Abstract
Protein Phosphatase 1 (PP1) is a major serine/threonine phosphatase in eukaryotes, participating in several cellular processes and metabolic pathways. Due to their low substrate specificity, PP1’s catalytic subunits do not exist as free entities but instead bind to Regulatory Interactors of Protein Phosphatase One (RIPPO), which regulate PP1’s substrate specificity and subcellular localization. Most RIPPOs bind to PP1 through combinations of short linear motifs (4–12 residues), forming highly specific PP1 holoenzymes. These PP1-binding motifs may, hence, represent attractive targets for the development of specific drugs that interfere with a subset of PP1 holoenzymes. Several viruses exploit the host cell protein (de)phosphorylation machinery to ensure efficient virus particle formation and propagation. While the role of many host cell kinases in viral life cycles has been extensively studied, the targeting of phosphatases by viral proteins has been studied in less detail. Here, we compile and review what is known concerning the role of PP1 in the context of viral infections and discuss how it may constitute a putative host-based target for the development of novel antiviral strategies.
Similar content being viewed by others
Introduction
Viruses are completely dependent on the host cell machinery [1] and manipulate diverse physiologic and metabolic host pathways to favour infection and ensure the proper formation of new infectious viral particles. Numerous studies have reported that protein phosphorylation is crucial for several steps of the life cycle of different viruses and that specific host protein kinases are hijacked by viruses to promote infection [2,3,4]. In fact, these proteins have been pinpointed as potential host-directed targets for the development of antiviral therapeutics (reviewed by García-Cárceles et al. [2]). While the role of host kinases has been extensively studied in the last decades, the role of host phosphatases in the context of viral infections is still poorly understood.
Protein phosphatase 1 (PP1), a member of the phosphoprotein phosphatase (PPP) family, catalyses an important fraction of protein Ser/Thr dephosphorylation events in eukaryotic cells [5]. This phosphatase is involved in the regulation of several cellular processes such as the cell cycle, transcription, protein synthesis, and apoptosis [6,7,8,9]. In mammalian cells, the PP1 catalytic subunit (PP1c) is encoded by three distinct genes that encode three isoforms—PP1α, PP1β/δ, and PP1γ – which are ubiquitously expressed in all tissues [10]. The PP1c isoforms have a nearly identical catalytic core (~ 90%) and mainly differ in their amino (N)- and carboxy (C)- terminal extremities [7]. All PP1c isoforms have poor subtract specificity, hence, no free PP1c pools are expected to exist in cells, to prevent uncontrolled and aberrant dephosphorylation events [11]. However, PP1 counteracts the activity of over 100 kinases, which is explained by the interaction with regulatory subunits, known as Regulatory Interactors of Protein Phosphatase One (RIPPO), that tightly control the substrate selectivity, localization, and activity of PP1c [6,7,8, 11, 12]. Currently, about 200 structurally unrelated vertebrate RIPPOs are known [13], enabling cells to generate a huge diversity of functionally distinct PP1 holoenzymes. Most RIPPOs have short linear motifs (SliMs) that mediate binding to PP1. The most common PP1-binding SLiMs are the so-called RVxF, SILK, MyPhoNE, and ΦΦ motifs [14, 15] that dock to surface grooves on the globular catalytic core of PP1c [12, 14, 16]. In addition, some RIPPOs interact with isoform-specific residues in the N- or C-termini of PP1c, accounting for their fairly selective binding to one PP1 isoform [7, 17, 18]. Most RIPPOs have multiple PP1-binding motifs that, together, create a high-affinity interaction interface with PP1c [14].
In recent years, some studies have reported that different viruses are able to hijack PP1 and subvert its activity to favour infection. Here, we review the current knowledge on the role of PP1 within different viruses’ life cycles, discuss its importance for the antiviral immune response, and suggest PP1 and its related mechanisms as potential host-based targets for the development of new antiviral therapies.
PP1 in viral infections
PP1 promotes Tat-induced transcription in human immunodeficiency virus infection
The human immunodeficiency virus (HIV) is an enveloped retrovirus from the Retroviridae family and its genome is composed of two copies of positive-sense single-stranded RNA. It is classified into two subtypes (HIV-1 and HIV-2), of which HIV-1 is the most prevalent and pathogenic [19]. Among HIV-1 encoded proteins, Tat is a potent transactivator expressed early in infection and has a crucial role in transcriptional activity increment. Tat promotes transcription initiation through interactions with Sp1 elements in the HIV-1 promotor [20], and transcriptional elongation through the recruitment of the host positive transcriptional elongation factor b (P-TEFb) to the transactivation response region (TAR) of the nascent viral mRNA [21]. Moreover, Tat promotes the formation of a super-elongation complex through the recruitment of other elongation factors and co-activators to the HIV-1 promotor [22, 23]. Earlier studies hypothesized a possible involvement of PP1 in Tat-induced transcription [24,25,26,27]. It was observed that the overexpression of the nuclear inhibitor of PP1 (NIPP1) led to a total block of Tat-induced transcription but had a weak impact on HIV-1 basal transcription [25, 26, 28]. The simultaneous co-expression of PP1γ and NIPP1 rescued the Tat-induced transcription [25, 28], whereas the PP1γ dead-mutant co-expression failed this rescue [27, 28]. Subsequent analyses showed an interaction between PP1γ and Tat, as well as their co-localization in the nucleus [27]. Tat possesses a consensus RVxF motif (35QVCF38) and the mutation of this PP1-binding SLiM impaired the PP1:Tat interaction, inhibited the PP1α redistribution into the nucleolus, and blocked Tat-induced transcription [27]. Together, these observations indicate that Tat acts as a viral RIPPO and enhances viral transcription elongation through dephosphorylation by PP1. But how does PP1 mediate the Tat-induced transcription? As described above, Tat recruits the transcription factor P-TEFb, a multimeric complex that includes cyclin-dependent kinase 9 (CDK9):cyclin-T1. This Ser/Thr kinase phosphorylates the C-terminal domain (CTD) of the largest subunit of RNA polymerase II (Pol II) [29, 30]. CDK9 activity is required in the early elongation phase to convert Pol II into a full elongation-competent polymerase (reviewed by Egloff [31]). When fully activated, P-TEFb is only composed of CDK9:Cyclin T1 and phosphorylated at CDK9 Thr186 [32,33,34]. This phosphorylation allows the interaction of CDK9 with 7SK small nuclear RNA (7SK snRNA) and hexamethylene bisacetamide-inducible protein (HEXIM1) that, together with La-related Protein 7 (LAPR7) and 7SK methylphosphate capping enzyme (MePCE), inhibit the P-TEFb complex [31, 35, 36]. P-TEFb is released from inhibition through dephosphorylation of CDK9 at Thr186 by PP1 [35, 37, 38]. During HIV-1 infection, Tat recruits P-TEFb to the TAR of the nascent viral mRNA and releases P-TEFb from its inhibitory complex through competition with HEXIM1 and 7SK snRNA [31, 39,40,41]. It has been suggested that Tat shuttles PP1 to the nucleus, where it dephosphorylates CDK9 Thr186 [27, 42]. The overexpression of NIPP1 or its central PP1-anchoring domain (cdNIPP1) resulted in increased levels of the CDK9 Thr186 phosphorylation [26, 28], supporting the proposed mechanism. Further research also implicated PP1 in CDK9 dephosphorylation at Ser175, promoting the up-regulation of the HIV-1 transcription [43, 44]. Together, these studies demonstrated that Tat acts as a viral RIPPO, inducing PP1 nuclear translocation and enhancing the HIV transcription elongation (Fig. 1A).
Schematic representation of the interplay between PP1 and different viruses. A HIV-1 Tat recruits PP1 for nuclear translocation, thereby promoting the P-TEFb dephosphorylation at the CDK9 Thr186 and enhanced transcription elongation of the viral genome. B EBOV and MARV VP30 are substrates of PP1 and their dephosphorylation shifts the RNA polymerase complex activity towards viral transcription. Since VP30 proteins are devoid of PP1-binding motifs, the PP1:VP30 interaction appears to be mediated by an unknown RIPPO. C In CREB-induced transcription, PP1 is recruited by HDAC1 to promote CREB dephosphorylation and inhibition. HBx binds to the CREB and interacts with the PP1:HDAC1 complex, to prevent CREB dephosphorylation. Thus, HBx promotes CREB-mediated HBV transcription. Also, PP1 dephosphorylates HBV Cp and it is encapsulated together with HBV pgRNA. D HSV-1 pUL21 and VZV pORF38 recruit PP1 to induce dephosphorylation of other viral or host proteins. During HSV-1 infection, the PP1:pUL21 complex dephosphorylates CERT, which contributes to host lipid traffic between the endoplasmic reticulum and the Golgi complex. For all figures, dash lines represent PP1 interactions that are not fully understood. Figure created with BioRender.com
PP1 fosters different activities of the RNA polymerase complex of Ebola and Marburg viruses
Filoviruses (Filoviridae family), including Ebola (EBOV) and Marburg (MARV) viruses, are enveloped non-segmented negative-sense singled stranded RNA viruses, responsible for viral haemorrhagic fevers and a high mortality rate in humans [45,46,47,48]. The EBOV RNA polymerase complex, formed by the large subunit of polymerase L, its cofactor virion protein (VP) 35, and the nucleoprotein (NP), is involved in viral RNA transcription and replication [49]. The transcriptional activator VP30 is also required, being a crucial regulator that switches the polymerase complex activity from replication to transcription [50]. The VP30 activity is regulated by its RNA binding capacity and phosphorylation state [51,52,53]. Dephosphorylated VP30 has a higher affinity for RNA and VP35, promoting transcription [51, 54]. On the other hand, phosphorylated VP30 has increased affinity for NP, thereby shifting the polymerase complex activity towards replication [52, 54]. The VP30 dephosphorylation is regulated by the host phosphatases PP1 and protein phosphatase 2A (PP2A) [55]. PP1 inhibition by cdNIPP1 overexpression led to increased levels of VP30 phosphorylation, suggesting that VP30 is a PP1 substrate [56]. However, when PP1 was knocked down by shRNA, there was neither an increase in VP30 phosphorylation nor a shift of RNA polymerase complex activity to replication [56]. This appeared to be mediated by the splicing factor that interacts with PQBP-1 and PP1 (SIPP1) upregulation, a RIPPO that shuttles PP1 from the nucleus to the cytoplasm [57], thereby promoting VP30 dephosphorylation and viral RNA transcription [56]. While phosphorylated EBOV VP30 shifts RNA polymerase activity from transcription to replication [50], phosphorylated MARV VP30 acts as a transcription repressor [58]. Although it was initially suggested that MARV VP30 was not required for viral transcription [59, 60], later studies proposed a key role of this protein in MARV transcription [58, 61]. The current hypothesis postulates that phosphorylated VP30 sequesters VP35 and NP, preventing MARV transcription [58]. Indeed, PP1 inhibition increased VP30 phosphorylation and its interaction with VP35 and NP, resulting in decreased MARV transcription [58].
It is not yet known how PP1 is recruited to dephosphorylate VP30 from EBOV or MARV since both VP30 lack a consensus PP1-binding SLiM. Recently, it was shown that EBOV NP possesses a conserved PP2A-binding motif that is necessary for PP2A recruitment and VP30 dephosphorylation [62]. An unknown RIPPO (either cellular or viral) may be involved in the recruitment of PP1 by filoviruses to promote VP30 dephosphorylation (Fig. 1B). Further research is needed to clarify the PP1 recruitment mechanism by EBOV and MARV.
