Virus-induced translational arrest through 4EBP1/2-dependent decay of 5'-TOP mRNAs restricts viral infection

The mosquito-transmitted bunyavirus, Rift Valley fever virus (RVFV), is a highly successful pathogen for which there are no vaccines or therapeutics. Translational arrest is a common antiviral strategy used by hosts. In response, RVFV inhibits two well-known antiviral pathways that attenuate translation during infection, PKR and type I IFN signaling. Despite this, translational arrest occurs during RVFV infection by unknown mechanisms. Here, we find that RVFV infection triggers the decay of core translation machinery mRNAs that possess a 5'-terminal oligopyrimidine (5'-TOP) motif in their 5'-UTR, including mRNAs encoding ribosomal proteins, which leads to a decrease in overall ribosomal protein levels. We find that the RNA decapping enzyme NUDT16 selectively degrades 5'-TOP mRNAs during RVFV infection and this decay is triggered in response to mTOR attenuation via the translational repressor 4EBP1/2 axis. Translational arrest of 5'-TOPs via 4EBP1/2 restricts RVFV replication, and this increased RNA decay results in the loss of visible RNA granules, including P bodies and stress granules. Because RVFV cap-snatches in RNA granules, the increased level of 5'-TOP mRNAs in this compartment leads to snatching of these targets, which are translationally suppressed during infection. Therefore, translation of RVFV mRNAs is compromised by multiple mechanisms during infection. Together, these data present a previously unknown mechanism for translational shutdown in response to viral infection and identify mTOR attenuation as a potential therapeutic avenue against bunyaviral infection

The Kaposi's Sarcoma-Associated Herpesvirus K5 E3 Ubiquitin Ligase Modulates Targets by Multiple Molecular Mechanisms▿

Kaposi's sarcoma-associated herpesvirus encodes two highly related membrane-associated, RING-CH-containing (MARCH) family E3 ubiquitin ligases, K3 and K5, that can down regulate a variety of cell surface proteins through enhancement of their endocytosis and degradation. In this report we present data that while K5 modulation of major histocompatibility complex class I (MHC-I) closely mirrors the mechanisms used by K3, alternative molecular pathways are utilized by this E3 ligase in the down regulation of intercellular adhesion molecule 1 (ICAM-1) and B7.2. Internalization assays demonstrate that down regulation of each target can occur through increased endocytosis from the cell surface. However, mutation of a conserved tyrosine-based endocytosis motif in K5 resulted in a protein lacking the ability to direct an increased rate of MHC-I or ICAM-1 internalization but still able to down regulate B7.2 in a ubiquitin-dependent but endocytosis-independent manner. Further, mutation of two acidic clusters abolished K5-mediated MHC-I degradation while only slightly decreasing ICAM-1 or B7.2 protein destruction. This same mutant abolished detectable ubiquitylation of all targets. These data indicate that while K5 can act as an E3 ubiquitin ligase to directly mediate cell surface molecule destruction, regulation of its targets occurs through multiple pathways, including ubiquitin-independent mechanisms

Kaposi's sarcoma-associated herpesvirus K3 and K5 proteins down regulate both DC-SIGN and DC-SIGNR.

Kaposi's sarcoma-associated herpesvirus (KSHV) is the etiological agent of multicentric Castleman's disease, primary effusion lymphoma and Kaposi's sarcoma. In this study, we show that like the C-type lectin DC-SIGN, the closely related DC-SIGNR can also enhance KSHV infection. Following infection, they are both targeted for down modulation and our data indicate that the KSHV MARCH-family ubiquitin ligase K5 is mediating this regulation and subsequent targeting for degradation of DC-SIGN and DC-SIGNR in the context of the virus. The closely related viral K3 protein, is also able to target these lectins in exogenous expressions studies, but only weakly during viral infection. In addition to requiring a functional RING-CH domain, several protein trafficking motifs in the C-terminal region of both K3 and K5 are important in regulation of DC-SIGN and DC-SIGNR. Further exploration of this modulation revealed that DC-SIGN is endocytosed from the cell surface in THP-1 monocytes, but degraded from an internal location with minimal endocytosis in HEK-293 cells. Pull-down data indicate that both K3 and K5 preferentially associate with immature forms of the lectins, mediating their ubiquitylation and degradation. Together, these data emphasize the molecular complexities of K3 and K5, while expanding the repertoire of targets of these two viral proteins

Wild-Type K5 and K3 cause enhanced endocytosis of DC-SIGN, but not DC-SIGNR.

