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Another approach to MARV vaccines is the use of viral vectors expressing MARV GP. To date, two different systems have been established based on replication-defective adenoviral vectors or recombinant VSV expressing MARV GP. The adenovirus-based vaccine successfully protects guinea pigs and NHPs, and provides cross-protection. High levels of cross-reactive MARV-specific IgG and T cell responses are induced, indicating an induction of an immune response [248, 254] . Preexisting immunity against the adenovirus strain Ad5 might pose a problem for its successful use in humans (reviewed in [245] ). |
The VSV-based vaccine completely protects NHPs and additionally has proven successful in post-exposure treatment (reviewed in [255] ). A single immunization with recombinant VSV expressing MARV Musoke GP resulted in 100% protection of cynomolgus macaques challenged by intramuscular injection or aerosol exposure and protected against RAVV Ravn and MARV Angola [13, 256, 257] . Although MARV-specific IgG were produced, only low levels of neutralizing antibodies were detected [13, 257] . Surprisingly, T cell-mediated responses were not observed in NHPs vaccinated with recombinant VSV expressing MARV GP [13, 256] . Safety is a concern for this vaccine, especially for immunocompromised individuals, as it is a replication-competent VSV vector. However, in all VSV-based filovirus vaccine studies VSV viremia was observed only shortly after immunization. Additionally, the VSV-based filovirus GP vaccine was well tolerated and protective in immunocompromised mice and NHPs and lacked neurovirulence in NHPs [258] [259] [260] (reviewed in [255] ). |
Cross-protection has not been observed in animals vaccinated with MARV-based vaccines and subsequently challenged with EBOV, while combined MARV and EBOV vaccines have been successful in protection against both viruses [252, 261, 262] . |
To date no approved treatment is available for MARV infection. Supportive care (fluids, anti-microbials, blood transfusions) has been the primary treatment of patients during MVD outbreaks. In the guinea pig model various treatments had some success as reflected by prolonged survival or increased survival rates. Applied treatments included cytokine inhibition, IFN treatment, or antibody transfer. The tested treatments were unsuccessful in the NHP model (reviewed in [16, 47] ). |
A third of EBOV-infected NHPs survived, however, following treatment with recombinant nematode coagulant protein 2, while only one of six MARV-infected animals survived [12, 263] . Treatment using antisense technology to block viral protein expression using phosphorodiamidate morpholino oligomers (PMO) beginning 30 to 60 minutes after MARV infection completely protected NHPs [264] . Additionally, a small molecule inhibitor showed complete protection of MARV-infected mice when administered 24h after infection but has not been tested in NHPs [265] . |
The VSV-based vaccine expressing MARV GP has also been demonstrated to be effective as a post-exposure treatment. A hundred percent survival of NHPs was observed when the vaccine was administered 20 to 30 minutes after MARV infection [266] . Delaying the time before treatment results in incomplete protection, although five of six animals or two of six animals still survived when given the treatment 1 or 2 days after MARV exposure, respectively [205] . These post-exposure treatments may be useful to prevent disease after known exposure to MARV, such as a laboratory accident, but the effective time frame during an outbreak might be too short and alternatives are needed. The Human Cytomegalovirus DNA Polymerase Processivity Factor UL44 Is Modified by SUMO in a DNA-Dependent Manner During the replication of human cytomegalovirus (HCMV) genome, the viral DNA polymerase subunit UL44 plays a key role, as by binding both DNA and the polymerase catalytic subunit it confers processivity to the holoenzyme. However, several lines of evidence suggest that UL44 might have additional roles during virus life cycle. To shed light on this, we searched for cellular partners of UL44 by yeast two-hybrid screenings. Intriguingly, we discovered the interaction of UL44 with Ubc9, an enzyme involved in the covalent conjugation of SUMO (Small Ubiquitin-related MOdifier) to cellular and viral proteins. We found that UL44 can be extensively sumoylated not only in a cell-free system and in transfected cells, but also in HCMV-infected cells, in which about 50% of the protein resulted to be modified at late times post-infection, when viral genome replication is accomplished. Mass spectrometry studies revealed that UL44 possesses multiple SUMO target sites, located throughout the protein. Remarkably, we observed that binding of UL44 to DNA greatly stimulates its sumoylation both in vitro and in vivo. In addition, we showed that overexpression of SUMO alters the intranuclear distribution of UL44 in HCMV-infected cells, and enhances both virus production and DNA replication, arguing for an important role for sumoylation in HCMV life cycle and UL44 function(s). These data report for the first time the sumoylation of a viral processivity factor and show that there is a functional interplay between the HCMV UL44 protein and the cellular sumoylation system. Most replicative DNA polymerases include a catalytic subunit, responsible for DNA polymerization, and a processivity factor that holds the catalytic subunit on DNA to allow continuous DNA synthesis. One of the best-studied processivity factors is proliferating cell nuclear antigen (PCNA) of eukaryotic DNA polymerases d and e [1] . PCNA, which belongs to the family of so-called ''sliding clamps'', has no inherent DNA-binding capacity, but with the aid of clamp loader proteins is assembled onto DNA as a toroidal homotrimer [2] . In addition to DNA replication, PCNA has been implicated in DNA recombination and repair, as well as in DNA methylation, chromatin remodeling, and cell cycle regulation [1] . Consistent with its pleiotropic functions, it interacts with a plethora of proteins [3] and undergoes a number of posttranslational modifications, including phosphorylation, acetyla-tion, ubiquitination and sumoylation, which are believed to regulate its subcellular localization, stability and protein binding specificity [4, 5, 6] . |
The human cytomegalovirus (HCMV) DNA polymerase includes a catalytic subunit, UL54 (the UL54 gene product), and an accessory, homodimeric subunit, UL44 (the UL44 gene product), that binds DNA without the aid of clamp loaders [7] yet wraps around DNA akin to PCNA [8] . While UL44 shows no apparent sequence homology with PCNA, there is striking structural similarity between UL44 and PCNA monomers [2, 9] . Similarly to PCNA, UL44 is a phosphoprotein [10] . Intriguingly, the phosphorylation state of UL44 has been shown to regulate its nuclear import rate by controlling its interaction with host cell factors [11, 12, 13] . The best-characterized function of UL44 during HCMV infection is that of binding to UL54 through a region named connector loop [14, 15, 16] , stimulating its activity and conferring processivity to the holoenzyme [17, 18] . However, UL44 continues to accumulate to strikingly high levels at late times after infection, when DNA replication is accomplished [19, 20] . Its early-late kinetics of transcription and the high level of expression suggest that UL44 might play additional roles during the viral life cycle. |