PP1 inhibits the hepatitis B virus CREB-induced transcription but promotes its genome packaging
Hepatitis B virus (HBV) is a small enveloped hepatotropic DNA virus from the Hepadnaviridae family and is responsible for acute and chronic infections worldwide [63]. The HBV X protein (HBx) is a multifunctional viral protein that modulates the expression of several cellular and viral genes through interactions with host transcriptional factors and promotes the evasion of different host signaling pathways [64]. Among the transcriptional factors, HBx binds to the cAMP response element binding protein (CREB), activating the transcription of the CREB-targeted genes and HBV DNA [64,65,66]. CREB activation requires its phosphorylation at Ser133, allowing the recruitment of CREB-binding protein (CBP)/p300 to promote transcription [67, 68]. CREB activity is reversed through dephosphorylation by PP1, following their recruitment by histone deacetylase 1 (HDAC1) [69, 70]. Upon HBV infection, HBx enables CREB-induced transcription through inhibition of its dephosphorylation by PP1 [65]. Though it was reported that HBx interacts with PP1:HDAC1, it does not disrupt the complex [65], suggesting that HBx sequesters this complex and avoids the CREB dephosphorylation (Fig. 1C).
Recently, PP1 was implicated in the dephosphorylation of HBV core protein (Cp), the main component of the HBV nucleocapsid [71], where pre-genomic RNA (pgRNA) is packaged and converted into viral DNA by DNA polymerase (DNA Pol) [72, 73]. The Cp CTD is composed of eight conserved Ser and Thr residues which are dynamically phosphorylated during virus infection [74, 75]. Phosphorylated Cp was observed intracellularly and in empty nucleocapsids, whereas unphosphorylated Cp was detected in mature nucleocapsids and in secreted pgRNA-containing virions [71, 76,77,78]. Cp dephosphorylation by PP1 appears to be necessary for pgRNA and DNA Pol encapsulation [71, 77, 79]. Indeed, the knockdown of PP1 by siRNA was associated with increased Cp phosphorylation levels and decreased pgRNA packaging [71]. Hu et al. also reported that PP1α and PP1β are co-packaged in pgRNA-containing nucleocapsids (Fig. 1C) [71]. A previous study reported the co-packaging of cyclin-dependent kinase 2 (CDK2), which plays a critical role in releasing HBV DNA from the nucleocapsid through the phosphorylation of the Cp N- and C-termini [80]. Noteworthy, PP1 is inhibited by CDK2 phosphorylation [81], suggesting that a posteriori PP1 inhibition is required for pgRNA release.
PP1 is recruited by Herpes Simplex virus 1 pUL21 to promote dephosphorylation of viral and host proteins
Herpesviruses are large enveloped double-strand DNA viruses that can remain in a dormant, latent state in host cells [82, 83]. Herpes Simplex virus (HSV) is an alpha-herpesvirus that infects epithelial cells and establishes latency in neurons [82, 83]. The HSV-1 tegument protein pUL21 was recently described as a PP1 interactor that regulates the phosphorylation status of viral and cellular proteins [84]. In early infection, pUL21 promotes the transport of the HSV capsid to the nucleus [85, 86], whereas in late infection pUL21 is required for the nuclear egress of the HSV capsids, and to promote the glycosylation of the viral glycoprotein gE [85, 87, 88]. Co-immunoprecipitation assays showed an interaction between pUL21, all three PP1 isoforms, and the ceramide transport protein (CERT) [84]. Dephosphorylated CERT participates in the ceramide exchange and lipid traffic between the endoplasmic reticulum and the Golgi complex [89, 90]. pUL21 acts as a bridge between PP1 and CERT, promoting CERT dephosphorylation. Consequently, dephosphorylated CERT modulates the cellular lipid composition of the endoplasmic reticulum and Golgi complex and/or the post-Golgi protein traffic [84, 91]. Nevertheless, the HSV-1 replication was not significantly altered when pUL21-mediated CERT dephosphorylation was abolished in keratinocytes and epithelial cells [91]. The Varicella-zoster virus (VZV), another alpha-herpesvirus, possesses an HSV-1 pUL21 homologue—the pORF38 protein – that also binds to PP1 but does not bind to CERT, suggesting that CERT recruitment is HSV-1-specific [84]. A full understanding of the role of pUL21 in HSV-1 infection and the CERT pathway modulation requires further investigation (Fig. 1D). Ma and colleagues also demonstrate that PP1 is an interactor of the porcine pseudorabies virus (PRV) pUL21, another orthologue of the HSV pUL21 [92]. Although the functional interaction between PRV pUL21 and PP1 was not explored, these results suggest that the binding of alpha-herpesviruses to PP1 is phylogenetically conserved.
Interestingly, pUL21 is devoid of a canonical PP1-binding SLiM but has a motif in the linker region, named Twenty-one Recruitment Of Protein Phosphatase One (TROPPO), that is conserved among the alpha herpesviruses (including VZV pORF38 and PRV pUL21) and has the consensus sequence φ-S-x-F-V-Q-[VI]-[KR]-x-I, where φ is a hydrophobic residue and x any amino acid (pUL21: 239VSEFVQVKHI248) [84]. Mutations in pUL21 Phe242 and Val243 impaired PP1 binding and significantly decreased viral replication [84]. Yet, HSV pUL21 mutants showed a remarkable capacity for adaption in that they recovered their replication potential through mutations in the pUS3 gene, a viral kinase involved in HSV nuclear egress [93]. Benedyk et al. reported that these pUS3 mutations reduced stability and kinase activity, suggesting that pUL21 may counteract pUS3 kinase through PP1 recruitment for the same substrates (e.g. pUL31) in the later phases of infection (Fig. 1D) [84].
PP1 plays important roles in the host cell response to viral infections
Viral proteins recruit PP1 to maintain translation through dephosphorylation of eIF2α
Upon viral infection, the production of double-strand RNA (dsRNA) induces activation of the dsRNA-dependent protein kinase R (PKR), which phosphorylates the eukaryotic initiation factor 2 subunit α (eIF2α) at Ser51 and blocks the translation initiation [94, 95]. Moreover, the accumulation of viral proteins in the endoplasmic reticulum (ER) induces the activation of the protein kinase R-like ER kinase (PERK), which also phosphorylates eIF2α at Ser51 [96]. Yet, this translational block is reversed through eIF2α dephosphorylation by PP1. Two RIPPOs, Growth arrest and DNA damage-inducible protein (GADD34, also known as PPP1R15A) and Protein Phosphatase 1 regulatory subunit 15B (PPP1R15B, also known as CREP) recruit PP1 to promote eIF2α dephosphorylation [97, 98]. However, viruses evolved mechanisms to prevent eIF2α phosphorylation by PKR and PERK or to promote eIF2α dephosphorylation through the recruitment of PP1 (Fig. 2) or PP2A [99].
PP1 is recruited by some viruses to rescue host translation activity. dsRNA sensing and viral protein accumulation in ER activates the PKR and PERK, respectively, culminating in eIF2α phosphorylation at Ser51 and host cell translation shut-off. This process is reversed by the PP1:GADD34 holoenzyme. HSV-1 γ34.5, ASFV DP71L, and TGEV protein 7 are viral orthologues of GADD34 and interact with PP1 to promote eIF2α dephosphorylation and protein synthesis rescue. Some viruses – DENV, HCV, NDV, HPV, PRV, and IBV –modulate the activity of host-cell PP1:GADD34. Figure created with BioRender.com
In HSV-1-infected cells, the neurovirulence factor γ34.5 is mainly responsible for keeping eIF2α dephosphorylated [100]. The C-terminus of γ34.5 has a primary structure similar to the C-terminal domain of GADD34 [101]. This γ34.5 region contains a validated RVxF motif (HSV-1: 192RVRF195) that mediates the recruitment of PP1α (Fig. 2) [102, 103]. Mutations of this motif abolished eIF2α dephosphorylation, impaired HSV-1 replication, and increased the sensitivity to interferon response [102, 104, 105]. The γ34.5 C-terminus also possesses an Ala-Arg-rich motif that directly binds to eIF2α [106,107,108,109], acting as a bridge between PP1 and the phosphorylated eIF2α. Furthermore, Meng et al. reported that HSV-1 γ34.5 enables the shuttling of ribosome biogenesis protein NOP53 from the nucleus to the cytoplasm, which was suggested to facilitate PP1α recruitment by γ34.5 [110]. Despite NOP53 seeming to be necessary for efficient HSV-1 viral replication [110], its role in the PP1 recruitment by the γ34.5 is still unknown. The African swine fever virus (ASFV) DP71L protein is another GADD34 viral orthologue [101]. The ASFV encodes a DP71L long form (184 aa) and a short form (70–72 aa), both possessing a RVxF motif: 124KVYF127 and 14KHVRF18, respectively [101, 111]. ASFV infection also induces a translation shut-off, being this response counter-acted by the PP1 recruitment mediated by the DP71L and subsequent dephosphorylation of the eIF2α (Fig. 2) [112,113,114]. In addition to the RVxF motif, DP71L also has a LSAVL motif (long form: position 169–173; short form: position 57–61) that is required to bridge the interaction between PP1 and eIF2α [111]. Noteworthy, the mutant ASFV ∆DP71L still possesses the capacity to induce eIF2α dephosphorylation, suggesting that other ASFV proteins may regulate the eFI2α phosphorylation [114].
The transmissible gastroenteritis virus (TGEV) protein 7 interacts with PP1 through a RVxF motif (58RVIF61) that is conserved in other α-coronaviruses’ (α-CoVs) protein 7, including the canine coronavirus (CcoV), porcine respiratory coronavirus (PRCV), and feline infectious peritonitis (FIPV) [115]. Co-immunoprecipitation assays showed that TGEV protein 7 interacts with both PP1 and eIF2α [115]. Compared to cells infected with wildtype TGEV, cells infected with TGEV lacking protein 7 (TGEV ∆7) had higher levels of phosphorylated eIF2α and showed premature protein synthesis inhibition, widespread apoptosis, and exacerbated immune response [115, 116]. TGEV ∆7-infected pigs presented increased tissue damage and proinflammatory response [115, 116]. Although protein 7 is not essential for TGEV replication, it appears to attenuate the virulence of the infection in both in vitro and in vivo models. Similarly, in infected mice with severe acute respiratory syndrome coronavirus (SARS-CoV), a beta coronavirus (β-CoV), the loss of the PP1 inhibitor Kepi was also associated with increased virus pathogenicity [117]. Hence, PP1 activity appears to be required to avoid severe pathological damage during TGEV and SARS-CoV infections.
Other viruses such as dengue virus (DENV) [118], hepatitis C virus (HCV) [119], the New Castle disease virus (NDV) [120], human papillomavirus (HPV) [121], PRV [122, 123], and infectious bronchitis virus (IBV) [124] appears to maintain protein translation and low eIF2α phosphorylation levels through stimulatory interactions with the host-cell PP1:GADD34 holoenzyme (Fig. 2). In addition, some of those viruses increase GADD34 expression during late infection, which contributes to increased dephosphorylation of eIF2α through the PP1:GADD34 [118, 120, 121, 123, 124]. Nevertheless, the GADD34 up-regulation should be carefully analysed since it is a downstream effect of the eIF2α phosphorylation (through the ATF4/Chop pathway) even in other stress-related conditions [125]. Therefore, the GADD34 increment may not be a specific phenomenon caused by a virus. Further research is required for a better understanding of the PP1:GADD34 holoenzyme role in those viral infections.
