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What is meant by 'proteolytic activation' of a virus?

What is meant by 'proteolytic activation' of a virus?


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Noob biologist here. I suspect that at the first stage this means 'activating the virus by lysis of a surface/spike protein'? Seems kind of obvious, but I put it here in case 'proteolytic activation' is not in fact merely a subset of 'virus activation', and means something slightly different.

Second, if the above interpretation is correct, what precisely do we mean by virus activation? Is it the release of viral RNA into the host cell cytosol? This would be my guess, as it is necessary (sufficient?) for replication of viral RNA to commence, and viral replication to occur. Another idea is that 'virus entering host cell' is enough to constitute viral activation, though I suspect some viruses can enter cells and remain dormant there. Hardly seems like activation to me.

Would really appreciate clarification of this terminology so I don't have misconceptions when reading literature.


Viruses all have some mechanism to enter cells' lipid membranes, and these need to be prevented from activating prematurely. Often this means the virus is in an inert state while it is assembled and trafficked out of the host cell, then activated by some chemical signal such as the low pH of inner endosomes. So "activation" is usually going to refer to a chemical change that removes the barriers to cell entry.

For example; in flaviviruses such as Zika or Dengue, the E glycoprotein that handles most of the host entry functions is assembled while bound to prM protein. Once virions are assembled with E in an inert arrangement, prM is proteolytically cleaved. When the new host cell absorbs the virion into an endosome and lowers the pH to digest it, the E glycoproteins rearrange into spikes that can penetrate the endosomal membrane and induce fusion with the viral membrane. If prM isn't cleaved, the virus can't infect cells. Here's a recent paper that discusses this process, with high resolution structures. For these viruses there are two rounds of activation, and only the first one is proteolytic.


Virus Activation

Cellular latency occurs following HIV provirus integration and is characterized by minimal transcriptional or translational activity of viral genes. Virus activation from a latent state is often the result of stimulation by mitogens, cytokines, or DNA-damaging agents. The regulation of viral latency remains elusive. Some of the cellular factors recruited to the LTR for the active transcription of viral genes include NF-κB, Sp-1, and TBP However, Rev, Tat, Vpu, and Nef have been implicated in mediating latency through their interactions with the LTR and replication cycle. Further understanding of the factors preventing induction of and reactivation from viral latency would be helpful for developing approaches to inhibit progression to disease. Activation of HIV from a latent state is a recent approach at bringing about a cure (see below).


Roles of Caspases in Inflammation and Apoptosis: Prospects as Drug Discovery Targets

Robert V. Talanian , Hamish J. Allen , in Annual Reports in Medicinal Chemistry , 1998

Posttranslational regulation

The rate-limiting step in caspase activation appears to be proteolytic activation . As described below, proteolytic caspase activation can be stimulated by induced proximity of caspase precursors or proteolysis by active caspases or other proteases. In activated human monocytic THP.1 cells, active caspase-1 has a very short half-life, being present at <1% of the level of precursor enzyme ( 10 ). In vitro, caspase-1 autodegradation at Asp381 ( 11 ) and reversible subunit dissociation ( 11,12 ) readily occur. In addition to being a potential contributor to inactivation, dissociation may allow rearrangement between different caspases. Murine caspase-11, probably equivalent to human caspase-4 or -5, physically interacts with and is required for murine caspase-1 activation ( 13 ). The in vitro observations of caspase precursor interdigitation ( 14 ) and of a reversible homodimeric equilibrium ( 11,12 ) suggests that formation of heteroligomeric species containing portions of up to four caspases is possible ( 11 ).


Targeting the Sonic Hedgehog Pathway in Brain Cancers: Advances, Limitations, and Future Directions

Novel Drug Therapeutics for SHH-Associated Gliomas and MBs

One venue of identifying novel drug targets is in understanding the pathways that regulate GLI1 activity outside of canonical SHH signaling ( Fig. 25.4 ). In addition, understanding the mechanisms driving MB resistance to SMO inhibitors is important.

Several kinases that regulate GLI1 protein stability and localization are as follows:

Protein kinase A (PKA) promotes proteolysis and downregulation of the activity of GLI1. PKA phosphorylation on GLI1 at its threonine 374 site results in GLI1 retention in the cytoplasm, precluding its transcriptional activity in the nucleus [92] . PKA also phosphorylates GLI1 at another site (serine 640), resulting in the interaction of GLI1 with the 14-3-3 protein and repression of GLI1 transcription [93] .

Ribosomal protein S6 kinase 1 (S6K1) phosphorylates GLI1 on serine 84, leading to GLI1 activation [94] .

Activation of AMPK reduces GLI1 protein levels and stability, thus downregulating the SHH-induced transcriptional activity [95,96] . AMPK phosphorylates GLI1 at serines 102 and 408 and threonine 1074 [95,96] . Importantly, cell lines containing GLI1 resistant to phosphorylation by AMPK exhibit increased oncogenic potential [95] .

Several important themes emerge from literature on the posttranslational modifications of GLI1. First, phosphorylation on different sites of GLI1 represents a complex code of sorts, a network of competing regulation. A basic question that remains to be elucidated is how these competing phosphorylation marks, such as concurrent phosphorylation on several sites, regulate GLI1 activity. Second, the examples of phosphorylation on GLI1 represent regulation by noncanonical (non-SHH) pathways. This is of therapeutic importance secondary methods of activating GLI1 such as phosphorylation by S6K1 represent potential sources of resistance to SHH pathway drug targets. Third, possible drug targets include activating the repressors (direct or indirect) of GLI1 activity and/or inhibiting the activators (direct or indirect) of GLI1 activity. Importantly, concurrent application of these agents with existing SMO inhibitors will be a promising avenue of research as combination treatment may improve both efficacy and prevent resistance.

The literature also reveals signaling pathways that cross talk with SHH signaling, although the exact mechanism of interaction is not yet fully understood. These interacting pathways may therefore reveal novel regulation points on SHH and GLI1 activity and are described as follows:

p53. Genome sequencing found that pediatric MBs are associated with mutated p53, whose status is an important diagnostic marker of patients with SHH-MB. p53 status plays a critical role in survival status of patients. Five-year survival rates are 81% for SHH-MB patients without p53 mutation, compared with 41% for patients with p53 mutation [97] . In mice with a single deletion of PTCH1, incidence of MB was 14%, but this incidence increased to > 95% with ablation of p53 [98] . MDM2, a negative regulator of p53, plays a role in SHH signaling as reduction in MDM2 resulted in decreased expression of GLI1 and GLI2 and small cerebella [99] . Consistent with these findings, reduction of MDM2 prevented MB tumorigenesis in PTCH1 heterozygous mice [99] . Thus, further examination on how MDM2 exactly regulates GLI1 and GLI2 expression may reveal novel regulation points.

cAMP. Several studies link the second messenger cAMP to the SHH signaling pathway and tumor pathogenesis. First, higher cAMP levels were found to be correlated with lower grade in brain tumors [100] . Conversely, the study found that lower adenylate cyclase and cAMP levels were associated with higher malignancy [100] . In a neurofibromatosis-1 model, researchers generated foci of decreased cAMP using phosphodiesterase-4A1 in mice cortical tissue and found that the majority of mice developed cortical growth whose cells exhibited tumor morphology [101] . Another study found that loss of the gene encoding G-protein Gαs resulted in SHH MB initiation [102] . Thus, increased cAMP levels may correlate with decreased tumor progression, a possibility that should be tested further for both MB and gliomas.

Bone morphogenic proteins (BMPs). BMPs have been shown to regulate SHH-mediated cell proliferation. BMP2 and BMP4 have been shown to arrest SHH-induced proliferation in primary cultures of granule cell precursors, allowing cellular differentiation. Addition of BMP2 to SHH-stimulated cultures resulted in downregulation of Smo and Gli1 mRNA levels [103] . Finally, overexpression of SMAD5 reduced the percentage of SHH-induced proliferating cells [103] . These results suggest that activation of the BMP2-SMAD5 axis may be an important targeting point in SHH MB.

bFGF. The basic FGF (bFGF) pathway appears to inhibit SHH-induced proliferation. Co-incubation of SHH with bFGF eliminated SHH-induced division [104] . Another study found that treatment of MB cells with bFGF prevented tumor formation following transplantation [105] .