To investigate this possibility, we conducted yeast two-hybrid (Y2H) screenings to search for cellular partners of UL44. To our surprise, Ubc9, an enzyme involved in the sumoylation process, was identified as a UL44 protein interaction partner. Sumoylation is a post-translational protein modification analogous to ubiquitination. It consists of reversible and covalent conjugation of SUMO (Small Ubiquitin-related MOdifier) to a protein target [21, 22] . In the sumoylation cascade, the C-terminus of SUMO is activated by an activating enzyme (E1), transferred to a conjugating enzyme (E2, that is Ubc9), and linked to a lysine residue of the substrate protein with the aid of a ligase (E3). Mainly, three SUMO paralogs (SUMO-1, -2, -3) have been identified so far [23, 24] . SUMO-2 and SUMO-3 are highly homologous to one another (95% identity) while they differ from SUMO-1 by 50%. Conjugation of SUMO-1 has been shown to play a functional role in a number of biological processes, ranging from nucleocytoplasmic transport to transcription, the maintenance of genome stability, nucleic acid DNA metabolism, cell signaling, and many others [21] , whereas the role of SUMO-2/23 modification is less clear. |
Here we report that the association of Ubc9 and UL44 leads to conjugation of SUMO molecules on multiple lysine residues. Both SUMO-1 and SUMO-2/3 were found to be conjugated to UL44. Sumoylation of UL44 was detected not only in vitro and in transiently transfected cells but, more importantly, also in HCMVinfected human cells during virus replication. Interestingly, we observed that binding of UL44 to DNA greatly stimulates SUMO conjugation to the protein both in vitro and in cells. In addition, we show that overexpression of SUMO-1 alters the intranuclear distribution of UL44 in HCMV-infected cells, and enhances both viral DNA replication and virus production in an Ubc9-dependent manner. These data represent the first report of sumoylation of a viral processivity factor and show that there is a complex interplay between the HCMV UL44 protein and the cellular sumoylation system. |
The Y2H plasmids expressing LexA-UL44 and LexA-Ubc9 were generated by cloning the UL44 and Ubc9 coding sequences from pRSET44 (a gift of P. F. Ertl, GlaxoSmithKline, UK) and pACT2-Ubc9 (from G. Gao, Chinese Academy of Sciences, Beijing, China) respectively, in pBTMK, derived from pBTM116 [25] . The pACT-UL44 and pACT2-Ubc9 plasmids, encoding GAD-UL44 and GAD-Ubc9 fusions, respectively, have been described in [26, 27] . The plasmid expressing GAD-UL54 was created by cloning the UL54 coding sequence from pRSET-Pol (a gift of P. F. Ertl) in pACT2 (Clontech). Plasmid pRSET44 was used to express 6His-UL44 in Escherichia coli. Plasmid pRSET-Ubc9 was constructed by cloning the Ubc9 coding sequence from pACT2-Ubc9 in pRSET (Invitrogen). Plasmid pCDNA3-PB1, used for in vitro transcription of the PB1 subunit of influenza A virus RNA polymerase, was described previously [28] . Plasmids pD15-GST and pD15-UL44, which express GST and GST-UL44, respectively, have been described in [29] . Plasmid pTE1E2S1 [30] was provided by H. Saitoh (Kusamoto University, Japan). Plasmids GFP-UL44, pDESTnV5-UL44, and pDESTnV5-UL44DNLS have been described previously [11, 31] . Plasmids pDsRed2-Ubc9 [27] and pDsRed2-UL53 [32] were kindly provided by G. Gao (Chinese Academy of Sciences, Beijing, China) and D. Camozzi (University of Bologna, Italy), respectively. Plasmid pCDNA3.1-UL44-FLAG (from M. Marschall, Universitat Erlangen-Numberg, Germany) was used to express C-terminally FLAG-tagged UL44 [33] , while plasmid pDEST-nFLAG [34] was used to express N-terminally FLAG-tagged UL44 [35] , UL44Dloop [36] and UL44L86A/L87A in mammalian cells. Plasmids pCDNA3-Ubc9, pCDNA3-Ubc9C93S, pCDNA3-HA-SUMO-1, pCDNA3-HA-SUMO-2, and pCDNA3-HA-SUMO-3 used in overexpression experiments in mammalian cells were a gift of R. T. Hay (University of Dundee, UK). The deletion mutants LexA-UL44 1-100 , LexA-UL44 1-200 , LexA-UL44 1-300 , LexA-UL44 1-350 , LexA-UL44 1-390 , and LexA-UL44 1-420 were generated by PCR amplification of plasmid pBTMK-UL44 with appropriate primers (Table S1 ). The LexA-UL44 114-433 , LexA-UL44 201-433 , and LexA-UL44 313-433 constructs were generated by deleting part of the UL44 coding sequence from pBTMK-UL44 with restriction enzymes. Plasmids pDESTnFLAG-UL44(1-300) and pDESTnFLAG-UL44(313-433) were generated using the Gateway Technology (see Supplementary Material and Methods in Text S1). All other UL44 mutants and the Ubc9C93S mutant were obtained by using the QuikChange mutagenesis kit (Stratagene) with primers containing appropriate nucleotide change(s). More details on plasmid construction and mutagenesis are given in Supplementary Material and Methods in Text S1. The sequences of all primers used in this work are reported in Table S1 . All DNA sequences were confirmed by sequencing. |
Y2H screenings and interaction assays. Growth media and standard methods for manipulating yeast cells were as described [37] . Saccharomyces cerevisiae strain L40 was transformed [38] with the bait plasmid pBTMK-UL44 and subsequently with either of two cDNA libraries fused to GAD (see Supplementary Material and Methods in Text S1). Primary transformants were selected for growth on -His-Leu-Trp dropout plates. His + colonies were thereafter analyzed for b-galactosidase activity by filter lift experiments [39] . Double positive clones were subjected to another cycle of screening (for further details see Supplementary Material and Methods in Text S1). cDNA inserts of interactor plasmids were sequenced and analyzed with BLAST (www.ncbi/ blastn). To quantify b-gal expression, the method of Breeden and Nasmyth [40] was used. |
Proteins. E. coli-expressed, purified GST and GST-or 6Histagged UL44 proteins were obtained as previously described [29] , with modifications (Supplementary Material and Methods in Text S1). In some preparations, samples were treated with polymin P as described [41] to eliminate residual bacterial nucleic acids. Preparation of UL44 SUMO-modified in E. coli was accomplished as described in Supplementary Material and Methods in Text S1. |