PP1 in Rig-I-like receptors’ activation and its modulation by measles virus and α-coronaviruses
RIG-I-like receptors (RLRs) are a family of cytosolic RNA sensors for the host defense against viral infections and include three members: retinoic-acid inducible gene I (RIG-I), melanoma differentiation association gene 5 (MDA5), and Laboratory of genetics and physiology 2 (LGP2) [126]. Although RIG-I and MDA5 recognize different viral RNA molecules, they are structurally similar and possess two caspase activation and recruitment domains (CARDs) at the N-terminus, which mediate the downstream signal transduction [126, 127]. Upon recognition of viral RNA, RIG-I or MDA5 oligomerizes, allowing the subsequent interaction with the mitochondrial antiviral-signalling protein (MAVS) at mitochondria and peroxisomes [128, 129]. MAVS is responsible for the induction of the subsequent signalling leading to the production of interferons (IFNs), interferon-stimulated genes (ISGs), and other cytokines [130]. To induce an INF response, MAVS recruits the tumour necrosis factor receptor-associated factor 3 (TRAF3) and TRAF family member-associated NF-κB activator (TANK). Subsequently, the TRAF3/TANK complex activates the IκB Kinase ε (IKKε) and TANK-binding kinase 1 (TBK1), two kinases responsible for the phosphorylation of the interferon regulatory factors (IRF) 3 and 7 [131]. Then, IRF3 and IRF7 are translocated to the nucleus and induce the expression of type I INF [130].
RLR activation is a tightly regulated process and different studies have demonstrated that it highly depends on protein phosphorylation [132]. In resting cells, RIG-I and MDA5 exist in an inactive conformational state where Thr and Ser residues in CARDs are phosphorylated (RIG-I: Ser8 and Thr170; MDA5: Ser88) [133,134,135]. Upon infection, the RIG-I/MDA5 CARDs are rapidly dephosphorylated by PP1α/γ (Fig. 3), followed by K63-linked polyubiquitination that triggers MAVS-dependent downstream signalling [135, 136]. RIG-I and MDA5 possess two PP1-binding SLiMs: a F-x-x-R/K-x-R/K motif at the CARD (RIG-I: 94FKKIEK99; MDA5: 18FRARVK23), and a RVxF motif at the helicase region (RIG-I: 292KVVFF296; MDA5: 397KISF400) [135]. While both MDA5 PP1-binding SLiMs are required for the interaction with PP1, only the RIG-I F-x-x-R/K-x-R/K motif appears to be necessary for this interaction [135]. However, the PP1 recruitment mechanism to dephosphorylate RLRs after viral dsRNA recognition is not fully understood. Recently, Acharya et al. reported that the Protein phosphatase 1 regulatory subunit 12C (PPP1R12C or R12C) mediates the RIG-I/MDA5 dephosphorylation by PP1 [137]. R12C participates in the actomyosin cytoskeleton dynamic regulation as part of myosin phosphatase complex [138]. Upon viral infection, the disturbance of the actin cytoskeleton induces the subcellular redistribution of R12C and the formation of the R12C:PP1:RLR complex, which dephosphorylates RIG-I/MDA5 and promotes the downstream signalling (Fig. 3) [137]. Thus, R12C acts as an upstream regulator of the RLR pathway, recruiting PP1 to RIG-I/MDA5 and promoting their activation. Still, further investigation is required to unravel other RIPPOs involved in the RIG-I/MDA5 dephosphorylation mediated by PP1.
PP1 is required for the RLRs activation, and its activity is modulated by MeV and α-CoVs. Upon dsRNA sensing, PP1 is recruited (by unknown RIPPOs) to dephosphorylate RIG-I/MDA5 and promote their activation. Recently, it was shown that as a consequence of the cytoskeleton disturbance, R12C recruits PP1 to dephosphorylate RIG-I and MDA5. In the early infection of dendritic cells, MeV interacts with the DC-SING receptor leading to the activation of Raf-1 kinase. Raf1 phosphorylate inhibitor 1 (I-1), a potent PP1 inhibitor, avoids RIG-I/MDA5 activation. In late MeV infection, protein V prevents MDA5 dephosphorylation by acting as a PP1-substrate. In α-CoV infections, the nsp7 binds to MDA5 N-terminus and blocks its dephosphorylation by PP1. Figure created with BioRender.com
The measles virus (MeV), a paramyxovirus, evolved several mechanisms to escape the IFN response [139,140,141], two of which are related to PP1 targeting to suppress the RIG-I/MDA5 dephosphorylation [142]. In the first mechanism, the MeV interacts with the C-type lectin DC-SIGN receptor in the dendritic cell (DC) surface and induces the phosphorylation and activation of protein kinase Raf1. Raf1 then phosphorylates inhibitor 1 (I-1), a specific inhibitor of the PP1:GADD34 holoenzyme [143]. The phosphorylated I-1 blocks RIG-I/MDA5 dephosphorylation and subsequent activation (Fig. 3) [143]. The second mechanism involves the MeV protein V, an accessory protein that is crucial to the suppression of the host IFN response [144, 145]. At the late infection stage, protein V antagonizes the MDA5 dephosphorylation by PP1α/γ and serves as a putative PP1 substrate, maintaining PP1 away from MDA5 (Fig. 3) [146]. The MeV protein V has a consensus RVxF motif in the C-terminus region (288RIWY291). Mutation or deletion of this motif reduced the PP1 binding to MeV protein V and increased MDA5 dephosphorylation [146]. It was also reported that the protein V of the Nipah virus (NiV), another paramyxovirus, interacts with PP1 [146]. However, contrary to MeV protein V, NiV protein V is devoid of PP1-binding SliMs [146]. This observation may suggest that PP1 recruitment by paramyxoviruses is required for their survival. Together, these studies showed that MeV avoids RLR activation by preventing both RIG-I and MDA5 dephosphorylation in a Raf1-dependent manner during early infection, and the MDA5 dephosphorylation in a protein V-dependent manner in the late infection [142, 143, 146].
Several studies have reported that various coronaviruses encode immunomodulatory proteins capable of suppressing RLR signalling [92, 147,148,149,150,151]. The α-CoV porcine epidemic diarrhoea virus (PEDV) non-structural protein 7 (nsp7) inhibits the RLR pathway by preventing the MDA5 dephosphorylation [152]. The nsp7 competes directly with PP1α/γ for binding to MDA5 and thereby prevents MDA5 Ser828 dephosphorylation and subsequent MAVS signalling [152, 153]. This mechanism also applies to other α-CoVs that encode orthologues of the PEDV nsp7 such as TGEV, swine acute diarrhoea syndrome coronavirus (SADS-CoV), and feline coronavirus (FCoV) [152].
PP1 is a negative regulator of the Toll-like receptor (TLR) pathway and is recruited by HSV-1 and enteroviruses to modulate TLR signaling
Toll-like receptors (TLR) are transmembrane glycoproteins located at the plasma membrane or endosomal membranes and sense a vast number of pathogen- and damage-associated molecular patterns [154]. Upon activation, TLRs trigger a downstream signalling cascade that culminates in the activation of transcription factors, interferon regulatory factors (IRFs), and nuclear factor kappa-light chain enhancer of activated B cells (NF-κB) [155]. Among the 10 known human TLRs, TLR3, TLR7, TLR8, and TLR9 are involved in the recognition of viral nucleic acids [156]. TLR7, TLR8, and TLR9 signal through the myeloid differentiation primary response protein 88 (MyD88) adaptor to activate the NF-κB or mitogen-activated protein kinase (MAPK) pathways, resulting in the expression of proinflammatory cytokines [156]. TLR7-9 also promotes the IFN response through the activation of IRF-5 and IRF-7. TLR3 signals through the TIR domain-containing adaptor protein inducing IFN-β (TRF) adaptor, leading to the activation of both IRF-3 and NF-κB signaling [157]. Phosphorylation by various kinases is crucial for TLR activation [158]. PP1 acts as a negative regulator of the TLR pathway by dephosphorylation of different mediators (Fig. 4).
PP1 is a negative regulator of the TLR pathway and is recruited by viruses to impair TLR signaling. PP1:GADD34 holoenzyme counteracts the activation of TAK1 and IKK, two crucial kinases in the TLR cascade. In resting cells, IKK is maintained in an inactive state by the PP1:GADD34:CUEDC2 complex. Upon viral infection, PP1 avoids the excessive activation of IRF3 and IRF7 and limits IFN-α/β expression. HSV-1 γ34.5 and enteroviruses (EV71, PV, CVA, and CVB) protein 2C recruit PP1 to dephosphorylate IKKβ and inhibit NF-κB activation. Figure created with BioRender.com
Three main Ser/Thr kinase families are conserved elements of TLR signaling: interleukin-1 receptor kinases (IRAKs), the tumor growth factor β-activated kinase (TAK1), and inhibitors of NF-κβ (IκB) kinase (IKK) complex [159]. TAK1 is activated by ubiquitination and phosphorylation at Thr184, Thr187, and Ser412 [160, 161]. PP1:GADD34 holoenzyme dephosphorylates TAK1 at Ser412, thereby preventing downstream NF-κB and MAPK activation [162]. Although all three PP1 isoforms can interact directly with TAK1, Ser412 dephosphorylation is dependent on GADD34, which possesses a TAK1-binding domain [162]. The IKK complex is formed by two Ser/Thr kinases (IKKα and IKKβ) and a regulatory scaffolding protein NF-kβ essential modifier (NEMO, also known as IKKγ) [163, 164]. In resting cells, IKK is sequestered in the cytoplasm by the PP1:GADD34:CUE domain–containing protein 2 (CUEDC2) complex that maintains IKK in a non-phosphorylated, inactive state [165]. In TNF-stimulated mouse embryonic fibroblasts, IKK is released from its inhibitory complex by tumor necrosis factor receptor-associated factor 2 (TRAF2) and is phosphorylated by the TNF receptor. After activation, the IKK is released from the TNF-receptor signaling complex and is available to interact with CUEDC2. GADD34 mediates the interaction between PP1 and CUEDC2 and promotes IKK dephosphorylation and inactivation [165]. A similar phenomenon was observed in LPS-stimulated cells (a TLR signaling inductor), being the IKK released from its inhibitory complex by TRAF6 [165]. Together, these results are indicative that PP1 negatively regulates specific steps of TLR signaling.
Upon viral recognition by TLRs, IRF3, and IRF7 are activated through phosphorylation of key Ser/Thr residues, and subsequently induce IFN production [166,167,168]. In NDV-infected cells, inhibition of PP1 enhanced IRF7 activation and IFN-α production, and impaired NDV replication [169]. PP1 binds to IRF7, which possesses two RVxF motifs (11RVLF14 and 408RVFF411), and reduces its activation through dephosphorylation [169]. Like IRF7, IRF3 possesses a RVxF motif (213RQVF216) that is necessary for the PP1 binding [170]. In turn, PP1 prevents full IRF3 activation, decreasing the TLR- and RLR-mediated IFN-β expression [170].The human enterovirus EV71 – the causative agent of hand, foot, and mouth disease – inhibits IKKβ and suppresses the NF-κβ signaling pathway [171]. The EV71 protein 2C forms a complex with PP1 and IKKβ, promoting IKKβ dephosphorylation and inactivation (Fig. 4). The EV71 protein 2C possesses three RVxF motifs (motif 1: 16KGLEG20; motif 2: 27KFIDW31; motif 3: 43KVEF46) that appear to be required for PP1 recruitment [171]. Three other enteroviruses – poliovirus (PV), coxsackievirus A (CVA), and coxsackievirus B (CVB) – also inhibit IKKβ dephosphorylation through PP1 recruitment by their protein 2C ortholog [171]. In HSV-1-infected DCs, γ34.5 binds to IKKα/β via the N-terminus and recruits PP1 via the C-terminus, preventing the IKKβ phosphorylation and NF-κβ activation (Fig. 4) [172]. This negatively impacts DC maturation, where the TLR-4/NF-κβ pathway is activated to promote the expression of inflammatory and co-stimulatory molecules that are essential for maturation [173]. Collectively, these studies highlight how viruses can subvert the regulatory role of PP1 in TLR signaling to promote their escape from the host immune response.
Although most studies highlighted the role of PP1 as a negative regulator of the TLR pathway, Opaluch et al. proposed that PP1 may positively regulate TLR signaling (Fig. 4) [174]. Upon TLR4/5/7 stimulation, PP1γ dephosphorylates TRAF6 and promotes downstream signaling events including NF-κB activation [174]. Yet, the full mechanism behind PP1:TRAF6 interaction and regulation remains poorly understood.