Survivin. Survivin was overexpressed in PTCH1-mutant MB cells, and MB tumor cells isolated from mice harboring a deletion of survivin demonstrated significantly reduced thymidine incorporation, suggesting decreased cellular division, along with cell cycle arrest in the G2/M phase [106] . These studies show that survivin plays a role in SHH-MB pathogenesis and poses another venue of investigation.


Bioinformatics explained: Proteolytic cleavage

Proteolytic cleavage is basically the process of breaking the peptide bonds between amino acids in proteins. This process is carried out by enzymes called peptidases, proteases or proteolytic cleavage enzymes.

Proteins often undergo proteolytic processing by specific proteolytic enzymes (proteases/peptidases) before final maturation of the protein. Proteins can also be cleaved as a result of intracellular processing of, for example, misfolded proteins. Another example of proteolytic processing of proteins is secretory proteins or proteins targeted to organelles, which have their signal peptide removed by specific signal peptidases before release to the extracellular environment or specific organelle.

  • N-terminal methionine residues are often removed after translation.
  • Signal peptides or targeting sequences are removed during translocation through a membrane.
  • Viral proteins that were translated from a monocistronic mRNA are cleaved.
  • Proteins or peptides can be cleaved and used as nutrients.
  • Precursor proteins are often processed to yield the mature protein.

Proteolytic cleavage of proteins has shown its importance in laboratory experiments where it is often useful to work with specific peptide fragments instead of entire proteins.

Proteases also have commercial applications. As an example proteases can be used as detergents for cleavage of proteinaceous stains in clothing.

The general nomenclature of cleavage site positions of the substrate were formulated by Schechter and Berger, 1967-68 [Schechter and Berger, 1967], [Schechter and Berger, 1968]. They designate the cleavage site between P1-P1', incrementing the numbering in the N-terminal direction of the cleaved peptide bond (P2, P3, P4, etc..). On the carboxyl side of the cleavage site the numbering is incremented in the same way (P1', P2', P3' etc. ). This is visualized in figure 16.27.


Figure 16 . 27 : Nomenclature of the peptide substrate. The substrate is cleaved between position P1-P1'.

Proteases often have a specific recognition site where the peptide bond is cleaved. As an example trypsin only cleaves at lysine or arginine residues, but it does not matter (with a few exceptions) which amino acid is located at position P1'(carboxyterminal of the cleavage site). Another example is trombin which cleaves if an arginine is found in position P1, but not if a D or E is found in position P1' at the same time. (See figure 16.28).


Figure 16 . 28 : Hydrolysis of the peptide bond between two amino acids. Trypsin cleaves unspecifically at lysine or arginine residues whereas trombin cleaves at arginines if asparate or glutamate is absent.

Bioinformatics approaches are used to identify potential peptidase cleavage sites. Fragments can be found by scanning the amino acid sequence for patterns which match the corresponding cleavage site for the protease. When identifying cleaved fragments it is relatively important to know the calculated molecular weight and the isoelectric point.


Hemagglutinin Cleavability, Acid Stability, and Temperature Dependence Optimize Influenza B Virus for Replication in Human Airways

Influenza A virus (IAV) and influenza B virus (IBV) cause yearly epidemics with significant morbidity and mortality. When zoonotic IAVs enter the human population, the viral hemagglutinin (HA) requires adaptation to achieve sustained virus transmission. In contrast, IBV has been circulating in humans, its only host, for a long period of time. Whether this entailed adaptation of IBV HA to the human airways is unknown. To address this question, we compared two seasonal IAVs (A/H1N1 and A/H3N2) and two IBVs (B/Victoria and B/Yamagata lineages) with regard to host-dependent activity of HA as the mediator of membrane fusion during viral entry. We first investigated proteolytic activation of HA by covering all type II transmembrane serine protease (TTSP) and kallikrein enzymes, many of which proved to be present in human respiratory epithelium. The IBV HA0 precursor is cleaved by a broader panel of TTSPs and activated with much higher efficiency than IAV HA0. Accordingly, knockdown of a single protease, TMPRSS2, abrogated spread of IAV but not IBV in human respiratory epithelial cells. Second, the HA fusion pH values proved similar for IBV and human-adapted IAVs (with one exception being the HA of 1918 IAV). Third, IBV HA exhibited higher expression at 33°C, a temperature required for membrane fusion by B/Victoria HA. This indicates pronounced adaptation of IBV HA to the mildly acidic pH and cooler temperature of human upper airways. These distinct and intrinsic features of IBV HA are compatible with extensive host adaptation during prolonged circulation of this respiratory virus in the human population.IMPORTANCE Influenza epidemics are caused by influenza A and influenza B viruses (IAV and IBV, respectively). IBV causes substantial disease however, it is far less studied than IAV. While IAV originates from animal reservoirs, IBV circulates in humans only. Virus spread requires that the viral hemagglutinin (HA) is active and sufficiently stable in human airways. We resolve here how these mechanisms differ between IBV and IAV. Whereas human IAVs rely on one particular protease for HA activation, this is not the case for IBV. Superior activation of IBV by several proteases should enhance shedding of infectious particles. IBV HA exhibits acid stability and a preference for 33°C, indicating pronounced adaptation to the human upper airways, where the pH is mildly acidic and a cooler temperature exists. These adaptive features are rationalized by the long existence of IBV in humans and may have broader relevance for understanding the biology and evolution of respiratory viruses.

Keywords: acid stability airway proteases hemagglutinin host adaptation influenza A virus influenza B virus membrane fusion temperature.

Copyright © 2019 American Society for Microbiology.

Figures

Human lung tissue and airway…

Human lung tissue and airway epithelial cell lines are rich in TTSP and…

IBV HA0 is efficiently cleaved…

IBV HA0 is efficiently cleaved by a broad range of TTSPs. (A) Experiment…

TTSP cleavage generates fusion-competent IBV…

TTSP cleavage generates fusion-competent IBV HA. (A) Experiment setup. HeLa cells expressing IBV…

Pseudoparticles carrying IBV HA are…

Pseudoparticles carrying IBV HA are efficiently activated by different TTSPs. (A) Experiment setup.…

TMPRSS2 is a crucial protease…

TMPRSS2 is a crucial protease for replication of IAV but not IBV. (A)…

MDCK cells contain high levels…

MDCK cells contain high levels of HA-activating TMPRSS4. (A and B) IAV or…

IBV HA exhibits a similar…

IBV HA exhibits a similar fusion pH as human-adapted IAV HAs. (A) Experiment…

IBV HA prefers a temperature…

IBV HA prefers a temperature of 33°C for protein expression, explaining temperature-restricted fusion…

Pseudoparticles carrying B/Victoria HA require…

Pseudoparticles carrying B/Victoria HA require 33°C for infectivity. (A) Experiment setup. Pseudoparticles carrying…


Results

Cleavage of SARS-CoV-2 S1/S2 site fluorescence resonance energy transfer (FRET)-substrates by furin

The S1/S2 cleavage site of the novel emerged SARS-CoV-2 has been shown to possess a minimal furin consensus motif of the sequence R-R-A-R↓ with an alanine instead of a basic residue in the P2 position ( Fig 1B (22, 23)). Only few furin substrates possess a nonbasic residue in the P2 position, such as Pseudomonas aeruginosa exotoxin A or Shiga toxin (10, 19). To test, whether the S1/S2 sequence of SARS-CoV-2 S protein is efficiently cleaved by furin, a small series of FRET substrates was synthesized ( Fig 2A ). All compounds possess a 3-nitrotyrosine amide as P4′ residue and a 2-amino-benzoyl fluorophore in the P7 position. The analogous sequences of the S proteins from MERS-CoV, SARS-CoV, and avian infectious bronchitis virus (IBV) strain Beaudette were prepared as reference substrates. Moreover, two FRET substrates of the SARS-CoV-2 S1/S2 cleavage site with P2 A → K and A → R mutations were synthesized, to evaluate whether they could constitute even more efficient cleavage sites for furin than the wild type. The FRET substrates were tested in an enzyme kinetic assay with human furin, and their cleavage efficiency is shown in Fig 2B . The FRET substrate of the SARS-CoV-2 S1/S2 cleavage site was efficiently cleaved by recombinant furin. In contrast, the monobasic SARS-CoV FRET substrate was not processed by furin. The MERS-CoV S1/S2 FRET substrate possessing a dibasic R-X-X-R motif was cleaved by furin �-fold less efficiently than the best substrates of this FRET series. The FRET substrate SARS-CoV-2_M1, which contains an optimized furin recognition site by virtue of an A → K mutation in the P2 position, was cleaved with similar efficiency compared with the wild-type sequence. However, substitution of A → R in the P2 position strongly enhanced cleavage by furin. As expected, the analogous reference sequence of IBV was also processed by furin very efficiently. The data show that the R-R-A-R motif at the S1/S2 cleavage site of SARS-CoV-2 S is efficiently cleaved by furin in vitro.