GST-pulldown assays. Assays were performed using GST and GST-UL44 and in vitro-translated UL54, PB1, or Ubc9 as previously described [29] , with modifications (Supplementary Material and Methods in Text S1). In vitro transcription-translation of proteins was performed from the appropriate plasmid by using the TNT T7 coupled transcription-translation system (Promega). The translation products were labeled with [ 35 S]methionine (Amersham Pharmacia Biotech). |
In vitro sumoylation assays. The assays to test in vitro sumoylation of UL44 by SUMO-1, -2, and -3 were performed using purified wild-type and mutant 6His-UL44 or GST-UL44 fusion proteins and the SUMOlink SUMO-1 kit from Active Motif or the SUMOylation Kit from Enzo Life Science according to the manufacturer's suggestions. In some experiments, double stranded (ds) DNA (e.g., activated calf thymus DNA from Amersham Pharmacia Biotec or salmon testes DNA from Sigma) or singlestranded (ss) DNA (e.g., single-stranded calf lung DNA, from Crinos, Como, Italy) was added to the reaction mixture at a final concentration of 500 nM. |
Human foreskin fibroblasts (HFF; from the American Type Culture Collection [ATCC]), HeLa (from ATCC), eco Phoenix (a generous gift from G. P. Nolan, Stanford, USA; [42] ), and Human Embryonic Kidney 293T (HEK 293T; from ATCC) cells were maintained in Dulbecco's modified Eagle's medium (DMEM, Life Biotechnologies) supplemented with 10% fetal calf serum (FCS), 100 U/ml penicillin and 100 mg/ml streptomycin (P/S). COS-1 cells (from ATCC) were maintained in DMEM with 5% FCS and P/S. The U373-SUMO-1 cell line, which constitutively expresses FLAG-tagged SUMO-1 [43] , and the control U373-Neo cell line, stably transfected with an empty vector carrying a Neomycin resistance marker [44] , were kindly provided by G. S. Hayward (Johns Hopkins University School of Medicine, Baltimore, USA) and were maintained in medium containing 0.5 mg/ml Neomycin (G418, Gibco-BRL). HCMV strain AD169 was purchased from ATCC. |
To analyze UL44 sumoylation in transient expression assays, HeLa or Phoenix cells were transfected for 48 h with appropriate plasmids using the calcium phosphate precipitation method. For analysis of UL44 sumoylation during HCMV replication, HFF, U373-Neo, and U373-SUMO-1 cells were mock-infected or infected with HCMV at a multiplicity of infection (MOI) of 5 PFU/cell. Cells were harvested at different time points after infection and analyzed by western blotting as described below. |
Transfected or infected cells were lysed in an appropriate volume of buffer I (5% SDS, 0.15 M Tris-HCl pH 6.8, 30% glycerol) diluted 1:3 in buffer II (25 mM Tris-HCl pH 8, 50 mM NaCl, 0.5% NP-40, 0.5% sodium deoxycholate, 0.1% SDS) supplemented with Complete protease inhibitors (Roche Molecular Biochemicals) and 5 mM N-ethylmaleimide (NEM). Lysates were then incubated on ice for 20 minutes and boiled at 95uC for 10 min. Proteins were separated by SDS-PAGE, electroblotted onto a polyvinylidene fluoride membrane (Bio-Rad), and analyzed by western blotting with indicated antibodies (details are given in Supplementary Material and Methods in Text S1). |
For immunoprecipitation analysis, lysates were diluted 1:5 in E1A buffer (50 mM Hepes pH 7.5, 250 mM NaCl, 0.1% NP-40) supplemented with Complete protease inhibitors and 5 mM NEM. Immunoprecipitation was performed with 2-3 mg of total lysate using a ratio of 3-7 mg antibody/mg of total proteins (see Supplementary Material and Methods in Text S1 for details on antibodies) and protein A-Sepharose beads. For co-immunoprecipitation analysis, cells were lysed in E1A buffer supplemented with Complete protease inhibitors and 5 mM NEM, and successively co-immunoprecipitations were performed with 1.5-5 mg of total lysate and 50 ml of 50% slurry of anti-FLAG-M2-Agarose beads (Sigma). |
Mass spectrometric identification of sumoylated lysine residues within UL44 was performed after in-gel-digestion of E. coliexpressed and SUMO-modified UL44 with endoproteinase Trypsin. Extracted peptides were analyzed by LC-MSMS on an Orbitrap Velos (ThermoFisherScientific) exactly under the conditions described in Hsiao et al. [45] . Data analysis was performed by the use of software ''ChopNSpice'' [45] in combination with MASCOT as search engine. See Supplementary Material and Methods in Text S1 for details. |
For confocal laser-scanning microscopy (CLSM) analysis, COS-1 were transfected using the Arrest-IN TM (Biosystems) reagent, according to the manufacturer's recommendations. At 24 h posttransfection, cells were fixed with 4% paraformaldehyde. Cells were imaged using a Leica TCS-SP2 confocal microscope equipped with a 636 oil immersion objective. |
For analysis of UL44 intranuclear localization in HCMVinfected U373-SUMO-1 and U373-Neo cell lines, cells were seeded at 2.5 6 10 5 /well on glass coverslips in 6-well plates and allowed to attach. The next day, cells were infected with HCMV AD169 at an MOI of 1 or of 5 PFU/cell. Cells were fixed in 4% paraformaldehyde in PBS for 15 min at room temperature, and then permeabilized with acetone for 2 min at -20uC. After washing extensively with PBS, cells were incubated first with 4% FBS in PBS for 1 h at room temperature and then with a primary mouse monoclonal antibody against UL44 (10-C50, Fitzgerald Industries International) at a dilution of 1:100 in FBS 4% in PBS for 1 h at 37uC. Cells were then washed extensively with 4% FBS in PBS and incubated with a secondary goat anti-mouse fluorescein-conjugated antibody (Ig-FITC, Chemicon International) at a dilution of 1:1000 for 1 h at 37uC. Cells were successively washed with PBS and mounted in 70% glycerol in PBS. For better visualization, cells were counterstained with Evans Blue and analyzed also for red fluorescence. CLMS analysis was then performed as described above. |
To analyze the effects of SUMO-1 overexpression on viral DNA synthesis, U373-Neo and U373-SUMO-1 cells transduced with either shUbc9 or non-silencing lentiviral particles (see below) or non-transduced, were seeded at a density of 5610 4 per well in 24well plates. The next day, cells were infected with HCMV AD169 at an MOI of 1 PFU/cell. At 72 h post-infection (p.i.), cells were collected and total DNA was extracted using the QiAmp DNA Extraction Kit (Qiagen). The levels of viral DNA were then determined by qPCR and normalized to the cellular b-globin gene copies as described [46] . |
To analyze the effects of SUMO-1 overexpression on virus production, virus yield assays were performed as described previously [47] , with some modifications. Briefly, U373-Neo and U373-SUMO-1 cells transduced with either shUbc9 or nonsilencing lentiviral particles (see below) or non-transduced, 5 610 4 cells per well were seeded in 24-well plates, incubated overnight, and infected with HCMV AD169 at an MOI of 1. At 120 h p.i., cells were subjected to one cycle of freezing and thawing, and titers were determined by transferring 100-ml aliquots from each of the wells to a fresh 96-well monolayer culture of HFF cells followed by 1:5 serial dilution across the plate. After incubation at 37uC for 7 days, cell monolayers were stained with crystal violet and plaques were counted. |