PP1-mediated dephosphorylation of myxovirus resistance 2 (MX2) is required for the innate response against HIV-1
The innate immune response mobilizes an antiviral state that stimulates the production of IFNs, which induces the expression of IFN-stimulated genes that participate in antiviral responses against viral pathogens [175]. One of the IFN-stimulated genes is MX2 (also named MXB), a GTPase related to MX1 that presents a potent inhibitory activity against HIV-1 [176,177,178,179], HSV-1, and HSV-2 replication [180, 181]. In HIV-1 infection, MX2 blocks the nuclear entry of the viral genome by binding to the HIV-1 capsid, which precludes viral DNA accumulation and integration into host chromosomes [182, 183]. Betancor et al. showed that the N-terminal domain (NTD) of MX2 interacts with PP1β:myosin phosphatase target subunit 1 (MYPT1) holoenzyme [184]. PP1 inhibition or depletion increased the MX2 NTD phosphorylation (Ser 14, 17, and 18), and reduced the ability of MX2 to bind to the HIV-1 capsid [184]. This suggests that PP1β regulates the antiviral activity of MX2. The NTD of MX2 has a F-x-x-R/K-x-R/K motif (8WPYRRR13) that also contributes to the binding of the PP1β:MYPT1 [184]. Together, these findings suggest that MX2 is regulated by phosphorylation, and its IFN-dependent activation may also induce MX2 dephosphorylation [184].
PP1 as potential target for antiviral therapies
Classical antiviral therapies are focused on the inhibition of viral processes by targeting viral proteins [185,186,187]. However, this strategy often fails due to viral adaptation, which leads to the emergence of drug-resistant mutants [188]. During viral infections, several interactions between viral and host proteins are established and maintained to facilitate virus surveillance and propagation. Therefore, targeting host-based mechanisms became an efficient alternative strategy to fight viruses, allowing the design of molecules that interfere with host proteins hijacked by them. Such molecules are expected to have a broad spectrum activity and to decrease viral drug resistance [189, 190].
For a long time, phosphatases were considered unattractive targets for the development of drug therapies [191]. The design of selective modulators for PP1’s active site is highly challenging due to its great conservation within the PPP family [192, 193]. However, PP1 holoenzymes can be selectively targeted by interference with PP1:RIPPO interactions [16, 194]. One effective approach is the design and synthesis of PP1-disrupting peptides that allow the disruption of PP1 holoenzymes’ interactions and the modulation of certain processes [195,196,197,198]. In this sense, the PP1-drug targeting landscape has been explored to fight viral infections. Through in silico modulation, a set of molecules was designed based on the HIV-1 Tat RVxF motif, leading to the synthesis of 1,2,3,4-tetrahydracridine (1H4) that decreases the interaction between PP1 and Tat, and significantly inhibits HIV-1 transcription and replication [199]. Based on the 1H4 structure, a library of compounds was designed to improve HIV-1 inhibition, of which compound 1E7-03 showed a strong HIV-1 inhibition capacity with a lower IC50 (five-fold compared to 1H4) [200]. 1E7-03 was also the first PP1-small target molecule that significatively inhibited HIV-1 transcription in mice [201]. While 1E7-03 had strong in vivo inhibitory capacity, it had lower metabolic stability being converted into two products (DP1 and DP3) that bind PP1 but present low cell penetration capacity [201]. Further structural improvement of 1E7-03 led to the synthesis of the compound HU-1a, which has an overall superior HIV-1 inhibitory capacity as well as improved metabolic stability [202]. The 1E7-03 treatment diminished the replication of the Rift Valley fever virus (RVFV) and the Venezuelan equine encephalitis virus (VEEV) [203, 204], suggesting that PP1 also plays a role in these viral infections. Indeed, Carey et al. showed that PP1 interacts with the VEEV capsid and regulates at least two phases of viral replication: the early post-entry and late viral assembly [204]. The same authors also reported that other alphaviruses – including Eastern Equine Encephalitis Virus (EEEV), West Equine Encephalitis Virus (WEEV), Sindbis virus (SINV), and Chikungunya virus (CHIKV) – also presented impaired viral replication after 1E7-03 treatment [204]. After EBOV infection, 1E7-03 inhibited VP30 dephosphorylation and prevented the EBOV transcription [56]. Surprisingly, 1E7-03 also strongly inhibited EBOV replication in a dose-dependent manner, which may result from the transcription/replication imbalance promoted by high levels of phosphorylated VP30 [56]. Two 1E7-03 analogues—1E7-07 and C31 – also prevented VP30 dephosphorylation and viral transcription [205, 206]. In MARV-infected cells, 1E7-03 increased VP30 phosphorylation and decreased MARV transcription and replication [58].
In latent HIV-infected cells, one major concern is that integrated proviruses can become activated and produce viral proteins when antiretroviral therapy is interrupted or drug resistance emerges [207,208,209]. Eradication of latent HIV-1 is challenging since all clinically approved anti-HIV drugs are ineffective unless the viral transcription is activated. Tyagi et al. discovered a sulfonamide-containing compound, named small-molecule activator of PP1 (SMAPP1) that induced HIV-1 transcription in latent HIV-1 infected cells [210]. SMAPP1 is a suitable compound for the kick-and-kill approach, where the HIV-1 is reactivated and then fought through an antiretroviral combination [210].
Concluding remarks
During viral infections, there is an extensive network of interactions occurring between viral and host proteins, not only to promote viral replication but also to increase the host antiviral response. As a multifaceted player that participates in several crucial processes for cell survival and maintenance, it does not come as a surprise that PP1 stands out as one of these modulated host cell factors.
In this review, we discussed the interplay between viral proteins and host-cell PP1 in the context of different viral infections. We highlight that viruses interact with PP1 via two mechanisms: (i) some viruses encode a viral RIPPO (HIV Tat, HBV HBx, HSV pUL21) that interacts with PP1 and promotes the dephosphorylation of host and/or viral proteins; or (ii) viruses encode a protein that is a PP1-substrate (EBOV/MARV VP30, HBV Cp, MeV protein V). Both mechanisms involve the recruitment of PP1 to promote viral propagation. Although this topic has been explored for about two decades, only a few studies present detailed mechanistic data on how PP1 is recruited and how its activity is crucial for virus infection. So far, the role of PP1 in HIV-1 infection is the best characterized. Importantly, some viruses such as filoviruses, herpesviruses, α-CoVs, and enteroviruses appear to retain PP1-binding motifs, suggesting that PP1-recruitment may be crucial for those viruses’ adaption across their evolution.
PP1 is also involved in the regulation of different host antiviral responses and, therefore, some viruses modulate PP1 activity to evade host defence mechanisms. In fact, HSV-1 and ASFV encode viral RIPPOs that are orthologs of host-cell GADD34 and bind to PP1 to induce the eIF2α dephosphorylation. Also, TGEV protein 7 mimics the GADD34 role to direct PP1 to reestablish the translation activity. Regarding RLRs and TLRs pathways, despite most kinases involved in Ser/Thr phosphorylation of these pathways have been widely studied, the phosphatases that counteract their activity remain poorly investigated. One aspect that needs to be clarified is how PP1 is recruited to dephosphorylate the different receptors and whether there are other RIPPOs (still unknown) that promote and direct PP1 activity to these substrates upon receptor activation. Intriguingly, PP1’s role in the cGAS-STING pathway, which is activated by cytosolic DNA (self and invaded viral or microbial) needs to be further explored in the context of viral infection. One study reported that PP1 regulates the cGAS dephosphorylation during mitosis [211], suggesting that PP1 may play a regulatory role in this pathway.
As stated in the previous section, the design and synthesis of compounds that disturb the interaction between PP1 and viral RIPPOs may be an effective strategy to fight viral infections. PP1-based antiviral therapies appear to have a broad spectrum of viral inhibitory activity, as shown with the 1E7-03 compound. Further and more detailed studies are necessary to complement the knowledge on how different viruses modulate and take advantage of PP1 activity and how this can be exploited to develop novel antiviral therapies.
References
Roychoudhury S, Das A, Sengupta P, Dutta S, Roychoudhury S, Choudhury AP, et al. Viral pandemics of the last four decades: pathophysiology. Health Impacts Perspect. 2020;15:9411.
García-Cárceles J, Caballero E, Gil C, Martínez A. Kinase inhibitors as underexplored antiviral agents. J Med Chem. 2022;65(2):935–54.
Keating JA, Striker R. Phosphorylation events during viral infections provide potential therapeutic targets. Rev Med Virol. 2012;22(3):166–81.
Keck F, Ataey P, Amaya M, Bailey C, Narayanan A. Phosphorylation of single stranded RNA Virus proteins and potential for novel therapeutic strategies. Viruses. 2015;7(10):5257–73.
Smith RJ, Cordeiro MH, Davey NE, Vallardi G, Ciliberto A, Gross F, et al. PP1 and PP2A use opposite phospho-dependencies to control distinct processes at the kinetochore. Cell Rep. 2019;28(8):2206-2219.e8.
Peti W, Nairn AC, Page R. Structural basis for protein phosphatase 1 regulation and specificity. FEBS J. 2013;280(2):596–611.
Korrodi-Gregório L, Esteves SLC, Fardilha M. Protein phosphatase 1 catalytic isoforms: specificity toward interacting proteins. Transl Res. 2014;164(5):366–91.
Li X, Wilmanns M, Thornton J, Köhn M. Elucidating human phosphatase-substrate networks. Sci Signal. 2013;6(275):1–15.
Ceulemans H, Bollen M. Functional diversity of protein phosphatase-1, a cellular economizer and reset button. Physiol Rev. 2004;84(1):1–39.
Yates B, Braschi B, Gray KA, Seal RL, Tweedie S, Bruford EA. Genenames.org: the HGNC and VGNC resources in 2017. Nucleic Acids Res. 2017;45(D1):619–25.
Verbinnen I, Ferreira M, Bollen M. Biogenesis and activity regulation of protein phosphatase 1. Biochem Soc Trans. 2017;45(1):89–99.
Wu D, De Wever V, Derua R, Winkler C, Beullens M, Van Eynde A, et al. A substrate-trapping strategy for protein phosphatase PP1 holoenzymes using hypoactive subunit fusions. J Biol Chem. 2018;293(39):15152–62.
Alanis-Lobato G, Andrade-Navarro MA, Schaefer MH. HIPPIE v2.0: enhancing meaningfulness and reliability of protein-protein interaction networks. Nucleic Acids Res. 2017;45(D1):D408-14.
Heroes E, Lesage B, Görnemann J, Beullens M, Van Meervelt L, Bollen M. The PP1 binding code: a molecular-lego strategy that governs specificity. FEBS J. 2013;280(2):584–95.
Peti W, Page R. Strategies to make protein serine/threonine (PP1, calcineurin) and tyrosine phosphatases (PTP1B) druggable: Achieving specificity by targeting substrate and regulatory protein interaction sites. Bioorg Med Chem. 2015;23(12):2781–5.
Chatterjee J, Beullens M, Sukackaite R, Qian J, Lesage B, Hart DJ, et al. Development of a peptide that selectively activates protein phosphatase-1 in living cells. Angew Chemie Int Ed. 2012;51(40):10054–9.
Bertran MT, Mouilleron S, Zhou Y, Bajaj R, Uliana F, Kumar GS, et al. ASPP proteins discriminate between PP1 catalytic subunits through their SH3 domain and the PP1 C-tail. Nat Commun. 2019;10(1):771.
Skene-Arnold TD, Luu HA, Uhrig RG, De Wever V, Nimick M, Maynes J, et al. Molecular mechanisms underlying the interaction of protein phosphatase-1c with ASPP proteins. Biochem J. 2013;449(3):649–59.