(A) Fluorescence resonance energy transfer substrates of the S protein S1/S2 sites of the indicated CoVs. M1 and M2 are mutants of the SARS-CoV-2 S1/S2 site with substitution of A → K or A → R in P2 position. IBV, avian infectious bronchitis virus strain Beaudette. Cleavage by furin is indicated in red. (B) Cleavage of the fluorescence resonance energy transfer substrates (20 μM) by furin (0.5 nM). Cleavage efficiency of SARS-CoV-2_M2 was set as 100%. (C) Cleavage of SARS-CoV-2 S by furin and TMPRSS2 in HEK293 cells. Cells were co-transfected with pCAGGS-S-Myc-6xHis and either empty vector or pCAGGS-TMPRSS2. Cells were then incubated in the absence or presence of aprotinin or furin inhibitor MI-1851 (50 μM each) for 48 h. Cell lysates were subjected to SDS–PAGE and Western blot analysis using antibodies against the C-terminal Myc-tag. For each Western blot lanes are spliced together from one immunoblot from one experiment. β-actin was used as loading control.

Source data are available for this figure.

SARS-Cov-2 spike protein is cleaved by both furin and TMPRSS2

We next examined whether the SARS-CoV-2 S protein is cleaved by endogenous furin in HEK293 cells. Cells were transiently transfected with pCAGGS plasmid encoding the SARS-CoV-2 S protein with a C-terminal Myc-6xHis-tag and incubated in the absence and presence of the potent synthetic furin inhibitor MI-1851 (cf. Fig S1 manuscript describing its synthesis submitted). At 48 h post transfection, cell lysates were subjected to SDS–PAGE and Western blot analysis using antibodies against the Myc epitope. As shown in Fig 2C (left panel), the uncleaved precursor S and the S2 subunit were detected in the absence of MI-1851, indicating that S is cleaved by endogenous proteases at the S1/S2 site in HEK293 cells. In contrast, S cleavage was efficiently prevented by MI-1851. The S1 subunit cannot be detected by the Myc-specific antibody (cf. Fig 1A ). However, S cleavage was not prevented by the trypsin-like serine protease inhibitor aprotinin ( Fig 2C , right panel, lane 2). Thus, the data indicate that SARS-CoV-2 S protein is cleaved by furin at the S1/S2 site in HEK293 cells.

Aprotinin contains three disulfide bonds (indicated by lines).

We then investigated SARS-CoV-2 S cleavage by TMPRSS2. Because HEK293 cells do not express endogenous TMPRSS2 (unpublished data see also www.proteinatlas.org), we co-transfected the cells with pCAGGS-S-Myc-6xHis and pCAGGS-TMPRSS2. Then, the cells were incubated in the absence or presence of MI-1851 to suppress S cleavage by endogenous furin. Interestingly, two S cleavage products of � and 80 kD, respectively, were detected upon co-expression of TMPRSS2 in the absence of MI-1851 ( Fig 2C , left panel), most likely S2 and S2′, as they can both be detected by the Myc-specific antibody (cf. Fig 1A ). In the presence of MI-1851, only a minor S2 protein band was detected. However, the amount of S2′ protein present in transient TMPRSS2-expressing cells was similar in MI-1851–treated and untreated cells, suggesting that S cleavage at the S2′ site is only caused by TMPRSS2 activity. The small amount of S2 protein detected in TMPRSS2-expressing cells in the presence of MI-1851 was likely due to residual furin activity rather than cleavage of S at the S1/S2 site by TMPRSS2. Cleavage of S by TMPRSS2 at the S2′ site was further supported by the reduction of S2′ protein in aprotinin treated cells ( Fig 2C , right panel). Together, the data show that SARS-CoV-2 S can be cleaved by furin and by TMPRSS2. The data further suggest that the proteases cleave S at different sites with furin processing the S1/S2 site and TMPRSS2 cleaving at the S2′ site.

Knockdown of TMPRSS2 prevents proteolytic activation and multiplication of SARS-CoV-2 in Calu-3 human airway epithelial cells

Next, we investigated whether TMPRSS2 is involved in proteolytic activation and multicycle replication of SARS-CoV-2 in Calu-3 human airway epithelial cells. To specifically knockdown TMPRSS2 activity, we previously developed an antisense peptide-conjugated phosphorodiamidate morpholino oligomer (PPMO) (25). PPMOs are single-stranded nucleic acid–like compounds, composed of a morpholino oligomer covalently conjugated to a cell-penetrating peptide, and can interfere with gene expression by sterically blocking complementary RNAs. PPMOs are water-soluble and achieve entry into cells and tissues without assisted delivery (reviewed in references 26 and 27). The previously developed PPMO T-ex5 interferes with splicing of TMPRSS2 pre-mRNA, resulting in the production of mature mRNA lacking exon 5 and consequently expression of a truncated TMPRSS2 form that is enzymatically inactive. Using T-ex5 PPMO-mediated knockdown of TMPRSS2 activity, we were able to identify TMPRSS2 as the major influenza A virus activating protease in Calu-3 cells and primary human airway epithelial cells and of influenza B virus in primary human type II pneumocytes (25, 28).

Here, Calu-3 cells were treated once with T-ex5 PPMO for 24 h before infection with SARS-CoV-2 to inhibit the production of normal TMPRSS2-mRNA and deplete enzymatically active TMPRSS2 present in the cells. The cells were then inoculated with SARS-CoV-2 at a low MOI of 0.001, further incubated without additional PPMO treatment for 72 h, and then fixed and immunostained using a rabbit serum originally produced against 2002 SARS-CoV. As shown in Fig 3A , a strong cytopathic effect (CPE) and efficient spread of SARS-CoV-2 infection was visible in Calu-3 cells treated with a negative-control PPMO of nonsense sequence designated as “scramble” as well as untreated cells that were used as controls. In contrast, no CPE and only small foci of infection were observed in T-ex5 PPMO-treated cells at 72 h p.i. ( Fig 3A ). To examine SARS-CoV-2 activation and multicycle replication in PPMO-treated cells in more detail, Calu-3 cells were treated with PPMO for 24 h before infection, then inoculated with virus at an MOI of 0.001 for 1 h 30 min, and incubated for 72 h in the absence of further PPMO, as described above. At different time points, virus titers in supernatants were determined by tissue culture infection dose 50% (TCID50) end point dilution. T-ex5 PPMO treatment dramatically reduced virus titers in Calu-3 cells, by 500- and 2,000-fold at 16 and 24 h p.i., respectively, and 90-fold at 48 h p.i. ( Fig 3B ).