Lentiviral particles were produced by transient transfection of HEK 293T cells with packaging plasmids helper D8.9 (Addgene) and helper Ampho (kindly provided by the Tissue Culture Facility at the IEO, Milan, Italy) and either pTRIPZ-shUBC9 vector (Open Biosystems) or non-silencing pTRIPZ control vector (Open Biosystems) by the calcium phosphate precipitation method. Supernatants were collected at 48 h post-transfection and used for the transduction of U373-SUMO-1 and U373-Neo target cells in the presence of polybrene (8 mg/ml, Sigma). Selection was done in puromycin (0.5 mg/ml, Invitrogen) for two weeks prior to 4-day doxycycline (1 mg/ml, Sigma) induction to obtain Ubc9 silencing. Ubc9-knocked-down cells were screened for Red Fluorescent Protein expression and used for further experiments. |
To identify cellular proteins that interact with UL44, Y2H screens were carried out with a bait consisting of full-length UL44 protein (amino acids 1-433) fused to the E. coli LexA protein. |
Control experiments demonstrated that the LexA-UL44 protein did not activate expression of either HIS3 or lacZ reporter gene by itself and, as expected [9, 14] , could both interact with UL54 and dimerize (Table 1 and data not shown). Thus, this bait was used to screen two different cellular cDNA libraries fused to S. cerevisiae GAL4 activation domain (GAD), one derived from human B lymphocytes [48] and the other from promyelocytic HL-60 cells [49] . A total of 167 or 85 colonies, respectively, were positive for both HIS3 and lacZ reporter genes. Plasmids encoding putative interactors of UL44 were isolated from double-positive clones and retransformed into yeasts expressing LexA-UL44 in order to confirm the interaction. The 28 and 13 positive clones after this retransformation, respectively, were sequenced. |
In total, from the two screenings we identified 7 cellular UL44binding proteins; here we report the identification of human Ubc9 as a specific interaction partner of UL44. In particular, 15 out of 28 clones of B lymphocytes library and 6 out of 13 clones of the HL-60 cells library contained the whole Ubc9 coding sequence, plus 59 and 39 untranslated regions which varied in length in different clones. By co-transformation experiments with each individual interactor clone expressing Ubc9 and the LexA vector, it was excluded that Ubc9 could activate the reporter genes in the absence of the bait protein (Table 1) . Ubc9 specifically interacted with UL44 in Y2H assays also in the reverse combination, i.e., with Ubc9 fused to LexA and UL44 fused to GAD (Table 1 ). In addition, in quantitative assays the b-galactosidase activity of yeasts expressing both UL44 and Ubc9 turned out to be comparable to that of yeasts expressing UL44 and UL54 (Table 1) , which served as a positive control. |
Having identified Ubc9 as a potential interaction partner of UL44 in Y2H screens, we wished to confirm their physical interaction by an independent experimental approach. For this purpose, pulldown assays with a purified GST-UL44 protein and in vitro-translated, 35 S-labeled Ubc9 were performed. As positive and negative controls, we also assayed the interaction between UL44 and in vitro-translated UL54 or the PB1 subunit of influenza A virus RNA polymerase. As expected, we could detect the interaction of UL54 [14, 29] , but not of PB1, with GST-UL44 (Fig. 1A) . Consistent with the Y2H results, Ubc9 specifically associated with GST-UL44, while no interaction with GST was observed (Fig. 1A) . |
Since our data suggested that UL44 can interact with Ubc9 both in vitro and in yeast cells, we sought to examine whether UL44 and Ubc9 could also co-localize in mammalian cells. To this end, aggregates distributed on a diffuse background fluorescence throughout the nucleus (Fig. 1B) . As previously reported [50] , when expressed alone Ubc9 exhibited a nuclear punctate pattern (Fig. 1B) . Upon co-expression of GFP-UL44 and DsRed2-Ubc9, co-localization of UL44 we analyzed the subcellular localization of GFP-UL44 when transiently expressed either alone or in the presence of DsRed2-Ubc9. As a negative control, we also expressed the DsRed2-UL53 fusion protein, which localizes to the cell nucleus but does not interact with UL44 [32] . In GFP-UL44-transfected cells, UL44 localized in a large number of discrete nuclear with Ubc9 was observed (Fig. 1B) . UL44 also colocalized with a catalytically impaired Ubc9 mutant (Ubc9C93S) [51] , but not with UL53 [32] . In addition, co-immunoprecipitation experiments confirmed that UL44 could physically interact with endogenous Ubc9 in mammalian cells. The specificity of the interaction was confirmed by the inability of UL44 to coimmunoprecipitate with Cyclin D1 (CycD1; Fig. 1C ). |
To further explore the interaction between UL44 and Ubc9, we sought to map the domain of UL44 that interacts with Ubc9. To this aim, several N-and C-terminal deletion mutants of UL44 fused to LexA were generated and tested for the ability to interact with GAD-Ubc9 by Y2H assays. Control western blot experiments with an anti-LexA antibody evidenced protein bands of the expected molecular mass for all mutants (data not shown). As shown in Fig. 1D , the truncated protein UL44 1-300 , lacking most of the C-terminal disordered region of UL44, exhibited interaction with Ubc9. Further C-terminal truncation of UL44 revealed that a protein fragment corresponding to the first 200 amino acids of UL44 (UL44 ) was still capable of interacting with Ubc9 ( Fig. 1D ). In contrast, the N-terminal 100 residues of UL44 (UL44 1-100 ) exhibited no interaction with Ubc9 (Fig. 1D ). Although unfolding of the UL44 1-100 mutant protein cannot be excluded, these results suggested that this region of UL44 may not contain sequences important and/or sufficient for Ubc9 binding. Therefore, we analyzed the effects of N-terminal truncations. Deletion of the N-terminal 113 residues of UL44 (UL44 114-433 ) did not impair the ability of UL44 to bind Ubc9. Similarly, the UL44 201-433 mutant, lacking the N-terminal 200 amino acids, interacted with Ubc9. Interestingly, a mutant that only expresses the C-terminal 121 residues of UL44 (UL44 313-433 ) still retained the ability to bind Ubc9 (Fig. 1D) . Control Y2H experiments showed that none of the truncated UL44 proteins was able to activate transcription by itself (data not shown). Mapping studies in mammalian cells expressing UL44 deletion mutants showed that both the UL44 1-300 and UL44 313-433 mutants could immunoprecipitate endogenous Ubc9, similarly to wild-type UL44 (Fig. S1 ). Thus, our results suggest that UL44 contains two domains capable of independently binding to Ubc9, located at the N-and Cterminus of the protein (likely within residues 1-200 and 313-433, respectively). |