<p><b>A</b>) Empty vector transduced THP-1 cells, as well as K3 wt, K3 Y/A, K5 wt and K5 Y/A expressing stable lines, were stained for the indicated surface markers. Histograms on the left show staining of these markers (filled lines) overlayed with isotype control (dashed lines) or with vector transduced THP-1 cells (open lines). As shown in the right bar graph, surface levels of DC-SIGN were determined by flow cytometry and mean channel fluorescence was normalized to vector transduced THP-1 cells. Data shown are an average of three independent experiments with error bars showing standard deviation. <b>B</b>) Indicated THP-1 cells lines were treated for 60 minutes with 80 µM dynasore or mock treated with DMSO at 37°C. Upon dynasore removal, cells were chased for 30 min at 37°C in complete medium and endocytosis was stopped by adding sodium azide to all samples. Half of the samples were stained for DC-SIGN surface levels on ice (left panel), while the other half was stained for total DC-SIGN levels after paraformaldehyd fixation and saponin permeabilisation (right panel). Mean channel fluorescence for DMSO treated vector transduced cells was set to 100%. The data shown are representative of three independent experiments. <b>C</b>) 293 cells stable expressing DC-SIGN or DC-SIGNR were transiently transfected with 4 µg GFP-tagged K3 wt, K5 wt or the Y/A mutant of either protein. After 36 to 48 hours, cells were subjected to dynasore treatment and release similar to that described in B except that three release time points were assayed. Mean channel fluorescence for surface DC-SIGN, DC-SIGNR, and MHC I were determined by flow cytometry for GFP positive (filled symbols) as well as GFP negative (open symbols) populations. For normalization, DMSO treated cells were set to 100 percent (indicated by the −60 min time point) and relative fluorescence following dynasore treatment (indicated by the 0 min time point) and release (indicated by the 10 min, 30 min and 60 min time points) was calculated. Data presented are representative of three independent experiments.</p

DC-SIGN and DC-SIGNR are ubiquitylated and degraded in a proteasomal- and lysosomal-dependent pathway by K3 and K5.

<p><b>A</b>) 293T cells were co-transfected with either 1.75 µg DC-SIGN (top panels) or 1.75 µg DC-SIGNR (bottom panels), 0.5 ug HA-tagged ubiquitin, and 1.75 µg wild-type or RING-CH mutant constructs of GST-tagged K3 or K5, as indicated. At 36–48 hours post-transfection, cells were collected and lysed. Pull-down (PD) was done using glutathione-sepharose beads, followed by SDS-PAGE and immunoblotting analysis. The precipitated proteins were probed with anti-HA antibodies. <b>B</b>) 293T cells were transiently transfected as in panel A. At 36–48 hours post-transfection cells were treated with chloroquine (ChQ ), MG132, or DMSO (solvent), as indicated. Cell lysates were then subjected to pull-down with glutathione beads followed by immunoprecipitation of DC-SIGN. Precipitated proteins were then subjected to western blot with an anti-HA antibody. Supernatants from the immunoprecipitation were subjected to TCA precipitation and then western blot with an anti-GST antibody. As additional controls, whole cell lysates were subjected to western blot with antibodies against DC-SIGN or HA-tagged ubiquitin. Data is representative of multiple experiments.</p

DC-SIGN and DC-SIGNR co-precipitate with K3 and K5.

<p><b>A</b>) 293T cells were transiently transfected with 1 µg DC-SIGN (left panels) or DC-SIGNR (right panels) together with 3 µg wild type (wt) or RING-CH mutant (mZn) constructs of GST-tagged K3 or K5, and as controls EglN1, an unrelated protein, and empty GST expression vector (−) as indicated. At 36–48 hours post-transfection, cells were harvested, lysed in NP40 lysis buffer and subjected to pull-down (PD) using glutathione-sepharose beads. Purified proteins were resolved by SDS-PAGE electrophoresis and subjected to WB analysis for DC-SIGN/R co-purification. GST pull-down was evaluated by reprobing the blots with a GST specific antibody. Even expression of DC-SIGN or DC-SIGNR was verified by WB of whole cell lysates (WCL).Closed arrowheads indicate GST-tagged EglNI, K3 or K5; open arrowheads indicate unfused GST. Data is representative of multiple experiments. <b>B and C</b>) Co-transfection of 293T cells with (<b>B</b>) DC-SIGN or (<b>C</b>) DC-SIGNR together with vector or GST-tagged wild-type K3 or K5 was repeated. Samples were split in half and subjected to either GST pull-down or immunoprecipitation for lectin proteins. Aliquots of the purified proteins were left untreated (UT) or digested with either EndoH (E) or PNGaseF (P). Following SDS-PAGE, changes in mobility were detected by WB for DC-SIGN or DC-SIGNR. Data is representative of multiple experiments.</p

K3 and K5 affect the stability of DC-SIGN and DC-SIGNR in a RING-CH domain-dependent mechanism.