(A) Multicycle replication of SARS-CoV-2 in T-ex5–treated Calu-3 cells. Cells were treated with 25 μM T-ex5 or control PPMO (scramble) for 24 h or remained without treatment (w/o). Cells were then inoculated with SARS-CoV-2 at a MOI of 0.001 for 1 h 30 min, the inoculum was removed and the cells further incubated in the absence of PPMO for 72 h. Cells were fixed and immunostained using a serum against SARS-CoV. Virus-positive cells are stained in blue. Scale bars indicate 500 μm. (B) Calu-3 cells were treated with PPMO for 24 h and then infected with SARS-CoV-2 for 72 h as described above. Virus titers in supernatants were determined by tissue culture infection dose 50% (TCID50) end point dilution at indicated time points. Results are mean values ± SD of three independent experiments. (C) Analysis of TMPRSS2-mRNA in PPMO-treated Calu-3 cells. Cells were treated with 25 μM T-ex5, scramble PPMO or remained untreated (w/o) for 24 h (lanes 1𠄴). T-ex5–treated cells were inoculated with SARS-CoV-2 as described above and incubated in the absence of PPMO for 72 h (lane 4). Total RNA was isolated and analyzed by RT-PCR using primers designed to amplify 1,228 nt of full-length TMPRSS2-mRNA. Full-length and truncated PCR products lacking exon 5 are indicated by filled and open arrow heads, respectively. (D) Effect of PPMO treatment on Calu-3 cell viability. Calu-3 cells were treated with scramble or T-ex5 PPMO (25 μM) for 24 h. Cell viability of untreated (w/o) cells was set as 100%. Results are mean values ± SD (n = 3).

Source data are available for this figure.

To confirm knockdown of enzymatically active TMPRSS2 expression, Calu-3 cells were treated with PPMO or remained untreated for 24 h, after which TMPRSS2-specific mRNA was isolated and analyzed by RT-PCR as described previously (25). Total RNA was analyzed with primers designed to amplify nucleotides 108𠄱,336 of TMPRSS2-mRNA. A full-length PCR product of 1,228 bp was amplified from untreated and scramble PPMO-treated Calu-3 cells, whereas a shorter PCR fragment of about 1,100 bp was amplified from T-ex5 PPMO-treated cells ( Fig 3C ). Sequencing revealed that the truncated TMPRSS2-mRNA lacked the entire exon 5 (data not shown). To further confirm that T-ex5 PPMO single dose treatment before infection still interferes with TMPRSS2-mRNA splicing at 72 h p.i., total RNA was isolated from infected cells at 72 h p.i. and amplified as described above. As shown in Fig 3C , most TMPRSS2-mRNA amplified from T-ex5-treated cells at 72 h p.i. lacked exon 5. The data demonstrate that T-ex5 was very effective at producing exon skipping in TMPRSS2-pre-mRNA and, thus, at inhibiting expression of enzymatically active protease, during the virus growth period in Calu-3 cells. However, a small band of the full-length PCR product was visible after 72 h p.i., indicating low levels of expression of enzymatically active TMPRSS2 at later time points of the virus growth period, which may explain the increase in virus titers observed at 48 h p.i. (cf. Fig 3B ). Cell viability was not affected by T-ex5 PPMO treatment of Calu-3 cells, as shown in Fig 3D and described previously (25, 28).

Together, our data identify TMPRSS2 as a host cell factor essential for SARS-CoV-2 activation and multiplication in Calu-3 cells and show that down-regulation of TMPRSS2 activity dramatically blocks SARS-CoV-2 replication.

Inhibition of either TMPRSS2 or furin activity suppresses multicycle replication of SARS-CoV-2 in human airway epithelial cells

We next investigated the efficacy of different inhibitors of trypsin-like serine proteases, also inhibiting TMPRSS2, on preventing SARS-CoV-2 activation by TMPRSS2 in Calu-3 cells. We used the natural broad-range serine protease inhibitor aprotinin from bovine lung and two prospective peptide mimetic inhibitors of TMPRSS2, MI-432 (29), and MI-1900 ( Fig S1 ). Aprotinin has long been known to prevent proteolytic activation and multiplication of influenza A virus in cell cultures and mice. Furthermore, inhalation of aerosolized aprotinin by influenza patients markedly reduced the duration of symptoms without causing side effects (30). MI-432 was shown to efficiently inhibit proteolytic activation and multiplication of influenza A virus in Calu-3 cells (31). The inhibitor MI-1900 is a monobasic and structurally related analog of the dibasic inhibitor MI-432.

To examine the antiviral efficacy of the protease inhibitors against SARS-CoV-2, Calu-3 cells were infected with the virus at a low MOI of 0.001 for 1 h 30 min, after which the inoculum was removed and the cells incubated in the presence of the inhibitors at the indicated concentrations for 72 h. The cells were fixed and immunostained using an antiserum against 2002 SARS-CoV. As shown in Fig 4 , strong CPE, with holes visible throughout the monolayer, and efficient spread of SARS-CoV-2 infection was observed in Calu-3 cells in the absence of protease inhibitors. Spread of SARS-CoV-2 infection and virus-induced CPE was efficiently inhibited by aprotinin treatment in a dose-dependent manner and only a few virus-positive cells were visible in Calu-3 cultures treated with 20 and 50 μM aprotinin. Even at a lower concentration of 10 μM, the spread of SARS-CoV-2 was greatly reduced and CPE markedly prevented. Treatment with peptide mimetic TMPRSS2 inhibitors MI-432 and MI-1900 also strongly prevented SARS-CoV-2 multiplication and CPE in Calu-3 cells in a dose-dependent manner, although less potently than aprotinin. At 20 or 50 μM of MI-432 or MI-1900, only small foci of infection were visible. At a concentration of 10 μM, virus spread and CPE in MI-432– or MI-1900–treated cells were still reduced compared with control cells. The data demonstrate that SARS-CoV-2 multiplication in Calu-3 human airway cells can be strongly suppressed by aprotinin and the synthetic TMPRSS2 inhibitors MI-432 and MI-1900.

Calu-3 cells were inoculated with SARS-CoV-2 at a low MOI of 0.001 and then incubated in the presence of inhibitors of TMPRSS2 (aprotinin, MI-432, and MI-1900), furin (MI-1851), and endosomal cathepsins (E64d), respectively, at the indicated concentrations. Cells were fixed and immunostained using a rabbit serum against SARS-CoV at 72 h p.i. Virus-positive cells are stained in blue or dark gray depending on the staining intensity. Cells infected in the absence of inhibitors (w/o), in the presence of DMSO (0.5%) and noninfected cells (mock) were used as controls. Scale bars indicate 500 μm. Images are representatives of three independent experiments.

The observed efficient cleavage of transient expressed SARS-CoV-2 S protein by furin in HEK293 cells prompted us to investigate if furin is involved in SARS-CoV-2 activation in Calu-3 cells. Hence, virus spread in Calu-3 cells was analyzed in the presence of the furin inhibitor MI-1851. Interestingly, MI-1851 strongly inhibited SARS-CoV-2 spread at even the lowest concentration of 10 μM, indicating that furin is critical for SARS-CoV-2 activation and multiplication in these cells ( Fig 4 ). Finally, to examine whether endosomal cathepsins are involved in SARS-CoV-2 activation in Calu-3 cells, multicycle virus replication was determined in the presence of the cathepsin inhibitor E64d. Cathepsin L was shown to cleave the S protein of 2002 SARS-CoV S close to the S1/S2 site (R667) at T678 in vitro (2). Here, strong CPE and foci of infection were observed in E64d-treated cells even at the highest dose of 50 μM, similar to that observed in DMSO-treated as well as untreated control cells, indicating that SARS-CoV-2 activation in Calu-3 cells is independent of endosomal cathepsins.

In sum, our data demonstrate that inhibition of either TMPRSS2 or furin strongly inhibits SARS-CoV-2 in Calu-3 human airway cells, indicating that both proteases are critical for S activation. In contrast, endosomal cathepsins are dispensable or not involved at all in SARS-CoV-2 activation in these cells.