The observation that UL44 interacts with the SUMO-conjugating enzyme Ubc9 prompted us to investigate the possibility that UL44 may be sumoylated. This hypothesis was first tested in a cellfree system by incubating a purified 6His-UL44 fusion protein with purified Aos1/Uba2 (E1), Ubc9 (E2), and SUMO-1. As a control, sumoylation of human p53 was also examined by the same assay. As expected [52, 53] , p53 was readily modified to give mainly a single mono-sumoylated product that reacted with both anti-p53 ( Fig. 2A , left bottom panel) and anti-SUMO-1 ( Fig. 2A , right bottom panel) antibodies (the bands appearing at high molecular weight, which are particularly visible in the anti-SUMO-1 panel, correspond to SUMO-E1 and -E2 enzymes conjugates). In a western blot analysis with an anti-UL44 antibody ( Fig. 2A , left top panel), three main slower migrating forms of UL44 were observed (lane 3). The appearance of the ,65, 80, and 95-kDa forms of UL44 was strictly dependent on the presence of SUMO-1, as the substitution of wild-type SUMO-1 with a mutant form which bears the Gly97-to-Ala change (SUMO-1 mut) and hence cannot be attached to target proteins, eliminated formation of these products ( Fig. 2A, left top panel, lane 4 ). In addition, SUMO-1 modification was abolished if either SUMO-1, Aos1/ Uba2, or Ubc9 was omitted from the reaction (not shown). A western blot analysis with an anti-SUMO-1 antibody confirmed that the slower migrating UL44 bands contained SUMO-1 ( Fig. 2A, right top panel) . Taken together, the above results established that UL44 is a substrate for in vitro SUMO-1 conjugation. |
Having shown that UL44 can be modified by SUMO-1, we wondered whether it could be conjugated also to SUMO-2 and SUMO-3. Thus, the in vitro sumoylation system was applied to purified 6His-UL44 in the presence of sumoylation enzyme components and activated forms of SUMO-2 or SUMO-3. In these assays, the RanGTPase-activating protein RanGAP1 [54] was used as a positive control and as previously observed [55] , was modified in the presence of SUMO-2 and SUMO-3 (data not shown). As shown in Fig. 2B , UL44 was also modified by either of the two SUMO peptides. |
To verify whether UL44 could also be sumoylated in mammalian cells, we expressed UL44-FLAG and HA-SUMO-1 fusion proteins, in the presence of either wild-type Ubc9 or the catalytically impaired mutant Ubc9C93S in Phoenix cells and analyzed cell lysates by western blotting with anti-FLAG, anti-HA, and anti-Ubc9 antibodies. As expected, several bands corresponding to SUMO-1-conjugated proteins that reacted with an anti-HA antibody were detected upon co-expression of wild-type Ubc9, while less SUMO-1 conjugation was observed in the presence of the Ubc9C93S mutant (Fig. 3A) . This was not due to differences in expression of the two Ubc9 variants, as evidenced by western blot analysis with the anti-Ubc9 antibody (Fig. 3A) . When UL44-FLAG was expressed in the absence of HA-SUMO-1 and Ubc9, a single band with an apparent molecular mass of ,50 kDa was detected using the anti-FLAG antibody. In contrast, slower migrating bands similar to those observed in in vitro sumoylated products ( Fig. 2A) were observed upon co-expression of UL44-FLAG with HA-SUMO-1 and wild-type Ubc9 (Fig. 3A) . These bands were significantly reduced in the presence of the Ubc9C93S mutant (Fig. 3A) , demonstrating that they were dependent on the catalytic activity of Ubc9, with the residual slower migrating bands due to the activity of wild-type endogenous Ubc9. |
To confirm that the observed products were indeed sumoylated forms of UL44, we expressed UL44-FLAG either in the absence or in the presence of HA-SUMO-1 and Ubc9, immunoprecipitated UL44-FLAG using an anti-FLAG antibody and analyzed the immunoprecipitated proteins by western blot. As expected, three slower migrating bands were detected by the anti-FLAG antibody from lysates of cells expressing UL44-FLAG in the presence of both HA-SUMO-1 and Ubc9 (Fig. 3B, left panel) . The ,50-kDa non-sumoylated form of UL44 could be immunoprecipitated both from cells expressing UL44-FLAG alone and from cells expressing UL44-FLAG together with HA-SUMO-1 and Ubc9, but not from Table 1 . UL44 interacts with human Ubc9 in yeast two-hybrid assays. LexA-UL44 / 2(,1) |
UL44, UL54, and Ubc9 proteins were fused to the C-terminus of LexA protein and/or of GAL4 activation domain (GAD). Fusion proteins were then assayed for interaction by qualitative b-galactosidase (b-gal) filter assays and by quantitative b-gal liquid assays. Ubc9, but not in immunoprecipitates obtained from cells expressing only UL44-FLAG (Fig. 3B, right panel) . Similar results were obtained in HeLa cells (Fig. S2) . Altogether, these results demonstrate that Ubc9 can mediate the conjugation of SUMO-1 to UL44 in mammalian cells. Moreover, sumoylation of UL44 in mammalian cells by both SUMO-2 and SUMO-3 could also be detected (Fig. S3) . |
Since sumoylation mainly occurs in the nucleus of mammalian cells [56] and UL44 is translocated to the host cell nucleus during HCMV infection [11] , we decided to investigate whether nuclear localization might be a prerequisite for conjugation of SUMO-1 to UL44. We therefore analyzed the ability of UL44bDNLS, a derivative of UL44 bearing point mutations within the nuclear localization signal (NLS, 425 PNTKKQK 431 ) which prevent UL44 nuclear accumulation [11] , to be modified by SUMO-1. Interestingly, mutations of UL44 NLS impaired the sumoylation of a V5-UL44 fusion protein (V5-UL44DNLS, Fig. 3C ), suggesting that SUMO-1 modification of UL44 most likely occurs into the nucleus or during nuclear import. |
We next sought to identify the SUMO-1 acceptor sites of UL44. A prediction analysis with the SUMOplot program (Abgent) identified seven residues in UL44 with a certain probability to be sumoylated: K73, K172, K224, K339, K371, K410, and K431. To test whether one or more of these lysines could be a SUMO-1 acceptor site, each of the candidate residues was conservatively mutated to arginine, both individually and in combination, and the mutant proteins were tested for in vitro sumoylation. None of the single point mutants exhibited a consistently altered SUMO-1 modification pattern as compared to wild-type UL44 (Fig. S4A ). Mutants carrying two or three K/R substitutions also showed a SUMO-1 modification pattern identical to that of wild-type UL44 ( Fig. S4A and data not shown) . Similar results were obtained when in vitro sumoylation reactions with SUMO-2/23 were performed (data not shown). We decided to also test the ability of these K/R mutants to be modified by SUMO-1 in mammalian cells. Consistent with the in vitro data, none of the tested mutations significantly affected the ability of UL44 to undergo SUMO-1 conjugation ( Fig. S4B and data not shown) . |