Deeks SG, Overbaugh J, Phillips A, Buchbinder S. HIV infection. Nat Rev Dis Prim. 2015;1(1):15035.
Das AT, Harwig A, Berkhout B. The HIV-1 Tat protein has a versatile role in activating viral transcription. J Virol. 2011;85(18):9506–16.
Rice AP. The HIV-1 Tat protein: mechanism of action and target for HIV-1 cure strategies. Curr Pharm Des. 2017;23(28):4098–102.
Sobhian B, Laguette N, Yatim A, Nakamura M, Levy Y, Kiernan R, et al. HIV-1 Tat assembles a multifunctional transcription elongation complex and stably associates with the 7SK snRNP. Mol Cell. 2010;38(3):439–51.
He N, Liu M, Hsu J, Xue Y, Chou S, Burlingame A, et al. HIV-1 Tat and host AFF4 recruit two transcription elongation factors into a bifunctional complex for coordinated activation of HIV-1 transcription. Mol Cell. 2010;38(3):428–38.
Bharucha DC, Zhou M, Nekhai S, Brady JN, Shukla RR, Kumar A. A protein phosphatase from human T cells augments Tat transactivation of the human immunodeficiency virus type 1 long-terminal repeat. Virology. 2002;296(1):6–16.
Ammosova T, Jerebtsova M, Beullens M, Voloshin Y, Ray PE, Kumar A, et al. Nuclear protein phosphatase-1 regulates HIV-1 transcription. J Biol Chem. 2003;278(34):32189–94.
Ammosova T, Washington K, Debebe Z, Brady J, Nekhai S. Dephosphorylation of CDK9 by protein phosphatase 2A and protein phosphatase-1 in Tat-activated HIV-1 transcription. Retrovirology. 2005;2(1):47.
Ammosova T, Jerebtsova M, Beullens M, Lesage B, Jackson A, Kashanchi F, et al. Nuclear targeting of protein phosphatase-1 by HIV-1 Tat protein. J Biol Chem. 2005;280(43):36364–71.
Ammosova T, Yedavalli VRK, Niu X, Jerebtsova M, Van Eynde A, Beullens M, et al. Expression of a protein phosphatase 1 inhibitor, cdNIPP1, increases CDK9 threonine 186 phosphorylation and inhibits HIV-1 transcription. J Biol Chem. 2011;286(5):3798–804.
Hsin J-P, Manley JL. The RNA polymerase II CTD coordinates transcription and RNA processing. Genes Dev. 2012;26(19):2119–37.
Kim W, LeBlanc B, Matthews WL, Zhang Z-Y, Zhang Y. Advancements in chemical biology targeting the kinases and phosphatases of RNA polymerase II-mediated transcription. Curr Opin Chem Biol. 2021;63:68–77.
Egloff S. CDK9 keeps RNA polymerase II on track. Cell Mol Life Sci. 2021;78(14):5543–67.
Larochelle S, Amat R, Glover-Cutter K, Sansó M, Zhang C, Allen JJ, et al. Cyclin-dependent kinase control of the initiation-to-elongation switch of RNA polymerase II. Nat Struct Mol Biol. 2012;19(11):1108–15.
Kwiatkowski N, Zhang T, Rahl PB, Abraham BJ, Reddy J, Ficarro SB, et al. Targeting transcription regulation in cancer with a covalent CDK7 inhibitor. Nature. 2014;511(7511):616–20.
Baumli S, Lolli G, Lowe ED, Troiani S, Rusconi L, Bullock AN, et al. The structure of P-TEFb (CDK9/cyclin T1), its complex with flavopiridol and regulation by phosphorylation. EMBO J. 2008;27(13):1907–18.
Chen R, Yang Z, Zhou Q. Phosphorylated Positive Transcription Elongation Factor b (P-TEFb) is tagged for inhibition through association with 7SK snRNA. J Biol Chem. 2004;279(6):4153–60.
Li Q, Price JP, Byers SA, Cheng D, Peng J, Price DH. Analysis of the large inactive P-TEFb complex indicates that it contains One 7SK molecule, a dimer of HEXIM1 or HEXIM2, and two P-TEFb molecules containing Cdk9 phosphorylated at threonine 186. J Biol Chem. 2005;280(31):28819–26.
Chen R, Liu M, Li H, Xue Y, Ramey WN, He N, et al. PP2B and PP1α cooperatively disrupt 7SK snRNP to release P-TEFb for transcription in response to Ca 2+ signaling. Genes Dev. 2008;22(10):1356–68.
Wang Y, Dow EC, Liang Y-Y, Ramakrishnan R, Liu H, Sung T-L, et al. Phosphatase PPM1A regulates phosphorylation of Thr-186 in the Cdk9 T-loop. J Biol Chem. 2008;283(48):33578–84.
Muniz L, Egloff S, Ughy B, Jády BE, Kiss T. Controlling cellular P-TEFb activity by the HIV-1 transcriptional transactivator Tat. Cullen BR, editor. PLoS Pathog. 2010;6(10):e1001152.
Barboric M, Yik JHN, Czudnochowski N, Yang Z, Chen R, Contreras X, et al. Tat competes with HEXIM1 to increase the active pool of P-TEFb for HIV-1 transcription. Nucleic Acids Res. 2007;35(6):2003–12.
D’Orso I, Frankel AD. HIV-1 Tat: its dependence on host factors is crystal clear. Viruses. 2010;2(10):2226–34.
Nekhai S, Ammosova T, Charles S, Jeang K. Regulation of HIV-1 transcription by protein phosphatase 1. FASEB J. 2007;21(6):3–9.
Ammosova T, Obukhov Y, Kotelkin A, Breuer D, Beullens M, Gordeuk VR, et al. Protein phosphatase-1 activates CDK9 by dephosphorylating Ser175. Harrich D, editor. PLoS One. 2011;6(4):e18985.
Mbonye UR, Gokulrangan G, Datt M, Dobrowolski C, Cooper M, Chance MR, et al. Phosphorylation of CDK9 at Ser175 enhances HIV transcription and is a marker of activated P-TEFb in CD4+ T lymphocytes. Emerman M, editor. PLoS Pathog. 2013;9(5):e1003338.
Emanuel J, Marzi A, Feldmann H. Filoviruses: ecology, molecular biology, and evolution. In: Advances in virus research. Cambridge: Elsevier Inc.; 2018. p. 189–221. https://doi.org/10.1016/bs.aivir.2017.12.002.
Hargreaves A, Brady C, Mellors J, Tipton T, Carroll MW, Longet S. Filovirus neutralising antibodies: mechanisms of action and therapeutic application. Pathogens. 2021;10(9):1201.
Towner JS, Khristova ML, Sealy TK, Vincent MJ, Erickson BR, Bawiec DA, et al. Marburgvirus genomics and association with a large hemorrhagic fever outbreak in Angola. J Virol. 2006;80(13):6497–516.
Timothy JWS, Hall Y, Akoi-Boré J, Diallo B, Tipton TRW, Bower H, et al. Early transmission and case fatality of Ebola virus at the index site of the 2013–16 west African Ebola outbreak: a cross-sectional seroprevalence survey. Lancet Infect Dis. 2019;19(4):429–38.
Mühlberger E. Filovirus replication and transcription. Future Virol. 2007;2(2):205–15.
Bach S, Demper J-C, Grünweller A, Becker S, Biedenkopf N, Hartmann RK. Regulation of VP30-dependent transcription by RNA sequence and structure in the genomic Ebola virus promoter. Dutch RE, editor. J Virol. 2021;95(5):2215–20.
Biedenkopf N, Schlereth J, Grünweller A, Becker S, Hartmann RK. RNA binding of Ebola virus VP30 is essential for activating viral transcription. Lyles DS, editor. J Virol. 2016;90(16):7481–96.
Biedenkopf N, Lier C, Becker S. Dynamic phosphorylation of VP30 is essential for Ebola virus life cycle. Lyles DS, editor. J Virol. 2016;90(10):4914–25.
Schlereth J, Grünweller A, Biedenkopf N, Becker S, Hartmann RK. RNA binding specificity of Ebola virus transcription factor VP30. RNA Biol. 2016;13(9):783–98.
Biedenkopf N, Hartlieb B, Hoenen T, Becker S. Phosphorylation of Ebola virus VP30 Influences the composition of the viral nucleocapsid complex. J Biol Chem. 2013;288(16):11165–74.
Modrof J, Mühlberger E, Klenk H-D, Becker S. Phosphorylation of VP30 impairs Ebola virus transcription. J Biol Chem. 2002;277(36):33099–104.
Ilinykh PA, Tigabu B, Ivanov A, Ammosova T, Obukhov Y, Garron T, et al. Role of protein phosphatase 1 in dephosphorylation of Ebola virus VP30 protein and its targeting for the inhibition of viral transcription. J Biol Chem. 2014;289(33):22723–38.
Llorian M, Beullens M, Lesage B, Nicolaescu E, Beke L, Landuyt W, et al. Nucleocytoplasmic shuttling of the splicing factor SIPP1. J Biol Chem. 2005;280(46):38862–9.
Tigabu B, Ramanathan P, Ivanov A, Lin X, Ilinykh PA, Parry CS, et al. Phosphorylated VP30 of marburg virus is a repressor of transcription. Dermody TS, editor. J Virol. 2018;92(21):1–20.
Mühlberger E, Lötfering B, Klenk H-D, Becker S. Three of the four nucleocapsid proteins of Marburg virus, NP, VP35, and L, are sufficient to mediate replication and transcription of Marburg Virus-specific monocistronic minigenomes. J Virol. 1998;72(11):8756–64.
Mühlberger E, Weik M, Volchkov VE, Klenk H-D, Becker S. Comparison of the transcription and replication strategies of Marburg virus and Ebola virus by using artificial replication systems. J Virol. 1999;73(3):2333–42.
Fowler T, Bamberg S, Möller P, Klenk H-D, Meyer TF, Becker S, et al. Inhibition of Marburg virus protein expression and viral release by RNA interference. J Gen Virol. 2005;86(4):1181–8.
Kruse T, Biedenkopf N, Hertz EPT, Dietzel E, Stalmann G, López-Méndez B, et al. The Ebola Virus nucleoprotein recruits the host PP2A-B56 phosphatase to activate transcriptional support activity of VP30. Mol Cell. 2018;69(1):136-145.e6.
Schweitzer A, Horn J, Mikolajczyk RT, Krause G, Ott JJ. Estimations of worldwide prevalence of chronic hepatitis B virus infection: a systematic review of data published between 1965 and 2013. Lancet. 2015;386(10003):1546–55.
Yen C-J, Yang S-T, Chen R-Y, Huang W, Chayama K, Lee M-H, et al. Hepatitis B virus X protein (HBx) enhances centrosomal P4.1-associated protein (CPAP) expression to promote hepatocarcinogenesis. J Biomed Sci. 2019;26(1):44.
Cougot D, Allemand E, Rivière L, Benhenda S, Duroure K, Levillayer F, et al. Inhibition of PP1 phosphatase activity by HBx: a mechanism for the activation of hepatitis B virus transcription. Sci Signal. 2012;5(205):1–12.
Tang H-MV, Gao W-W, Chan C-P, Cheng Y, Chaudhary V, Deng J-J, et al. Requirement of CRTC1 coactivator for hepatitis B virus transcription. Nucleic Acids Res. 2014;42(20):12455–68.
Steven A, Friedrich M, Jank P, Heimer N, Budczies J, Denkert C, et al. What turns CREB on? And off? And why does it matter? Cell Mol Life Sci. 2020;77(20):4049–67.
Dyson HJ, Wright PE. Role of intrinsic protein disorder in the function and interactions of the transcriptional coactivators CREB-binding Protein (CBP) and p300. J Biol Chem. 2016;291(13):6714–22.
Canettieri G, Morantte I, Guzmán E, Asahara H, Herzig S, Anderson SD, et al. Attenuation of a phosphorylation-dependent activator by an HDAC–PP1 complex. Nat Struct Mol Biol. 2003;10(3):175–81.