Growth kinetics of SARS-CoV-2 in protease inhibitor–treated Calu-3 cells

To analyze inhibition of SARS-CoV-2 activation and multiplication by the different protease inhibitors in more detail, we performed virus growth kinetics in inhibitor-treated cells. Calu-3 cells were inoculated with SARS-CoV-2 at an MOI of 0.001 and then incubated in the presence of 10 or 50 μM of the different protease inhibitors. At 16, 24, 48, and 72 h p.i., the viral titer in supernatants was determined by TCID50 titration. Untreated cells and cells treated with DMSO alone were used as controls. SARS-CoV-2 replicated to high titers within 24� h in untreated and DMSO-treated cells Calu-3 cells ( Fig 5A ). Aprotinin suppressed virus replication 25- to 100-fold compared with control cells even at a concentration of 10 μM at 16-48 h p.i. The TMPRSS2 inhibitor MI-432 only slightly affected virus replication at a concentration of 10 μM but reduced virus titers 75-fold at 24 h p.i. at 50 μM. Treatment of cells with TMPRSS2 inhibitor MI-1900 reduced virus titers in a dose-dependent manner and caused strong inhibition of SARS-CoV-2 replication at 50 μM with 35- to 280-fold reduced viral titers compared with control cells. The furin inhibitor MI-1851 efficiently suppressed SARS-CoV-2 multiplication in Calu-3 cells, producing a 30- to 190-fold reduction in virus titers at a dose of 10 μM. In contrast, virus multiplication was not affected by treatment with the cathepsin inhibitor E64d, consistent with the data shown in Fig 4 . To provide evidence that inhibition of SARS-CoV-2 replication in inhibitor-treated cells was not caused by cytotoxic effects, we analyzed cell viability in Calu-3 cells treated with 50 μM of the different inhibitors for 72 h. As shown in Fig 5B , evaluation of cell viability revealed no significant cytotoxicity by any of the inhibitors under conditions used in the virus growth experiments.

(A) Calu-3 cells were inoculated with SARS-CoV-2 at a low MOI of 0.001 and then incubated in the absence (w/o) or presence of inhibitors of TMPRSS2 (aprotinin, MI-432, and MI-1900), furin (MI-1851), and endosomal cathepsins (E64d), respectively, or DMSO (0.5%), at the indicated concentrations. At 16, 24, 48, and 72 h postinfection (p.i.), supernatants were collected, and virus replication was determined by tissue culture infection dose 50% (TCID50) titration at indicated time points. Data are mean values ± SD of three to five independent experiments. (B) Effect of inhibitor treatment on cell viability. Calu-3 cells were treated with the indicated protease inhibitor (50 μM) for 72 h. Untreated cells (w/o) and DMSO treated cells were used as controls. Cell viability of untreated cells was set as 100%. Results are mean values ± SD (n = 3). (C) Antiviral activity of combinations of TMPRSS2 and furin inhibitors against SARS-CoV-2 in human airway epithelial cells. Calu-3 cells were inoculated with SARS-CoV-2 at an MOI of 0.001 as described above and then incubated in the presence of single protease inhibitors or inhibitor combinations at the indicated concentrations. Virus titers in supernatants were determined by TCID50 at 16, 24, 48, and 72 h p.i. Data are mean values ± SD of three independent experiments. (D) Calu-3 cells were treated with PPMO for 24 h, then infected with SARS-CoV-2 as described above and incubated in the absence of PPMO (w/o, scramble and T-ex5) and with or without 50 μM of furin inhibitor treatment (MI-1851) for 72 h. At 16, 24, 48, and 72 h p.i., supernatants were collected, and viral titers were determined by TCID50 at indicated time points. Data are mean values ± SD (n = 2).

The data demonstrate that SARS-CoV-2 replication can be efficiently reduced by inhibiting either TMPRSS2 or furin activity, demonstrating that both proteases are crucial for SARS-CoV-2 activation.

Treatment of SARS-CoV-2–infected Calu-3 cells with a combination of TMPRSS2 and furin inhibitors

Finally, we wished to examine whether the combination of inhibitors against TMPRSS2 and furin shows a synergistic antiviral effect. Therefore, Calu-3 cells were infected with virus as described above and incubated in the presence of aprotinin, MI-432, or MI-1900 in combination with MI-1851 at 10 and 50 μM each, respectively, for 72 h. Virus titers in supernatants were determined at indicated time points. Single-dose treatment of each inhibitor and untreated cells were used as controls. As shown in Fig 5C , the combination of 10 μM of MI-1851 with either aprotinin or MI-432 showed enhanced antiviral activity against SARS-CoV-2 and 10- to 30-fold reduced virus titers compared with 10 μM of each inhibitor alone and also reduced viral titer four to eightfold more than that observed with either of the single inhibitors at 50 μM. A combination of 50 μM each of MI-432 and MI-1851 reduced virus titers 10- to 32-fold compared with 50 μM of each inhibitor alone and thereby dramatically suppressed SARS-CoV-2 multiplication 100- to 250-fold compared with untreated or DMSO-treated cells. In contrast, treatment of Calu-3 cells with 50 μM each of MI-1851 and aprotinin did not cause further suppression of virus titers compared with the combination of 10 μM of each inhibitor. The combination of 10 μM each of MI-1851 and MI-1900 did not show enhanced antiviral activity compared with single inhibitor treatments at 10 μM. However, treatment of cells with 50 μM each of MI-1900 and MI-1851 caused fivefold reduction in viral titers when compared with cells treated with 50 μM of each inhibitor alone and thereby SARS-CoV-2 multiplication in Calu-3 cells was markedly reduced compared to control cells. We furthermore examined the antiviral activity of a combination of T-ex5 PPMO and furin inhibitor MI-1851 against SARS-CoV-2 in Calu-3 cells. As shown in Fig 5D , combined treatment of Calu-3 cells with 25 μM T-ex5 PPMO and 50 μM MI-1851 almost completely blocked SARS-CoV-2 replication with a nearly 15,000-fold reduction in virus titers at 24 h p.i., and reduced virus titers 500-fold at 48 h p.i. compared with control cells. Combination of T-ex5 and MI-1851 was synergistic and caused 30- to 10-fold lower virus titers at 16 and 24 h p.i. compared with single inhibitor-treated cells. The data demonstrate that efficient inhibition of S cleavage by a combination of TMPRSS2 and furin inhibitors can dramatically block SARS-CoV-2 replication in human airway epithelial cells. Furthermore, our data show that combination of TMPRSS2 and furin inhibitors can act synergistically to produce inhibition of SARS-CoV-2 activation and multiplication at lower doses than single protease inhibitor treatment.

In conclusion, our data demonstrate that both TMPRSS2 and furin cleave the SARS-CoV-2 S protein and are essential for efficient virus multicycle replication in Calu-3 human airway cells. The results indicate that TMPRSS2 and furin cleave S at different sites𠅏urin at the S1/S2 site and TMPRSS2 at the S2′ site𠅊nd suggest that TMPRSS2 and furin cannot compensate for each other in SARS-CoV-2 S activation. Our data further demonstrate that inhibition of either one of these critical proteases can render the S protein of SARS-CoV-2 unable to efficiently mediate virus entry and membrane fusion and, therefore, provides a promising therapeutic approach for treatment of COVID-19.


Results

Role of Mpl and the PLCs in Mouse Virulence and Plaque Formation

A series of deletions was generated into the PLC-, Mpl-, and ActA-encoding genes to evaluate the requirements for proPC-PLC proteolytic activation. The mutations were introduced into the chromosome of L. monocytogenes strains SLCC-5764 and 10403S. Proteolytic activation of proPC-PLC was first evaluated in vitro, using SLCC-5764 and isogenic mutant strains. SLCC-5764 is a hypersecreting strain of L. monocytogenes that facilitates in vitro analysis of virulence factors, such as ActA and PC-PLC, not normally expressed in vitro (Camilli et al., 1993). ProPCPLC (Mr 33 kD) and the processed form of the enzyme (Mr 28 kD) were identified by Coomassie blue staining and Western immunoblotting. ProPC-PLC was secreted in comparable amounts from each plcB positive strain (Fig. 1, a and b), indicating that the internal deletions in mpl and actA genes, which are located within the same operon and upstream of the plcB gene (Portnoy et al., 1992), did not affect plcB gene expression. Similarly, the internal deletion in plcA, which is located within the same operon and upstream of prfA, the transcriptional regulator encoding gene (Portnoy et al., 1992), did not affect plcB gene expression (Fig. 1, a and b, lane 3). The processed form of PC-PLC was secreted in comparable amounts from SLCC5764 and isogenic plcA and actA mutant strains (Fig. 1, a and b, lanes 2, 3, and 5). Enzymatic activity, as determined by the egg yolk gel overlay assay, comigrated with the processed form of PC-PLC (compare Fig. 1,b to 1 c, lanes 2, 3, and 5), confirming that PI-PLC and ActA are not required for the proteolytic activation of proPC-PLC in vitro and that the processed form is the active form of the enzyme. A processed form of PC-PLC was detected in minute amounts from the mpl mutant strain (Fig. 1, a and b, lane 6). However, there was no detectable PC-PLC activity associated with this mutant (Fig. 1 c, lane 6), even after increasing the amount of protein loaded on the gel fourfold (data not shown). This result is consistent with Mpl being required for activation of proPC-PLC in broth culture as previously reported (Poyart et al., 1993).