These results suggested that UL44 might possess multiple lysine residues that could alternatively serve as SUMO-1 acceptors, as reported for other proteins [57, 58] , and/or that UL44 might be sumoylated on lysine residues other than those predicted by SUMOplot. UL44 contains 31 lysines, most of which are solventexposed in the crystal structure [9] and therefore potentially accessible to SUMO molecules, which makes it difficult to identify the target lysines by mutational approaches. Therefore, we attempted to map the sites of UL44 where SUMO-1 is conjugated by mass spectrometry analysis. To this end, we expressed UL44 in an E. coli expression/modification system that produces SUMOconjugated proteins [30] . The 6His-tagged UL44 construct was co-expressed in E. coli with the pTE1E2S1 plasmid, which contains a linear fusion of genes for E1 and E2 enzymes and SUMO-1 under the control of an IPTG-inducible promoter [30] . To confirm sumoylation of UL44 with this system, UL44 was purified from the bacterial cultures expressing UL44 alone or in combination with the SUMO conjugation system and analyzed by western blotting with both the anti-UL44 (Fig. S5, left panel) and the anti-SUMO-1 (Fig. S5, right panel) antibody. Shifted bands with an apparent molecular mass of ,65, 80, and 95 kDa, similar to those detected in in vitro reactions ( Fig. 2A) and in cells (Fig. 3) , were observed in UL44 purified from bacteria cotransformed with pTE1E2S1, but not in UL44 purified from bacteria expressing only UL44 (Fig. S5) . Then, to identify lysine acceptor sites by mass spectrometry, sumoylated UL44 was separated by SDS-PAGE, in-gel-digested with trypsin and peptides were analyzed by LC-MSMS. Upon application of the software ''ChopNSpice'' for database search, we identified 16 sumoylation sites in UL44, including the predicted sites K172, K339, K371, K410, and K431. Table 2 summarizes the UL44 peptides that have been found to be sumoylated. The corresponding MS spectra are shown in Fig. S6 . |
These results indicated that UL44 possesses multiple SUMO target lysines that are located throughout the protein, in accordance with the observation that Ubc9 could bind to both N-and C-terminal portions of UL44 ( Fig. 1D and Fig. S1 ). Mutagenesis of several of these lysines in combination caused a strong decrease of UL44 expression (data not shown), likely due to protein misfolding and/or instability, making impossible to analyze the sumoylation state of the mutated protein and to compare it to that of wild-type UL44. |
UL44 sumoylation is stimulated by binding to DNA. As mentioned above, UL44 possesses a structural fold similar to that of the eukaryotic processivity factor PCNA [9] . In addition, like UL44, PCNA is SUMO-conjugated and its sumoylation involves both a consensus and a non-consensus site [4] . Since it has been shown that PCNA needs DNA to be sumoylated efficiently [59] , we wished to investigate whether UL44 might behave similarly and its sumoylation could be stimulated by the presence of DNA. Thus, we performed in vitro sumoylation experiments in the absence and in the presence of DNA using as a substrate a purified 6His-UL44 fusion protein treated with polymin P to eliminate residual bacterial nucleic acids. In a western blot analysis with an anti-UL44 antibody (Fig. 4A, left panel) , only a faint band corresponding to mono-sumoylated UL44 was observed in the absence of DNA (lane 2). Upon addition of dsDNA (e.g., activated calf thymus DNA) to the reaction mixture, a marked increase of the mono-sumoylated product and the appearance of bi-and trisumoylated forms of UL44 were observed (Fig. 4A , lane 4 of left panel). A western blot analysis with an anti-SUMO-1 antibody confirmed that these bands indeed contained SUMO-1 (data not shown). Similar results were obtained when GST-UL44 was used as a substrate (Fig. S7) . Thus, like PCNA sumoylation, UL44 sumoylation is strongly stimulated by the presence of DNA. |
To investigate whether the nature of DNA could influence the stimulation of UL44 sumoylation, we also performed in vitro sumoylation reactions in the presence of different DNA substrates. Similar stimulation levels were obtained when different dsDNAs were added, regardless of their sequence (data not shown), in keeping with previous observations that UL44 binds DNA in a sequence-independent manner [7] . In contrast, consistently less stimulation of UL44 sumoylation was observed in the presence of ssDNA (Fig. 4A, right panel) , for which UL44 has been shown to possess an apparent affinity lower than for dsDNA [7] . This suggested that UL44 sumoylation could depend on binding to To analyze UL44 sumoylation in vitro, purified 6His-UL44 was incubated in the absence or the presence of sumoylation enzymes and either wild-type SUMO-1 (SUMO-1 wt) or a mutant form of SUMO-1 (SUMO-1 mut) which cannot be covalently linked to substrates. The reaction products were analyzed by western blotting with anti-UL44 and anti-SUMO-1 antibodies. As a positive control, in vitro sumoylation of p53 was also analyzed. (B) Purified 6His-UL44 was incubated in the absence or the presence of sumoylation enzymes and either SUMO-2 or SUMO-3 and analyzed by western blotting with anti-UL44 and anti-SUMO-2/23 antibodies. For all panels, the arrowhead indicates the unmodified form of UL44 or p53 and the asterisks indicate the respective sumoylated forms. doi:10.1371/journal.pone.0049630.g002 DNA. To test this hypothesis, we analyzed the ability of two UL44 mutants, FLAG-UL44Dloop and FLAG-UL44L86A/L87A, which are defective for DNA binding [36] , to be modified by SUMO-1 in mammalian cells. FLAG-UL44Dloop contains three point mutations within a UL44 flexible loop ( 163 HTRVKRNVKKAP 174 ) involved in UL44-DNA interaction [8, 9] , and is therefore impaired in DNA binding [36] . FLAG-UL44L86A/L87A carries two point mutations preventing dimerization of UL44 and strongly impairing the UL44-DNA interaction both in vitro and in vivo [9, 36, 60] . Both FLAG-UL44Dloop (Fig. 4B) and FLAG-UL44L86A/L87A (Fig. 4C) , when coexpressed with HA-SUMO-1 and Ubc9, exhibited strongly reduced sumoylation levels when compared to FLAG-UL44. Importantly, co-immunoprecipitation experiments demonstrated that the reduced sumoylation of FLAG-UL44Dloop and FLAG-UL44L86A/L87A was not due to an impairment of binding of the mutant proteins to Ubc9, since the two mutants precipitated with endogenous Ubc9 at levels comparable to those of the wild-type protein (Fig. 4D ). In addition, the presence of DNA did not stimulate the sumoylation of the UL44Dloop or UL44L86A/L87A mutants in in vitro reactions (Fig. 4E) . Finally, since the UL44Dloop mutant contains a substitution (K167N) involving a potential SUMO target lysine, we wished to exclude the possibility that the reduced sumoylation levels of FLAG-UL44Dloop might be due to alteration of a putative SUMO acceptor site rather than an impairment of DNA binding. To this end, the K167 residue was conservatively mutated to arginine and the mutant protein was tested for in vitro sumoylation in the presence of DNA. The K167R mutant showed a SUMO-1 modification pattern identical to that of wild-type UL44 (Fig. 4F) . |