Sakamoto K, Karelina K, Obrietan K. CREB: a multifaceted regulator of neuronal plasticity and protection. J Neurochem. 2011;116(1):1–9.
Hu Z, Ban H, Zheng H, Liu M, Chang J, Guo J-T. Protein phosphatase 1 catalyzes HBV core protein dephosphorylation and is co-packaged with viral pregenomic RNA into nucleocapsids. Hu J, editor. PLoS Pathog. 2020;16(7):e1008669.
Venkatakrishnan B, Zlotnick A. The structural biology of hepatitis B virus: form and function. Annu Rev Virol. 2016;3(1):429–51.
Seeger C, Mason WS. Molecular biology of hepatitis B virus infection. Virology. 2015;479–480:672–86.
Jung J, Hwang SG, Chwae Y-J, Park S, Shin H-J, Kim K. Phosphoacceptors threonine 162 and serines 170 and 178 within the carboxyl-terminal RRRS/T motif of the hepatitis B virus core protein make multiple contributions to hepatitis B virus replication. Sandri-Goldin RM, editor. J Virol. 2014;88(16):8754–67.
Perlman DH, Berg EA, O’Connor PB, Costello CE, Hu J. Reverse transcription-associated dephosphorylation of hepadnavirus nucleocapsids. Proc Natl Acad Sci. 2005;102(25):9020–5.
Su P-Y, Yang C-J, Chu T-H, Chang C-H, Chiang C, Tang F-M, et al. HBV maintains electrostatic homeostasis by modulating negative charges from phosphoserine and encapsidated nucleic acids. Sci Rep. 2016;6(1):38959.
Zhao Q, Hu Z, Cheng J, Wu S, Luo Y, Chang J, et al. Hepatitis B virus core protein dephosphorylation occurs during pregenomic RNA encapsidation. Ou J-HJ, editor. J Virol. 2018;92(13):2139–17.
Basagoudanavar SH, Perlman DH, Hu J. Regulation of hepadnavirus reverse transcription by dynamic nucleocapsid phosphorylation. J Virol. 2007;81(4):1641–9.
Ning X, Basagoudanavar SH, Liu K, Luckenbaugh L, Wei D, Wang C, et al. Capsid phosphorylation state and hepadnavirus virion secretion. McFadden G, editor. J Virol. 2017;91(9):1–16.
Luo J, Xi J, Gao L, Hu J. Role of hepatitis B virus capsid phosphorylation in nucleocapsid disassembly and covalently closed circular DNA formation. Siddiqui A, editor. PLOS Pathog. 2020;16(3):e1008459.
Liu CWY, Wang R-H, Dohadwala M, Schönthal AH, Villa-Moruzzi E, Berndt N. Inhibitory phosphorylation of PP1α catalytic subunit during the G1/S transition. J Biol Chem. 1999;274(41):29470–5.
Louten J. Herpesviruses. In: Essential human virology. Cambridge: Elsevier; 2016. p. 235–56. https://doi.org/10.1016/B978-0-323-90565-7.00013-7.
Payne S. Family herpesviridae. In: Viruses. Cambridge: Elsevier; 2017. p. 269–78. https://doi.org/10.1016/B978-0-12-803109-4.00034-9.
Benedyk TH, Muenzner J, Connor V, Han Y, Brown K, Wijesinghe KJ, et al. pUL21 is a viral phosphatase adaptor that promotes herpes simplex virus replication and spread. DeLuca NA, editor. PLoS Pathog. 2021;17(8):e1009824.
Le Sage V, Jung M, Alter JD, Wills EG, Johnston SM, Kawaguchi Y, et al. The herpes simplex virus 2 UL21 protein is essential for virus propagation. J Virol. 2013;87(10):5904–15.
Mbong EF, Woodley L, Frost E, Baines JD, Duffy C. Deletion of U L 21 causes a delay in the early stages of the herpes simplex virus 1 replication cycle. J Virol. 2012;86(12):7003–7.
Han J, Chadha P, Meckes DG, Baird NL, Wills JW. Interaction and interdependent packaging of tegument protein UL11 and glycoprotein E of herpes simplex virus. J Virol. 2011;85(18):9437–46.
Gao J, Finnen RL, Sherry MR, Le Sage V, Banfield BW. Differentiating the roles of UL16, UL21, and Us3 in the nuclear egress of herpes simplex virus capsids. Sandri-Goldin RM, editor. J Virol. 2020;94(13):1–16.
Kumagai K, Kawano-Kawada M, Hanada K. Phosphoregulation of the Ceramide Transport Protein CERT at Serine 315 in the Interaction with VAMP-associated Protein (VAP) for inter-organelle trafficking of ceramide in mammalian cells. J Biol Chem. 2014;289(15):10748–60.
Kumagai K, Hanada K. Structure, functions and regulation of CERT, a lipid-transfer protein for the delivery of ceramide at the ER–Golgi membrane contact sites. FEBS Lett. 2019;593(17):2366–77.
Benedyk TH, Connor V, Caroe ER, Shamin M, Svergun DI, Deane JE, et al. Herpes simplex virus 1 protein pUL21 alters ceramide metabolism by activating the interorganelle transport protein CERT. J Biol Chem. 2022;298(11):102589.
Ma Z, Bai J, Jiang C, Zhu H, Liu D, Pan M, et al. Tegument protein UL21 of alpha-herpesvirus inhibits the innate immunity by triggering CGAS degradation through TOLLIP-mediated selective autophagy. Autophagy. 2022;00(00):1–21.
Masoud Bahnamiri M, Roller RJ. Mechanism of nuclear lamina disruption and the role of pUS3 in herpes simplex virus 1 nuclear egress. Longnecker RM, editor. J Virol. 2021;95(10):e02432-20.
Dalton LE, Healey E, Irving J, Marciniak SJ. Phosphoproteins in stress-induced disease. In: Progress in molecular biology and translational science. 1st ed. Cambridge: Elsevier Inc.; 2012. p. 189–221. https://doi.org/10.1016/B978-0-12-396456-4.00003-1.
Sudhakar A, Ramachandran A, Ghosh S, Hasnain SE, Kaufman RJ, Ramaiah KVA. Phosphorylation of Serine 51 in Initiation Factor 2α (eIF2α) Promotes Complex Formation between eIF2α(P) and eIF2B and causes inhibition in the guanine nucleotide exchange activity of eIF2B. Biochemistry. 2000;39(42):12929–38.
Fusade-Boyer M, Dupré G, Bessière P, Khiar S, Quentin-Froignant C, Beck C, et al. Evaluation of the antiviral activity of Sephin1 treatment and its consequences on eIF2α phosphorylation in response to viral infections. Front Immunol. 2019;10:1–12.
Jousse C, Oyadomari S, Novoa I, Lu P, Zhang Y, Harding HP, et al. Inhibition of a constitutive translation initiation factor 2α phosphatase, CReP, promotes survival of stressed cells. J Cell Biol. 2003;163(4):767–75.
Novoa I, Zeng H, Harding HP, Ron D. Feedback inhibition of the unfolded protein response by GADD34-mediated dephosphorylation of eIF2α. J Cell Biol. 2001;153(5):1011–22.
Liu Y, Wang M, Cheng A, Yang Q, Wu Y, Jia R, et al. The role of host eIF2α in viral infection. Virol J. 2020;17(1):1–15.
Chou J, Roizman B. The herpes simplex virus 1 gene for ICP34.5, which maps in inverted repeats, is conserved in several limited-passage isolates but not in strain 17syn+. J Virol. 1990;64(3):1014–20.
Rojas M, Vasconcelos G, Dever TE. An eIF2α-binding motif in protein phosphatase 1 subunit GADD34 and its viral orthologs is required to promote dephosphorylation of eIF2α. Proc Natl Acad Sci. 2015;112(27):E3466–75.
He B, Gross M, Roizman B. The γ134.5 protein of herpes simplex virus 1 has the structural and functional attributes of a protein phosphatase 1 regulatory subunit and is present in a high molecular weight complex with the enzyme in infected cells. J Biol Chem. 1998;273(33):20737–43.
He B, Gross M, Roizman B. The γ 1 34.5 protein of herpes simplex virus 1 complexes with protein phosphatase 1α to dephosphorylate the α subunit of the eukaryotic translation initiation factor 2 and preclude the shutoff of protein synthesis by double-stranded RNA-activated protein. Proc Natl Acad Sci. 1997;94(3):843–8.
Cheng G, Brett M-E, He B. Val193 and Phe195 of the γ134.5 protein of herpes simplex virus 1 are required for viral resistance to interferon-α/β. Virology. 2001;290(1):115–20.
Verpooten D, Feng Z, Valyi-Nagy T, Ma Y, Jin H, Yan Z, et al. Dephosphorylation of eIF2α mediated by the γ 1 34.5 protein of herpes simplex virus 1 facilitates viral neuroinvasion. J Virol. 2009;83(23):12626–30.
Cheng G, Gross M, Brett M-E, He B. AlaArg motif in the carboxyl terminus of the γ 1 34.5 protein of herpes simplex virus type 1 is required for the formation of a high-molecular-weight complex that dephosphorylates eIF-2α. J Virol. 2001;75(8):3666–74.
Li Y, Zhang C, Chen X, Yu J, Wang Y, Yang Y, et al. ICP34.5 protein of herpes simplex virus facilitates the initiation of protein translation by bridging eukaryotic Initiation Factor 2α (eIF2α) and protein phosphatase 1. J Biol Chem. 2011;286(28):24785–92.
Cerveny M, Hessefort S, Yang K, Cheng G, Gross M, He B. Amino acid substitutions in the effector domain of the γ134.5 protein of herpes simplex virus 1 have differential effects on viral response to interferon-α. Virology. 2003;307(2):290–300.
Zhang C, Tang J, Xie J, Zhang H, Li Y, Zhang J, et al. A conserved domain of herpes simplex virus ICP34.5 regulates protein phosphatase complex in mammalian cells. FEBS Lett. 2008;582(2):171–6.
Meng W, Han S-C, Li C-C, Dong H-J, Wang X-J. Multifunctional viral protein γ34.5 manipulates nucleolar protein NOP53 for optimal viral replication of HSV-1. Cell Death Dis. 2018;9(2):103.
Barber C, Netherton C, Goatley L, Moon A, Goodbourn S, Dixon L. Identification of residues within the African swine fever virus DP71L protein required for dephosphorylation of translation initiation factor eIF2α and inhibiting activation of pro-apoptotic CHOP. Virology. 2017;504:107–13.
Galindo I, Hernáez B, Muñoz-Moreno R, Cuesta-Geijo MA, Dalmau-Mena I, Alonso C. The ATF6 branch of unfolded protein response and apoptosis are activated to promote African swine fever virus infection. Cell Death Dis. 2012;3(7):e341–e341.
Rivera J, Abrams C, Hernáez B, Alcázar A, Escribano JM, Dixon L, et al. The MyD116 African swine fever virus homologue interacts with the catalytic subunit of protein phosphatase 1 and activates its phosphatase activity. J Virol. 2007;81(6):2923–9.
Zhang F, Moon A, Childs K, Goodbourn S, Dixon LK. The African swine fever virus DP71L protein recruits the protein phosphatase 1 catalytic subunit to dephosphorylate eIF2α and inhibits CHOP induction but is dispensable for these activities during virus infection. J Virol. 2010;84(20):10681–9.
Cruz JLG, Sola I, Becares M, Alberca B, Plana J, Enjuanes L, et al. Coronavirus gene 7 counteracts host defenses and modulates virus virulence. PLoS Pathog. 2011;7(6):e1002090.
Cruz JLG, Becares M, Sola I, Oliveros JC, Enjuanes L, Zúñiga S. Alphacoronavirus protein 7 modulates host innate immune response. J Virol. 2013;87(17):9754–67.