The virulence of wild-type strain 10403S and isogenic mutants was evaluated in vivo using the mouse infection model. The ability of these mutants to spread cell to cell was evaluated in a mouse fibroblast cell line by a plaquing assay. The results are reported in Fig. 2 and Table II. The single plcA and plcB mutants show respective reductions of 12% and 33% in plaque sizes, and the double plcA, plcB mutant shows a reduction of 66% in plaque size. In the mouse infection model, the plcA mutant shows a minor increase in LD50, the plcB mutant is 1 log less virulent, whereas the double plcA, plcB mutant is 2.5 logs less virulent than the wild-type strain. These results are consistent with previously reported data, suggesting that the PLCs have overlapping functions (Smith et al., 1995). The double mpl, plcB mutant was phenotypically identical to the single plcB mutant, consistent with Mpl function being related to the activation of proPC-PLC. Similarly, the triple plcA, plcB, mpl mutant was phenotypically identical to the double plcA, plcB mutant. A single mutation in the mpl gene resulted in a 29% decrease in plaque size, also consistent with Mpl being required for proPC-PLC activation. However, the mpl mutant was as virulent as the wild-type strain in the mouse infection model, which is inconsistent with Mpl function being required for the activation of proPC-PLC. In addition, a double plcA, mpl mutant had a 47% decrease in plaque size, which differs significantly (P < 0.0001) from the 66% decrease observed with the double plcA, plcB mutant. The double mpl, plcA mutant was ∼2 logs less virulent than wild type but not as attenuated as the double plcA, plcB mutant in the mouse infection model. Taken together, these data clearly show a role for Mpl in bacterial cell-to-cell spread and reveal a role for Mpl in virulence as seen in the double mpl, plcA mutant. However, unlike the plcB mutant, the virulence of the single mpl mutant was not attenuated in mice, and that of the double mpl, plcA mutant was not as attenuated as the double plcA, plcB mutant. These results raised the possibility that intracellular activation of proPC-PLC may proceed in the absence of Mpl.

Intracellular Activation of ProPC-PLC

To directly evaluate the role of Mpl in the intracellular activation of proPC-PLC, we examined the production and processing of proPC-PLC in a tissue-culture model of infection. At 4 h after infection, J774 cells were pulse labeled for 30 min with [ 35 S]methionine and lysed, and both forms of PC-PLC were immunoprecipitated using affinity-purified antibodies. The precursor and processed forms of PCPLC were identified based on their relative migration on SDS-PAGE, and by comparison with cells infected with the plcB mutant strain. Both forms of the enzyme were present in cells infected with either the wild-type strain or the mpl mutant (Fig. 3, lanes 1 and 3), indicating that Mpl was not essential for intracellular processing of proPCPLC. Furthermore, results from an egg yolk gel overlay assay from infected cells indicated that the processed form of PC-PLC generated in cells infected with either the wildtype strain or the mpl mutant had enzymatic activity (Fig. 4, lanes 2 and 5). Phospholipase activity was not detected in uninfected cells or cells infected with the plcB mutant (Fig. 4, lanes 1 and 3).

Subcellular Localization of PC-PLC

Double immunofluorescence staining and confocal microscopy was performed at 5 h after infection to determine the site of PC-PLC and L. monocytogenes localization. Distinctively, only a small proportion of intracellular bacteria stained positively for PC-PLC, and PC-PLC staining colocalized with L. monocytogenes (Fig. 5,A). The pattern of PC-PLC staining suggested a vacuolar localization. Double immunofluorescence staining of PC-PLC and Lamp1 was performed to define more precisely the site of PC-PLC localization. Lamp1 is an endosomal/lysosomal marker (Chen et al., 1985 Lewis et al., 1985) that increases in concentration with endosomal maturation (Berón et al., 1995 Pitt et al., 1992). Colocalization of PC-PLC and Lamp1 was observed (Fig. 5, D and E), but the amount of Lamp1 colocalizing with PC-PLC varied considerably. This staining pattern suggested that PC-PLC was concentrated in vacuoles that had fused with endosomes and lysosomes. PC-PLC staining was not observed in filopodia-like structures, but we cannot eliminate the possibility that this could be due to a lack of sensitivity of the technique used.

To directly address whether PC-PLC localized to secondary vacuoles formed upon bacterial cell-to-cell spread, J774 cells were infected with a 10-fold lower multiplicity of infection to facilitate identification of primary and secondary infected cells at 5 and 6 h after infection. Double immunofluorescence staining of PC-PLC and L. monocytogenes indicated that the majority of PC-PLC localized to cells at the periphery of infected foci, presumably in secondary vacuoles formed during bacterial cell-to-cell spread. An example of an infection focus at 6 h after infection is shown in Fig. 5 B.

The Mpl-dependent and -independent Pathways of ProPC-PLC Proteolytic Activation

The above results suggested that secondary vacuoles formed during bacterial cell-to-cell spread fused with vesicles of the endocytic pathway (Berón et al., 1995 Gruenberg and Maxfield, 1995 Pitt et al., 1992). To assess the role of vacuolar acidification and lysosomal enzymes in proPC-PLC processing, we used two enzyme inhibitors. The first, bafilomycin A1, is a specific inhibitor of the vacuolar proton pump ATPase (Yoshimori et al., 1991), which serves to acidify a vacuolar compartment. The second is a peptidyldiazomethane, Z-FA-CHN2, which specifically inactivates cysteine proteases by alkylation of the reactive site cysteine residue (Leary et al., 1977). Z-FA-CHN2 has strong affinity for cathepsins B and L (Crawford et al., 1988 Kirschke and Shaw, 1981 Wilcox and Mason, 1992), which are lysosomal acid cysteine proteases, but does not react with the calpains (Crawford et al., 1988), which are cytosolic calcium-activated neutral cysteine proteases. The results showed that both bafilomycin A1 and Z-FA-CHN2 blocked processing of proPC-PLC in cells infected with the mpl mutant (Fig. 6, lanes 5 and 6). In wild-type infected cells, processing of proPC-PLC was not affected by Z-FACHN2, but no processing was detected in cells treated with bafilomycin A1 (Fig. 6, lanes 2 and 3), although the protein was detected by immunofluorescence in secondary vacuoles (data not shown). Neither bafilomycin A1 nor Z-FACHN2 inhibited activation of proPC-PLC in broth culture (data not shown). These results indicate that there are two intracellular pathways of activation of proPC-PLC: an Mpl-mediated pathway and a cysteine protease-mediated pathway. Both pathways were blocked by a specific inhibitor of the vacuolar proton pump ATPase, suggesting that vacuolar acidification is a prerequisite to the intracellular processing of proPC-PLC.

Intracellular Activity of PC-PLC on Phosphatidylcholine and Sphingomyelin

Intracellular activation of proPC-PLC was mediated by either Mpl or a cysteine protease, raising the possibility that PC-PLC activity might vary depending on the activating protease. To address that point, we investigated the activities of PC-PLC generated in cells infected with either the wild-type strain or the mpl mutant. PC-PLC was immunoprecipitated from infected cells, and the enzymatic assay was performed directly on PC-PLC bound to antibodies on protein A–Sepharose beads. Hydrolysis of 3 H-PC and 14 C-sphingomyelin by immunoprecipitated PC-PLC was measured as described in Materials and Methods. In either case, PC-PLC was capable of mediating PC and sphingomyelin hydrolysis (Table III). When corrected for the number of bacteria per dish, the ability of PC-PLC generated by the wild-type strain and the mpl mutant to hydrolyze PC and sphingomyelin was essentially the same, although a small shift in substrate preference was observed. Phospholipase activity was not detected on immunoprecipitates from uninfected cells or cells infected with the plcB mutant (data not shown).