<p><b>A</b>) 293 cells stably expressing wild-type K3 or K5, or the RING-CH mutant of either viral protein were transiently transfected with 2 µg DC-SIGN or DC-SIGNR constructs. At ∼48 hpt,cells were lysed in RIPA buffer and 30 µg of normalized lysate were loaded per sample. Protein levels of DC-SIGN or DC-SIGNR were determined by WB, and then blots were reprobed for lamin B as a loading control. Data is representative of at least three independent experiments. <b>B</b>) THP-1 cell stably expressing the indicated K5 constructs or empty vector were stained for cell surface levels of DC-SIGN. Solid histogram, empty vector; grey histogram, K5 construct; dotted histogram, isotype control. <b>C</b>) The same THP-1 cell lines used in Panel B, were lysed and subjected to western blotting with either a DC-SIGN (H-200) or GAPDH (O411) antibody, as loading control. Data is representative of at least three independent experiments.</p

Infectivity of KSHV is enhanced in the presence of DC-SIGN and DC-SIGNR.

<p><b>A</b>) 293T cells were transfected with empty pcDNA3 vector or expression constructs for DC-SIGN or DC-SIGNR. After 24 hours, cells were infected with 20 µl Bac16ΔK3ΔK5 or left uninfected as controls. Cells were harvested after additional 24 hours and surface stained with a DC-SIGN/R antibody (H-200) and analyzed by flow cytometry. Top three panels show transfected cells stained for DC-SIGN/R followed by PE- (DC-SIGN), FITC- (DC-SIGNR) or both (vector) conjugated secondary antibodies. Bottom panels shows KSHV infection of 293T cells transiently expressing DC-SIGN or DC-SIGNR. <b>B, left panels</b>) 293 cell lines stably expressing a vector construct, DC-SIGN or DC-SIGNR were fluorescently stained for surface expression of DC-SIGN or DC-SIGNR. The mean channel fluorescence is indicated in the upper right hand corner. Open histograms – secondary antibody alone; shaded histograms – DC-SIGN or DC-SIGNR staining. <b>B, right panel</b>) 293 pcDNA3, DC-SIGN or DC-SIGNR stable cell lines were pre-incubated with a control antibody (anti-ICAM1, 7 µg/ml), with mannan (100 µg/ml) or a monoclonal antibody specific for DC-SIGN (MAB161; 7 µg/ml) for 30 minutes on ice. These cells were then infected with wild type KSHV (Bac16 or rKSHV.219) at an MOI of 0.01. After 72 hours cells were harvested and evaluated for infection by flow cytometry measuring GFP expression. Infection rates were normalized to 293 pcDNA3 cells treated with the control antibody. The fold increase in relative infectivity is indicated. Data are representative of four independent experiments with two performed in triplicate. <b>C</b>) 293 pcDNA3, DC-SIGN or DC-SIGNR stable lines were infected with 50 µl of concentrated Bac16 wildtype (wt), or mutants with deletion of K3 only (ΔK3), K5 only (ΔK5), or deletion of both K3 and K5 (ΔK3ΔK5) as indicated. At 72 hours post-infection, the cells were stained for surface expression of DC-SIGN, DC-SIGNR or MHC class I. GFP fluorescence was used as a marker for infection. MHC I staining is shown for infected 293 DC-SIGNR cells. Inset numbers indicate percentage of cells in each quadrant.</p

A novel class of TMPRSS2 inhibitors potently block SARS-CoV-2 and MERS-CoV viral entry and protect human epithelial lung cells.

The host cell serine protease TMPRSS2 is an attractive therapeutic target for COVID-19 drug discovery. This protease activates the Spike protein of severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) and of other coronaviruses and is essential for viral spread in the lung. Utilizing rational structure-based drug design (SBDD) coupled to substrate specificity screening of TMPRSS2, we have discovered covalent small-molecule ketobenzothiazole (kbt) TMPRSS2 inhibitors which are structurally distinct from and have significantly improved activity over the existing known inhibitors Camostat and Nafamostat. Lead compound MM3122 (4) has an IC50 (half-maximal inhibitory concentration) of 340 pM against recombinant full-length TMPRSS2 protein, an EC50 (half-maximal effective concentration) of 430 pM in blocking host cell entry into Calu-3 human lung epithelial cells of a newly developed VSV-SARS-CoV-2 chimeric virus, and an EC50 of 74 nM in inhibiting cytopathic effects induced by SARS-CoV-2 virus in Calu-3 cells. Further, MM3122 blocks Middle East respiratory syndrome coronavirus (MERS-CoV) cell entry with an EC50 of 870 pM. MM3122 has excellent metabolic stability, safety, and pharmacokinetics in mice, with a half-life of 8.6 h in plasma and 7.5 h in lung tissue, making it suitable for in vivo efficacy evaluation and a promising drug candidate for COVID-19 treatment