McDermott JE, Mitchell HD, Gralinski LE, Eisfeld AJ, Josset L, Bankhead A, et al. The effect of inhibition of PP1 and TNFα signaling on pathogenesis of SARS coronavirus. BMC Syst Biol. 2016;10(1):93.
Umareddy I, Pluquet O, Wang QY, Vasudevan SG, Chevet E, Gu F. Dengue virus serotype infection specifies the activation of the unfolded protein response. Virol J. 2007;4(1):91.
Ruggieri A, Dazert E, Metz P, Hofmann S, Bergeest J-P, Mazur J, et al. Dynamic oscillation of translation and stress granule formation mark the cellular response to virus infection. Cell Host Microbe. 2012;12(1):71–85.
Liao Y, Gu F, Mao X, Niu Q, Wang H, Sun Y, et al. Regulation of de novo translation of host cells by manipulation of PERK/PKR and GADD34-PP1 activity during Newcastle disease virus infection. J Gen Virol. 2016;97(4):867–79.
Kazemi S, Papadopoulou S, Li S, Su Q, Wang S, Yoshimura A, et al. Control of α subunit of eukaryotic translation initiation Factor 2 (eIF2α) phosphorylation by the human papillomavirus type 18 E6 oncoprotein: implications for eIF2α-dependent gene expression and cell death. Mol Cell Biol. 2004;24(8):3415–29.
Xu S, Chen D, Chen D, Hu Q, Zhou L, Ge X, et al. Pseudorabies virus infection inhibits stress granules formation via dephosphorylating eIF2α. Vet Microbiol. 2020;247:108786.
Zhu T, Jiang X, Xin H, Zheng X, Xue X, Chen J-L, et al. GADD34-mediated dephosphorylation of eIF2α facilitates pseudorabies virus replication by maintaining de novo protein synthesis. Vet Res. 2021;52(1):148.
Wang X, Liao Y, Yap PL, Png KJ, Tam JP, Liu DX. Inhibition of protein kinase R activation and upregulation of GADD34 expression play a synergistic role in facilitating coronavirus replication by maintaining de novo protein synthesis in virus-infected Cells. J Virol. 2009;83(23):12462–72.
Urra H, Dufey E, Lisbona F, Rojas-Rivera D, Hetz C. When ER stress reaches a dead end. Biochim Biophys Acta Mol Cell Res. 2013;1833(12):3507–17.
Rehwinkel J, Gack MU. RIG-I-like receptors: their regulation and roles in RNA sensing. Nat Rev Immunol. 2020;20(9):537–51.
Rodriguez KR, Bruns AM, Horvath CM. MDA5 and LGP2: accomplices and antagonists of antiviral signal transduction. Goff SP, editor. J Virol. 2014;88(15):8194–200.
Kell AM, Gale M. RIG-I in RNA virus recognition. Virology. 2015;479–480:110–21.
Ferreira AR, Marques M, Ramos B, Kagan JC, Ribeiro D. Emerging roles of peroxisomes in viral infections. Trends Cell Biol. 2022;32(2):124–39.
Chan C-P, Jin D-Y. Cytoplasmic RNA sensors and their interplay with RNA-binding partners in innate antiviral response: theme and variations. RNA. 2022;28(4):449–77.
Paz S, Vilasco M, Werden SJ, Arguello M, Joseph-Pillai D, Zhao T, et al. A functional C-terminal TRAF3-binding site in MAVS participates in positive and negative regulation of the IFN antiviral response. Cell Res. 2011;21(6):895–910.
Liu Y, Olagnier D, Lin R. Host and viral modulation of RIG-I-mediated antiviral immunity. Front Immunol. 2017;7:1–12.
Maharaj NP, Wies E, Stoll A, Gack MU. Conventional Protein Kinase C-α (PKC-α) and PKC-β negatively regulate RIG-I antiviral signal transduction. J Virol. 2012;86(3):1358–71.
Nistal-Villán E, Gack MU, Martínez-Delgado G, Maharaj NP, Inn K-S, Yang H, et al. Negative role of RIG-I serine 8 phosphorylation in the regulation of interferon-β production. J Biol Chem. 2010;285(26):20252–61.
Wies E, Wang MK, Maharaj NP, Chen K, Zhou S, Finberg RW, et al. Dephosphorylation of the RNA sensors RIG-I and MDA5 by the phosphatase PP1 is essential for innate immune signaling. Immunity. 2013;38(3):437–49.
Hu MM, Liao CY, Yang Q, Xie XQ, Shu HB. Innate immunity to RNA virus is regulated by temporal and reversible sumoylation of RIG-I and MDA5. J Exp Med. 2017;214(4):973–89.
Acharya D, Reis R, Volcic M, Liu G, Wang MK, Chia BS, et al. Actin cytoskeleton remodeling primes RIG-I-like receptor activation. Cell. 2022;185(19):3588-3602.e21.
Tan I, Ng CH, Lim L, Leung T. Phosphorylation of a novel myosin binding subunit of protein phosphatase 1 reveals a conserved mechanism in the regulation of actin cytoskeleton. J Biol Chem. 2001;276(24):21209–16.
Mura M, Combredet C, Najburg V, Sanchez David RY, Tangy F, Komarova AV. Nonencapsidated 5’ copy-back defective interfering genomes produced by recombinant measles viruses are recognized by RIG-I and LGP2 but not MDA5. García-Sastre A, editor. J Virol. 2017;91(20):1–22.
Runge S, Sparrer KMJ, Lässig C, Hembach K, Baum A, García-Sastre A, et al. In vivo ligands of MDA5 and RIG-I in measles virus-infected cells. Mossman KL, editor. PLoS Pathog. 2014;10(4):e1004081.
Ayasoufi K, Pfaller CK. Seek and hide: the manipulating interplay of measles virus with the innate immune system. Curr Opin Virol. 2020;41:18–30.
Seya T. Measles virus takes a two-pronged attack on PP1. Cell Host Microbe. 2014;16(1):1–2.
Mesman AW, Zijlstra-Willems EM, Kaptein TM, de Swart RL, Davis ME, Ludlow M, et al. Measles virus suppresses RIG-I-like receptor activation in dendritic cells via DC-SIGN-mediated inhibition of PP1 phosphatases. Cell Host Microbe. 2014;16(1):31–42.
Nagano Y, Sugiyama A, Kimoto M, Wakahara T, Noguchi Y, Jiang X, et al. The measles virus V protein binding site to STAT2 overlaps that of IRF9. López S, editor. J Virol. 2020;94(17):e01169-20.
Mandhana R, Qian LK, Horvath CM. Constitutively active MDA5 proteins are inhibited by paramyxovirus v proteins. J Interf Cytokine Res. 2018;38(8):319–32.
Davis ME, Wang MK, Rennick LJ, Full F, Gableske S, Mesman AW, et al. Antagonism of the phosphatase PP1 by the measles virus V protein is required for innate immune escape of MDA5. Cell Host Microbe. 2014;16(1):19–30.
Yang Y, Ye F, Zhu N, Wang W, Deng Y, Zhao Z, et al. Middle East respiratory syndrome coronavirus ORF4b protein inhibits type I interferon production through both cytoplasmic and nuclear targets. Sci Rep. 2015;5(1):17554.
Fang P, Fang L, Xia S, Ren J, Zhang J, Bai D, et al. Porcine deltacoronavirus accessory protein NS7a antagonizes IFN-β production by competing With TRAF3 and IRF3 for binding to IKKε. Front Cell Infect Microbiol. 2020;10:1–16.
Ji L, Wang N, Ma J, Cheng Y, Wang H, Sun J, et al. Porcine deltacoronavirus nucleocapsid protein species-specifically suppressed IRF7-induced type I interferon production via ubiquitin-proteasomal degradation pathway. Vet Microbiol. 2020;250:108853.
Siu K-L, Yeung ML, Kok K-H, Yuen K-S, Kew C, Lui P-Y, et al. Middle East respiratory syndrome coronavirus 4a protein is a double-stranded RNA-binding protein that suppresses PACT-induced activation of RIG-I and MDA5 in the innate antiviral response. Perlman S, editor. J Virol. 2014;88(9):4866–76.
Fang P, Fang L, Ren J, Hong Y, Liu X, Zhao Y, et al. Porcine deltacoronavirus accessory protein NS6 antagonizes interferon beta production by interfering with the binding of RIG-I/MDA5 to double-stranded RNA. J Virol. 2018;92(15):1–17.
Zhang J, Fang P, Ren J, Xia S, Zhang H, Zhu X, et al. Porcine epidemic diarrhea virus nsp7 inhibits MDA5 dephosphorylation to antagonize type I interferon production. Sinclair A, editor. Microbiol Spectr. 2023;11(2):e0501722.
Takashima K, Oshiumi H, Takaki H, Matsumoto M, Seya T. RIOK3-mediated phosphorylation of MDA5 interferes with its assembly and attenuates the innate immune response. Cell Rep. 2015;11(2):192–200.
Gao D, Li W. Structures and recognition modes of toll-like receptors. Proteins. 2017;85(1):3–9.
Fitzgerald KA, Kagan JC. Toll-like receptors and the control of immunity. Cell. 2020;180(6):1044–66.
Sartorius R, Trovato M, Manco R, D’Apice L, De Berardinis P. Exploiting viral sensing mediated by Toll-like receptors to design innovative vaccines. NPJ Vaccines. 2021;6(1):127.
Komal A, Noreen M, El-Kott AF. TLR3 agonists: RGC100, ARNAX, and poly-IC: a comparative review. Immunol Res. 2021;69(4):312–22.
Seumen CHT, Grimm TM, Hauck CR. Protein phosphatases in TLR signaling. Cell Commun Signal. 2021;19(1):1–18.
Brown J, Wang H, Hajishengallis GN, Martin M. TLR-signaling networks: an integration of adaptor molecules, kinases, and cross-talk. J Dent Res. 2011;90(4):417–27.
Ouyang C, Nie L, Gu M, Wu A, Han X, Wang X, et al. Transforming Growth Factor (TGF)-β-activated Kinase 1 (TAK1) activation requires phosphorylation of serine 412 by Protein Kinase A Catalytic Subunit α (PKACα) and X-linked Protein Kinase (PRKX). J Biol Chem. 2014;289(35):24226–37.
Scholz R, Sidler CL, Thali RF, Winssinger N, Cheung PCF, Neumann D. Autoactivation of transforming growth factor β-activated Kinase 1 is a sequential bimolecular process. J Biol Chem. 2010;285(33):25753–66.
Gu M, Ouyang C, Lin W, Zhang T, Cao X, Xia Z, et al. Phosphatase holoenzyme PP1/GADD34 negatively regulates TLR response by inhibiting TAK1 serine 412 phosphorylation. J Immunol. 2014;192(6):2846–56.
Gilmore TD. Introduction to NF-κB: players, pathways, perspectives. Oncogene. 2006;25(51):6680–4.
Hinz M, Scheidereit C. The IκB kinase complex in NF-κB regulation and beyond. EMBO Rep. 2014;15(1):46–61.
Li H-Y, Liu H, Wang C-H, Zhang J-Y, Man J-H, Gao Y-F, et al. Deactivation of the kinase IKK by CUEDC2 through recruitment of the phosphatase PP1. Nat Immunol. 2008;9(5):533–41.
Panne D, McWhirter SM, Maniatis T, Harrison SC. Interferon regulatory factor 3 is regulated by a dual phosphorylation-dependent switch. J Biol Chem. 2007;282(31):22816–22.
Mcwhirter SM, Fitzgerald KA, Rosains J, Rowe DC, Golenbock DT, Maniatis T. IFN-regulatory factor 3-dependent gene expression is defective in Tbk1-deficient mouse embryonic fibroblasts. 2003
Lin R, Mamane Y, Hiscott J. Multiple regulatory domains control IRF-7 activity in response to virus infection. J Biol Chem. 2000;275(44):34320–7.