Proteolytic Activation of ProPC-PLC in the Absence of Bacterial Cell-to-Cell Spread

The above results indicated that PC-PLC localized to Lamp1-positive vacuoles (Fig. 5) and that vacuolar acidification was a prerequisite to proPC-PLC activation (Fig. 6, lanes 2 and 5). These results suggested that active PC-PLC would not be generated in the cytosol of infected cells. To further investigate the intracellular requirements for proPCPLC activation, we monitored the presence of PC-PLC in cells infected with an actA mutant of L. monocytogenes that is defective in actin-based motility, and consequently fails to spread cell to cell. ProPC-PLC was immunoprecipitated in similar amounts in cells infected with either the wild-type strain or the actA mutant, indicating that proPCPLC was synthesized by cytosolic bacteria, but processing of proPC-PLC, although observable, was very inefficient in the cytosol of cells infected with the actA mutant (Fig. 7, compare lanes 1 and 5).

To eliminate the possibility that ActA was directly involved in proPC-PLC processing, we infected cells with the wild-type strain, and then blocked bacterial cell-to-cell spread by adding cytochalasin D (Dabiri et al., 1990 Tilney and Portnoy, 1989), an inhibitor of actin polymerization, 30 min before labeling. Again, the precursor form of PC-PLC was present in these infected cells but the processed form was not detectable, indicating that ActA was not directly responsible for proPC-PLC processing (Fig. 7, lane 4). Moreover, the amount of processed PC-PLC was largely reduced when actin polymerization was blocked as late as 10 min into the pulse, and it was barely detectable when actin polymerization was blocked 10 min before labeling (Fig. 7, lanes 2 and 3). Results from the cytochalasin D time course experiment indicated that continuous bacterial spreading was required for proPC-PLC processing. Not surprisingly, PC-PLC activity, as measured by the egg yolk gel overlay assay, was detected neither in cells infected with the actA mutant strain (Fig. 4, lane 4) nor in wild-type infected cells treated with cytochalasin D (data not shown). Therefore, proPC-PLC proteolytic activation, but not synthesis, occurred predominantly in secondary vacuoles.

Cytosolic Degradation of ProPC-PLC

Our observations indicated that proPC-PLC proteolytic activation was inefficient in the absence of bacterial cellto-cell spread (Fig. 7). However, in the absence of proteolytic activation, proPC-PLC did not appear to accumulate intracellularly, suggesting that proPC-PLC was rapidly degraded when secreted in the cytosol of the host cell. Proteolytic degradation of cytosolic proteins is mostly proteasome dependent (Rock et al., 1994). We investigated the role of the proteasome in the intracellular degradation of proPC-PLC using two aldehyde tripeptide inhibitors: LLnL and LLM. LLnL blocks the proteasome activity, while LLM does not (Rock et al., 1994 Vinitsky et al., 1992).

The intracellular stability of proPC-PLC was evaluated by a pulse-chase experiment. Cells infected with ActA − bacteria were treated with either LLnL or LLM at 3 h after infection and the inhibitors were present during the entire pulse-chase experiment. At 3 h and 50 min after infection, cells were pulse labeled with [ 35 S]methionine for 10 min, and then chased for specific periods of time. A role for the proteasome in proPC-PLC degradation was demonstrated by the observation that proPC-PLC was stabilized in infected cells treated with LLnL (Fig. 8, compare lanes 5–8 to lanes 1–4), but not in those treated with LLM (Fig. 8, lanes 9–12). Indeed, the half-life of proPC-PLC was <15 min in either untreated or LLM-treated cells, and >60 min in LLnL-treated cells. The intracellular stability of proPC-PLC was comparable in cells infected with either the wild-type strain, the mpl mutant, or the actA mutant (data not shown). Yet, in wild-type and Mpl − infected cells, proPC-PLC chased into active PC-PLC (data not shown). These results suggested an additional level of PC-PLC regulation by proteolytic degradation of the precursor. ProPC-PLC secreted into the cytosol was degraded by host proteases and consequently had a short half-life.


Nesfatin-1

Processing of the Precursor

The NUCB2 protein contains several cleavage recognition sites for prohormone convertases (PC). The proteolytic processing of NUCB2 by PC-1/3 is assumed to generate several products including nesfatin-1 (amino acids 1–82), nesfatin-2 (amino acids 85–163) and nesfatin-3 (amino acids 166–396, Fig. 1 B). 22 In their landmark study, Oh-I and colleagues reported the presence of the 82 amino acid peptide in rat cerebrospinal fluid, whereas the mature peptide was not detectable in hypothalamic tissue. 22 Subsequent reports did not detect mature nesfatin-1 (9.7 kDa) in the brain 9,10 or plasma, 29 whereas full length NUCB2 as well as small amounts of exogenous synthetic nesfatin-1 were detectable by Western blot. 29 So far, most studies describing the expression at the protein level did not distinguish between NUCB2 and nesfatin-1 as the antibodies used recognized both molecules. Because of these findings, it remains to be clarified whether nesfatin-1 or full length NUCB2 is the biologically active molecule since NUCB2 also shows biological activity upon third ventricular injection in rats. 22 Among the known cleavage products, only nesfatin-1 influences food intake, whereas nesfatin-2 and nesfatin-3 do not. 22


Interferons: Meaning, Production and Applications

Interferons are natural glycoproteins produced by virus-infected eukaryotic cells which protect host cells from virus infection. They were discovered by Isaacs and Lindenmann in 1957 in course of a study of the effect of UV-inactivated influenza virus on chick chorioallantoic membrane kept in an artificial medium.

They observed that the infected membrane produced a soluble substance in the medium which could inhibit the multiplication of active influenza virus inoculated in fresh chick chorioallantoic membranes. The substance was called interferon because it interfered with intra­cellular multiplication of viruses.

Later observations confirmed that such host-produced antiviral substances were common to many viruses. Viral interference is a phenomenon observed when multiplication of one virus is inhibited by another virus. For instance, when influenza-A virus is inoculated into the allantoic cavity of an embryonated egg followed after 24 hr by influenza-B virus, the multiplication of influenza-B virus is partly or completely inhibited. The reason why influenza-B virus cannot multiply is that the influenza-A virus infected cells produce interferon which partly or totally inhibits multiplication of B virus. The interferon also protects cells from influenza A virus.

Characteristics of Interferons:

An outstanding feature of interferons is that they are host-cell-specific and not virus-specific. This means that interferons produced by mouse or chicken will not protect human cells against the same virus which induced interferon in the experimental animals. On the other hand, an interferon produced by a virus X in an animal will protect the animal also from other viruses.

This is because interferons do not interact directly with the viruses. But they induce the virus infected cells to synthesize antiviral proteins which inhibit viral multiplication. These proteins have a wide inhibitory spectrum. As a result, not only the interferon-inducing virus, but others are also inhibited.

The reason why interferon produced by one species does not protect another species is that the same virus produces different interferons in different species. It has been observed that interferons produced by different host species following infection by the same virus differ in molecular weight as well as in other properties, like isoelectric point etc. Not only different species produce different interferons, different tissues of the same animal produce different interferons.

All types of interferons are proteins having a comparatively low molecular weight ranging between 15,000 to 40,000 Daltons. Hence, they are non-dialyzable and destroyed by proteolytic enzymes. Interferons are fairly stable at low pH (pH2) and can withstand moderate temperature being stable at 37°C for an hour or so. They are produced in minute amounts by the infected cells as a longer precursor having 23 amino acid residues more than the mature molecule.

Human interferons are of three main types. These are called alpha interferons (α-IFN), beta-interferons (β-IFN) and gamma-interferons (γ-IFN). Alpha-interferon contains many subtypes. The total subtypes exceed 20 in number.

It is produced by the B-lymphocytes, monocytes and macrophages. β-IFN is produced by the fibroblasts in the connective tissues. γ-IFN is synthesized by the T-lymphocytes after they are activated by antigens. α-IFN has been shown to be coded by as many as 20 distinct chromosomal genes, indicating thereby that the subtypes of this interferon represent a family of closely related proteins.

β-IFN appears to be a glycoprotein. It is coded by a single human gene. All the genes of α-IFN and β-IFN are located on the short arm of human chromosome 9. α-IFN proteins are all 166 amino acid long (except one). They are non-glycosylated and the proteins are monomeric. The single β-IFN protein is also 166 amino acid long and a glycoprotein. It is dimeric.