Altogether, these results suggest that UL44 is preferentially modified by SUMO-1 when it is bound to DNA as a dimer. |
Having demonstrated that UL44 is sumoylated by Ubc9 in vitro and in transfected cells, we sought to investigate whether a similar modification also occurs naturally in HCMV-infected cells. Protein lysates of HFFs infected with HCMV and collected at different times post-infection (p.i.) were analyzed by western blotting with an anti-UL44 antibody. Two main bands of 65 and 80 kDa were observed above the primary UL44 band of 50 kDa during the whole time course of lytic infection, being detectable already at 24 h p.i. (Fig. 5A) . A third band of ,95 kDa was also visible from 48 h p.i. These subforms were similar in electrophoretic mobility to the UL44 bands covalently modified by SUMO-1 in transfected cells (Fig. 3A) . A densitometric analysis of the protein bands (Fig. 5B ) revealed that the relative amount of the sumoylated forms increased during the course of HCMV infection, becoming ,50% of total UL44 protein at 120 h p.i. |
To confirm that the slower migrating forms indeed represent UL44 molecules conjugated to SUMO-1, lysates from HCMVinfected cells at 120 h p.i. or from mock-infected cells were immunoprecipitated with an antibody against UL44. Subsequent- ly, the immunocomplexes were analyzed by western blotting with an anti-SUMO-1 antibody. Three main bands of the expected molecular mass (,65, 80, and 95 kDa) were recognized in the anti-UL44 immunoprecipitate (Fig. 5C, left panel) . Finally, the same immunocomplexes were analyzed by western blotting with the anti-UL44 antibody to demonstrate that the SUMO-1 crossreactive proteins were indeed modified UL44 forms (Fig. 5C, right panel) . Thus, these results clearly indicate that UL44 is covalently modified by SUMO-1 in HCMV-infected cells. |
Considering the difficulties in expressing a UL44 mutant completely impaired in sumoylation, whose activities could be compared to that of wild-type UL44, to gain some insight on the functional role of UL44 sumoylation in the context of HCMV replication we sought to undertake a different approach. We overexpressed SUMO-1 in virus-infected cells and analyzed the effects on the intracellular distribution of UL44, as the targeting to specific subcellular domains is one of most common biological effect exerted by the conjugation of SUMO to a substrate protein. |
It has been previously shown that during HCMV replication UL44 localizes to large globular intranuclear structures that correspond to viral DNA replication compartments [61, 62, 63] . A U373-MG cell line that constitutively overexpresses FLAG-SUMO-1 and control U373-Neo cells were mock-infected or infected with HCMV at an MOI of 1 or of 5 PFU/cell and the intracellular localization of UL44 was successively analyzed by indirect immunofluorescence with an anti-UL44 antibody. Control western blotting experiments (Fig. S8A ) confirmed that UL44 is sumoylated in the HCMV-infected U373 cells. In fact, slower migrating bands of the expected molecular mass and similar to the UL44 sumoylated forms observed in infected HFFs (Fig. 5A) were detected. Furthermore, as expected, they increased upon SUMO-1 overexpression. In immunofluorescence assays, the nuclei of mock-infected cells were oval-shaped with no anti-UL44 staining (Fig. 6A, upper panels) . Control HCMV-infected U373-Neo cells showed deformed nuclei, many of which exhibited a kidney shape (Fig. 6A, upper panels) . Indeed, it has been observed that infection by HCMV causes this kind of distortions in nuclear shape [64, 65] . Moreover, UL44 showed a globular fluorescent pattern consistent with previously described viral replication compartments in HCMV-infected cells [61, 66] . In contrast, in HCMV-infected U373-SUMO-1 cells UL44 staining was more distributed throughout the nucleus and, especially at the lower MOI (MOI 1), also failed to coalesce into any recognizable globular structures (Fig. 6A, upper panels) . Therefore, overexpression of SUMO-1 during HCMV replication appears to alter the intranuclear distribution of UL44, likely leading to significantly decreased localization of UL44 in viral DNA replication compartments. Importantly, the intranuclear distribution of another HCMV protein localizing to the replication compartments, i.e. UL57, the single-stranded DNA (ssDNA)-binding protein [61, 67] , appeared not to change upon SUMO-1 overexpression (Fig. 6A, lower panels) . |
To investigate whether sumoylation is indeed involved in the altered intranuclear distribution of UL44, an RNAi approach was employed to suppress the sumoylation system of the cells by silencing Ubc9 since Ubc9 is the only unique and essential enzyme in the SUMO-conjugating pathway [21] . U373-Neo and U373-SUMO-1 cells were transduced with a lentivirus expressing shUbc9, followed by selection with puromycin and induction with doxycyline to establish Ubc9-knocked-down cell lines (U373-Neo shUbc9 and U373-SUMO-1 shUbc9, respectively). As a control, U373-SUMO-1 cells were also transduced with a lentivirus expressing a non-silencing shRNA sequence (U373-SUMO-1 NS). As shown in Figure 6B , Ubc9 was almost completely silenced in cells infected with the shUbc9 lentivirus (U373-SUMO-1 shUbc9) as compared to the cells transduced with the control lentivirus (U373-SUMO-1 NS) and to nontransduced cells (U373-Neo and U373-SUMO-1). To examine the effect of Ubc9 knock-down on intranuclear distribution of UL44, these cell lines were infected with HCMV. As shown in Fig. 6A (upper panels) , upon doxycycline induction, the nuclear UL44 staining in Ubc9-knocked-down U373-SUMO-1 cells was similar to that of U373 cells not overexpressing SUMO-1 (U373-Neo and U373-Neo shUbc9; Fig. 6A , upper panels, and Fig. S8B ). In contrast, the U373-SUMO-1 cells transduced with the non-silencing lentivirus (U373-SUMO-1 NS, Fig. 6A , upper panels) retained an altered intranuclear distribution of UL44 similar to that observed in the non-transduced cells (U373-SUMO-1) or in the shUbc9-transduced U373-SUMO-1 cells with no doxycycline induction (data not shown). Altogether, these results established that the altered intranuclear distribution of UL44 in HCMV-infected cells upon SUMO-1 overexpression depends on Ubc9-mediated sumoylation. |
These observations raised the question whether the altered intranuclear distribution of UL44 observed upon SUMO-1 overexpression affected the viral replication efficiency. To examine the effects of SUMO-1 overexpression on viral DNA replication, U373-Neo and U373-SUMO-1 cells were infected with HCMV at an MOI of 1 and viral DNA levels were measured by quantitative real-time PCR. Viral DNA production from U373-SUMO-1 cells was ,two-fold higher than that from the control cells (U373-Neo and U373-Neo shUbc9; Fig. 6C ). This increase was not observed in Ubc9-knocked down cells (U373-SUMO-1 shUbc9), while the U373-SUMO-1 cells transduced with the non-silencing lentivirus (U373-SUMO-1 NS) exhibited augmented viral DNA levels like the non-transduced U373-SUMO-1 cells (Fig. 6C) . |