Wang L, Zhao J, Ren J, Hall KH, Moorman JP, Yao ZQ, et al. Protein phosphatase 1 abrogates IRF7-mediated type I IFN response in antiviral immunity. Eur J Immunol. 2016;46(10):2409–19.
Gu M, Zhang T, Lin W, Liu Z, Lai R, Xia D, et al. Protein phosphatase PP1 negatively regulates the Toll-like receptor- and RIG-I-like receptor-triggered production of type I interferon by inhibiting IRF3 phosphorylation at serines 396 and 385 in macrophage. Cell Signal. 2014;26(12):2930–9.
Li Q, Zheng Z, Liu Y, Zhang Z, Liu Q, Meng J, et al. 2C proteins of enteroviruses suppress IKKβ phosphorylation by recruiting protein phosphatase 1. López S, editor. J Virol. 2016;90(10):5141–51.
Jin H, Yan Z, Ma Y, Cao Y, He B. A herpesvirus virulence factor inhibits dendritic cell maturation through protein phosphatase 1 and IκB Kinase. J Virol. 2011;85(7):3397–407.
Dorrington MG, Fraser IDC. NF-κB signaling in macrophages: dynamics, crosstalk, and signal integration. Front Immunol. 2019;10:705.
Opaluch AM, Schneider M, Chiang C, Nguyen QT, Maestre AM, Mulder LCF, et al. Positive regulation of TRAF6-dependent innate immune responses by protein phosphatase PP1-γ. Allen IC, editor. PLoS One. 2014;9(2):e89284.
Chelbi-Alix MK, Wietzerbin J. Interferon, a growing cytokine family: 50 years of interferon research. Biochimie. 2007;89(6–7):713–8.
Bulli L, Apolonia L, Kutzner J, Pollpeter D, Goujon C, Herold N, et al. Complex interplay between HIV-1 capsid and MX2-independent alpha interferon-induced antiviral factors. Sundquist WI, editor. J Virol. 2016;90(16):7469–80.
Liu Z, Pan Q, Ding S, Qian J, Xu F, Zhou J, et al. The interferon-inducible MxB protein inhibits HIV-1 infection. Cell Host Microbe. 2013;14(4):398–410.
Dicks MDJ, Betancor G, Jimenez-Guardeño JM, Pessel-Vivares L, Apolonia L, Goujon C, et al. Multiple components of the nuclear pore complex interact with the amino-terminus of MX2 to facilitate HIV-1 restriction. Emerman M, editor. PLoS Pathog. 2018;14(11):e1007408.
Betancor G, Dicks MDJ, Jimenez-Guardeño JM, Ali NH, Apolonia L, Malim MH. The GTPase domain of MX2 Interacts with the HIV-1 capsid, enabling its short isoform to moderate antiviral restriction. Cell Rep. 2019;29(7):1923-1933.e3.
Schilling M, Bulli L, Weigang S, Graf L, Naumann S, Patzina C, et al. Human MxB protein is a pan-herpesvirus restriction factor. Sandri-Goldin RM, editor. J Virol. 2018;92(17):1–11.
Crameri M, Bauer M, Caduff N, Walker R, Steiner F, Franzoso FD, et al. MxB is an interferon-induced restriction factor of human herpesviruses. Nat Commun. 2018;9(1):1980.
Kane M, Yadav SS, Bitzegeio J, Kutluay SB, Zang T, Wilson SJ, et al. MX2 is an interferon-induced inhibitor of HIV-1 infection. Nature. 2013;502(7472):563–6.
Goujon C, Moncorgé O, Bauby H, Doyle T, Ward CC, Schaller T, et al. Human MX2 is an interferon-induced post-entry inhibitor of HIV-1 infection. Nature. 2013;502(7472):559–62.
Betancor G, Jimenez-Guardeño JM, Lynham S, Antrobus R, Khan H, Sobala A, et al. MX2-mediated innate immunity against HIV-1 is regulated by serine phosphorylation. Nat Microbiol. 2021;6(8):1031–42.
Ahmed AA, Abouzid M. Arbidol targeting influenza virus A Hemagglutinin; a comparative study. Biophys Chem. 2021;277:106663.
Fatma B, Kumar R, Singh VA, Nehul S, Sharma R, Kesari P, et al. Alphavirus capsid protease inhibitors as potential antiviral agents for Chikungunya infection. Antiviral Res. 2020;179:104808.
Mudgal R, Mahajan S, Tomar S. Inhibition of Chikungunya virus by an adenosine analog targeting the SAM-dependent nsP1 methyltransferase. FEBS Lett. 2020;594(4):678–94.
Irwin KK, Renzette N, Kowalik TF, Jensen JD. Antiviral drug resistance as an adaptive process. Virus Evol. 2016;2(1):vew014.
Mahajan S, Choudhary S, Kumar P, Tomar S. Antiviral strategies targeting host factors and mechanisms obliging +ssRNA viral pathogens. Bioorg Med Chem. 2021;46:116356.
Kumar N, Sharma S, Kumar R, Tripathi BN, Barua S, Ly H, et al. Host-directed antiviral therapy. Clin Microbiol Rev. 2020;33(3):116356.
Mullard A. Phosphatases start shedding their stigma of undruggability. Nat Rev Drug Discov. 2018;17(12):847–9.
Fontanillo M, Zemskov I, Häfner M, Uhrig U, Salvi F, Simon B, et al. Synthesis of highly selective submicromolar microcystin-based inhibitors of Protein Phosphatase (PP)2A over PP1. Angew Chemie Int Ed. 2016;55(45):13985–9.
Choy MS, Swingle M, D’Arcy B, Abney K, Rusin SF, Kettenbach AN, et al. PP1: tautomycetin complex reveals a path toward the development of PP1-specific inhibitors. J Am Chem Soc. 2017;139(49):17703–6.
Köhn M. Miklós Bodanszky Award Lecture: advances in the selective targeting of protein phosphatase-1 and phosphatase-2A with peptides. J Pept Sci. 2017;23(10):749–56.
Wang T, Gao H, Li W, Liu C. Essential role of histone replacement and modifications in male fertility. Front Genet. 2019;10:1–15.
Fischer TH, Eiringhaus J, Dybkova N, Saadatmand A, Pabel S, Weber S, et al. Activation of protein phosphatase 1 by a selective phosphatase disrupting peptide reduces sarcoplasmic reticulum Ca 2+ leak in human heart failure. Eur J Heart Fail. 2018;20(12):1673–85.
Silva JV, Freitas MJ, Santiago J, Jones S, Guimarães S, Vijayaraghavan S, et al. Disruption of protein phosphatase 1 complexes with the use of bioportides as a novel approach to target sperm motility. Fertil Steril. 2021;115(2):348–62.
Potenza DM, Janicek R, Fernandez-Tenorio M, Niggli E. Activation of endogenous protein phosphatase 1 enhances the calcium sensitivity of the ryanodine receptor type 2 in murine ventricular cardiomyocytes. J Physiol. 2020;598(6):1131–50.
Ammosova T, Platonov M, Yedavalli VRK, Obukhov Y, Gordeuk VR, Jeang K-T, et al. Small molecules targeted to a non-catalytic “RVxF” binding site of protein phosphatase-1 inhibit HIV-1. Kehn-Hall K, editor. PLoS One. 2012;7(6):e39481.
Ammosova T, Platonov M, Ivanov A, SaygideğerKont Y, Kumari N, Kehn-Hall K, et al. 1E7–03, a small molecule targeting host protein phosphatase-1, inhibits HIV-1 Transcription. Br J Pharmacol. 2014;171(22):5059–75.
Lin X, Kumari N, DeMarino C, Saygideğer Kont Y, Ammosova T, Kulkarni A, et al. Inhibition of HIV-1 infection in humanized mice and metabolic stability of protein phosphatase-1-targeting small molecule 1E7-03. Oncotarget. 2017;8(44):76749–69.
Lin X, Sajith AM, Wang S, Kumari N, Choy MS, Ahmad A, et al. Structural Optimization of 2,3-Dihydro-1H-cyclopenta[b ]quinolines targeting the noncatalytic RVxF site of protein phosphatase 1 for HIV-1 inhibition. ACS Infect Dis. 2020;6(12):3190–211.
Baer A, Shafagati N, Benedict A, Ammosova T, Ivanov A, Hakami RM, et al. Protein phosphatase-1 regulates Rift Valley fever virus replication. Antiviral Res. 2016;127:79–89.
Carey BD, Ammosova T, Pinkham C, Lin X, Zhou W, Liotta LA, et al. Protein phosphatase 1α Interacts with Venezuelan equine encephalitis virus capsid protein and regulates viral replication through modulation of capsid phosphorylation. García-Sastre A, editor. J Virol. 2018;92(15):e02068-17.
Lin X, Ammosova T, Choy MS, Pietzsch CA, Ivanov A, Ahmad A, et al. Targeting the non-catalytic RVxF site of protein phosphatase-1 with small molecules for Ebola virus inhibition. Front Microbiol. 2019;10:1–17.
Ammosova T, Pietzsch CA, Saygideğer Y, Ilatovsky A, Lin X, Ivanov A, et al. Protein phosphatase 1–targeting small-molecule C31 inhibits Ebola virus replication. J Infect Dis. 2018;218(suppl_5):S627-35.
Wensing AM, Calvez V, Günthard HF, Johnson VA, Paredes R, Pillay D, et al. 2017 update of the drug resistance mutations in HIV-1. Top Antivir Med. 2016;24(4):132–3.
Vanhamel J, Bruggemans A, Debyser Z. Establishment of latent HIV-1 reservoirs: what do we really know? J Virus Erad. 2019;5(1):3–9.
Henderson LJ, Johnson TP, Smith BR, Reoma LB, Santamaria UA, Bachani M, et al. Presence of Tat and transactivation response element in spinal fluid despite antiretroviral therapy. AIDS. 2019;33(Supplement 2):S145–57.
Tyagi M, Iordanskiy S, Ammosova T, Kumari N, Smith K, Breuer D, et al. Reactivation of latent HIV-1 provirus via targeting protein phosphatase-1. Retrovirology. 2015;12(1):63.
Zhong L, Hu M-M, Bian L-J, Liu Y, Chen Q, Shu H-B. Phosphorylation of cGAS by CDK1 impairs self-DNA sensing in mitosis. Cell Discov. 2020;6(1):26.
Funding
This research was funded by the Institute for Biomedicine—iBiMED (grant number UIDB/04501/2020) and by individual grants to P.O.C. (grant number 2020.10111.BD) and D.R. (CEECIND/03747/2017) financed by the Portuguese Foundation for Science and Technology (FCT) of the Portuguese Ministry of Science and Higher Education and by the European Union (QREN, FEDER and COMPETE frameworks). It was also funded by EU through the Horizon 2020 program: H2020-WIDESPREAD-2020-5 ID-952373.
Author information
Authors and Affiliations
Contributions
Pedro O. Corda: conceptualization; writing - original draft; writing – review and editing; figures preparation. Mathieu Bollen: writing – review and editing. Daniela Ribeiro: conceptualization; writing – review and editing. Margarida Fardilha: conceptualization; writing – review and editing.
Corresponding authors
Ethics declarations
Ethics approval and consent to participate
Not applicable.
Consent for publication
Not applicable.
Competing interests
The authors declare no competing interests.
Additional information
Publisher’s Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.
About this article
Cite this article
Corda, P.O., Bollen, M., Ribeiro, D. et al. Emerging roles of the Protein Phosphatase 1 (PP1) in the context of viral infections. Cell Commun Signal 22, 65 (2024). https://doi.org/10.1186/s12964-023-01468-8
Received:
Accepted:
Published:
DOI: https://doi.org/10.1186/s12964-023-01468-8