Production of Interferons:

Interferons are produced by living animal cells, both in vivo as well as cultured cells. Interferon production and its antiviral activity require expression of cellular genes, and these functions are blocked by inhibitors of transcription and translation. Thus, virus-infected host cells fail to produce interferon in presence of actinomycin D, an inhibitor of eukaryotic RNA polymerase. When the inhibitor is added after 2 hr of infection, interferon production is not inhibited, suggesting that transcription is completed by that time.

Interferon production starts after initiation of viral maturation and continues for 20 to 50 hr after that. Then the production stops, due to formation of a repressor which presumably is formed or activated only when the interferon concentration in the producing cell exceeds a certain threshold concentration. Most of the interferon is transported from the producing cell to other neighbouring cells.

The substance in a virus that is responsible for interferon synthesis by the host cell is known as interferon inducer. The nature of this substance was identified by Merigan (1970) as double-stranded RNA. The activity seems to reside in polyribonucleotide’s with a high helical content. The double- stranded RNA viruses — like reoviruses — can act as interferon inducer without replication. Single- stranded RNA viruses can act as inducers only after replication when they form double-stranded replicative intermediates. DNA-viruses can also induce interferons, presumably due to overlapping transcription of viral DNA as observed in case of vaccinia vinus (Fig. 6.39).

Fungal viruses which have mostly double-stranded RNA genomes are also efficient inducers of interferons. Some synthetic polymers containing riboinosinic acid, ribocytidylic acid (Poly I: C) as well as those containing riboadenylic acid and ribouridylic acid (Poly A: U) are also good inducers. All interferon inducers are characterized by high molecular weight, high density of anionic groups and resistance to enzymatic degradation. DNA and DNA-RNA hybrids have been found to be ineffective as interferon inducers.

The induction of interferon synthesis concerns α- and β-interferon’s which belong to a single class, called Type I. Gamma-interferon belongs to a separate class, called Type II. The human Υ-interferon is the single representative of its type. The gene coding the y-interferon protein is located on the long arm of chromosome 12. The gene has three introns, while the genes of α- and β- interferons are without any introns. Gamma-interferon (human) has 146 amino acids and is an N-glycosylated tetrameric protein. It is induced by antigenic stimulation of T-lymphocytes.

In presence of the inducer which is viral ds-RNA, the α- and β-interferon genes of the host chromosome(s) are activated to produce interferon m-RNAs. Those are then translated intoα- and β- interferon proteins. These proteins at first accumulate in the producing cell and eventually leave the cell to reach neighbouring host cells.

As the interferon concentration in the producing cell rises above a threshold level, it activates another gene of the producing cell which codes for a repressor protein which feeds back and stops further synthesis of interferon. As a result, virus-infected cells generally produce only small quantities of interferons.

The interferon molecules that leave the producing cell reach the neighbouring uninfected host cells and interact with the cell membrane or nuclear membrane receptors of these cells. Thereby these cells are induced to synthesise antiviral proteins. These antiviral proteins are the actual agents that provides protection to these host cells against viral infection.

Mechanism of Action of Interferons:

Type I interferons include α-IFN and β-IFN. These interferons do not interact with the viruses directly causing their inhibition, but they induce the formation of antiviral proteins which are activated to inhibit viral multiplications. These interferon-regulated proteins (IRPs) act presumably by blocking synthesis of the macromolecular components necessary for viral multiplication.

A general scheme for mechanism of action of type I interferons is shown in Fig. 6.40:

Several interferon regulated host proteins (IRPs) have been identified, though all of them have not been fully characterized. Among the better known of these proteins are a protein kinase and an enzyme catalyzing the formation of a short polymer of adenylic acid, the 2′, 5′-oligoadenylate synthetase (2′-5′ A synthetase).

The protein kinase is induced by Type I interferons. It has to be activated by ds-RNA. The activated kinase catalyses phosphorylation of initiation factor (el F-2) thereby causing inhibition of protein synthesis (Fig. 6.41).

The 2′-5′-oligoadenylate synthetase is an enzyme also induced by Type I interferons which requires activation by ds-RNA like the protein kinase. The activated synthetase acts as an activator of an endonuclease, RNase L. The activated RNAse degrades viral ss-RNA (Fig. 6.42).

Another group of proteins, called Mx-proteins induced by α- and β-IFN are known to possess intrinsic antiviral activity, although the exact molecular mechanism by which they inhibit viral multiplication is not known. Mx-proteins have been reported to play a major controlling role in infections caused by influenza viruses in experimental animals as well as in humans.

Type II interferon includes g-IFN which is also known as immune IFN. Although g-IFN also possesses anti-viral activity, its major role is in the immunity through activation of cytotoxic T-lymphocytes which can destroy virus infected cells. Besides T-lymphocytes, other naturally occurring killer cells like macrophages and monocytes are also activated by g-IFN. Thus, in contrast to that of Type I interferons, the antiviral effect of g-IFN is expressed through activating the killer cells of the body which destroy the virus-infected cells.

Type II interferon induces the major histocompatibility antigens of human cells. Expression of these antigens is essential for immuno-potent cells to present foreign antigens to the T-lymphocytes during generation of specific immune responses.

IFN induced expression of these major histocompatibility antigens represents an important contribution of the antiviral activity of g-IFN through enhancement of the activity of cytotoxic T-lymphocytes. The activation of cytotoxic T-lymphocytes by y-IFN also implies its possible role in elimination of cancer cells which are recognized by the immune system of the body as foreign objects.

Applications of Interferons:

Interferons could be ideal agents for combating viral diseases. They inhibit viral multiplication at such low concentration which is non-toxic to uninfected cells. One interferon can inhibit many viruses. But there are certain draw-backs which stand in their use.

Firstly, for application in humans, interferon must be of human origin, though interferons produced in monkey kidney cell cultures are also effective in humans. Interferons are produced in very small quantities and it is difficult to get them in sufficient quantity in pure form for clinical application. Another factor is that interferons are effective only for short periods and as such can be used against only acute infections, like influenza.

The difficulty of obtaining sufficient quantity of pure interferon for clinical use has been overcome by cloning the α-IFN and β-IFN human genes in bacteria and yeast. By growing these transgenic organisms in mass culture, it has been possible to obtain clinically usable interferons in sufficiently large quantities. Alpha-interferon has been marketed in 1984 under the trade name Intron A.

In the following years, this biotechnologically produced interferon has been approved for clinical use against diseases like genital herpes caused by herpes-virus, hepatitis B and C. Beta-interferon has also been biotechnologically produced and marketed under the trade name Betaseron. It has been used in a disease called multiple sclerosis. A recombinant g-interferon has been found effective against an inherited chronic disease, called granulomatous disease.

The neutrophils of the affected individual are unable to kill the infectious bacteria. Application of y-IFN to such persons restores the ability of the neutrophils to kill bacteria. As the disease is chronic and inherited, the affected persons must take g-IFN throughout their life to remain normal.

Interferons are not only antiviral, but they have also anticancer activity. Clinical trials have shown that interferons have effect against only some types of tumours. Alpha-interferon has been approved for treating hairy-cell leukemia, and Kaposi’s sarcoma, a cancer that occurs in AIDS patients.

Gamma-interferon has been mainly used as an immuno-stimulant in cancer patients. Resistance against tumours in the body is controlled by the immune response against tumour antigens. The cytotoxic T-lymphocytes recognize these antigens as foreign and destroy them. Gamma-interferon can stimulate the cytotoxic function of T-lymphocytes and other natural killer cells of the body, thereby helping to control the tumour cells.


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Affiliations

Institute of Molecular Virology and Cell Biology, Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Greifswald-Insel Riems, Germany

E. M. Abdelwhab, Jutta Veits & Thomas C. Mettenleiter

Department of Experimental Animal Facilities and Biorisk Management, Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Greifswald-Insel Riems, Germany

Reiner Ulrich & Jens P. Teifke

Institute of Epidemiology, Friedrich-Loeffler-Institut, Federal Research Institute for Animal Health, Greifswald-Insel Riems, Germany



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