M. and S.P.R.; writing—original draft preparation, A.M.; writing—review and editing, S.P.R. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was funded by startup funds for S.P.R.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** All figures were created using Biorender.com (accessed on 21 May 2021).

**Conflicts of Interest:** The authors declare no conflict of interest.

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