We also examined the effects of SUMO-1 overexpression on virus yield. The titers of viral particles produced from nontransduced U373-SUMO-1 and from transduced U373-SUMO-1 shUbc9 and U373-SUMO-1 NS cells after infection with HCMV at an MOI of 1 were determined and compared to those produced from infected U373-Neo and U373-Neo shUbc9 control cells. A 2-3-fold increase in viral progeny titers was observed in U373-SUMO-1 and U373-SUMO-1 NS with respect to U373-Neo, while the U373-SUMO-1 shUbc9 cells exhibited yields of Ubc9, and anti-vinculin antibodies (left panel). Cell lysates were incubated with anti-FLAG-M2-Agarose beads and the immunoprecipitated samples were analyzed by western blotting with anti-UL44 and anti-Ubc9 antibodies (right panel). (E) The sumoylation in vitro of wild-type 6His-UL44 and mutant 6His-UL44Dloop and 6His-UL44L86A/L87A proteins was carried out as in (A) and analyzed by western blotting with an anti-UL44 antibody. (F) The sumoylation in vitro of a UL44 mutant bearing the K167R substitution in the flexible loop of UL44 involved in DNA binding was carried out in the presence of DNA and compared to that of wild-type UL44. For all panels, the arrowhead indicates the unmodified form of UL44 or free SUMO-1 and the asterisks indicate the sumoylated forms. doi:10.1371/journal.pone.0049630.g004 infectious virus similar to those of the U373-Neo and U373-Neo shUbc9 cells (Fig. 6D) . |
Thus, the altered intranuclear distribution of UL44 upon SUMO-1 overexpression appears not to compromise HCMV replication, but conversely, SUMO-1 overexpression causes a positive effect on virus production. |
In this study we report that UL44, a viral ortholog of PCNA, is sumoylated on multiple lysines by the cellular factor Ubc9. Importantly, a consistent portion of UL44 is SUMO-modified in HCMV-infected human cells, resulting in ,50% of the protein being modified at late times during virus replication. From a structural point and functional of view, UL44 and PCNA share blotting with anti-SUMO-1 (left panel) and anti-UL44 (right panel) antibodies. For all panels, the arrowhead indicates the unmodified form of UL44, the arrow indicates the immunoglobulin G heavy chain (IgG hc) and the asterisks indicate the sumoylated UL44 forms. doi:10.1371/journal.pone.0049630.g005 some remarkable similarities and some differences. Monomers of UL44 and PCNA are structurally very similar, despite having extremely different primary sequences [2, 9] . However, while PCNA forms toroidal-homotrimers, UL44 binds to dsDNA as a head-to-head homodimer [7, 9] . In addition, PCNA must be loaded onto DNA in an ATP-dependent process by so-called clamp loaders [68] ; in contrast, UL44 directly binds DNA without the need for ATP hydrolysis or accessory proteins [7, 14, 18] . |
Similarities and differences also emerge from the comparison of the sumoylation processes of UL44 and PCNA. The most striking similarity is the DNA-dependence of such post-translational modification: in the case of PCNA, clamp loading rather than the mere presence of DNA was shown to be important for stimulation, implying a change in the properties of PCNA upon loading that enhances its capacity to be sumoylated [59] . This could also be the case for UL44: in fact, point mutations preventing DNA binding [8, 9, 36, 60] strongly impaired UL44 sumoylation in cells (Fig. 4C) and abolished the ability of dsDNA to stimulate UL44 sumoylation in vitro (Fig. 4E) . Furthermore, the ability of DNA to stimulate UL44 sumoylation appears to correlate with the efficiency of the UL44-DNA interaction. In fact, addition of dsDNA -for which UL44 has a ,3-to 8-fold higher affinity than for ssDNA [7] -caused a much stronger increase of UL44 sumoylation as compared to ssDNA (Fig. 4A) . In this context, it is important to note that point mutations impairing DNA binding and sumoylation in cells did not compromise the UL44-Ubc9 interaction (Fig. 4D) , suggesting that binding to DNA does not promote UL44 sumoylation by facilitating its binding to Ubc9. |
Another similarity between UL44 and PCNA sumoylation is that both proteins are modified on canonical (K127 for PCNA, and K410 for UL44) and non-canonical residues (K164 for PCNA, and all other sumoylation sites for UL44, see Table 2 ). However, while PCNA is exclusively sumoylated at the Nterminus, both N-and C-terminal residues of UL44 can be modified. In addition, according to our MS-analysis UL44 can be alternatively sumoylated at 16 different sites, while K127 and K164 appear to be the only target sites in PCNA. This flexibility of UL44 in terms of sumoylation target sites, which is reminiscent of the ones described for Daxx and the small hepatitis delta antigen [57, 58] , is arguably the main difference from PCNA. This makes it extremely difficult to study the physiological importance of UL44 sumoylation, also because mutation of several of these lysines caused protein instability. Currently it is therefore impossible to test if, like in the case of budding yeast PCNA, SUMOmodification of UL44 also plays a role in DNA repair/ recombination [69] . |
In terms of functional effects, sumoylation is known to regulate protein activity and/or intracellular location [21, 22] . As for the latter, the targeting to specific subcellular domains is one of the best-characterized biological effects exerted by the conjugation of SUMO to a substrate protein. This effect is exemplified by the targeting of cellular protein RanGAP1 to the cytosolic side of the nuclear pore complex upon sumoylation [54, 70] . As a first, preliminary attempt to characterize the role of UL44's sumoylation, here we show that overexpression of SUMO-1 in the context of HCMV replication alters the intranuclear distribution of UL44 as it appears to result in a more diffuse pattern and in decreased localization of UL44 in viral DNA replication compartments (Fig. 6A) , suggesting that sumoylation of UL44 may retarget the protein to other nuclear site(s). Importantly, Ubc9 knock-down studies confirmed that sumoylation is responsible for such altered intranuclear pattern of UL44. The observation that SUMO-1 overexpression causes a positive effect on HCMV replication suggests that sumoylation of UL44 could be important for its function(s) in the context of the virus life cycle, although effects on HCMV replication mediated by sumoylation of other viral or cellular proteins cannot be excluded. However, at the moment understanding the molecular details of how SUMO alters the intranuclear distribution of UL44 is rather difficult. Most likely, sumoylation does not affect the functions of UL44 as a DNA polymerase processivity factor -the only role currently well established for UL44. In fact, several reports have shown that the UL44 protein expressed in E. coli, which is non-sumoylated, is capable of performing all known biochemical activities related to this role (e.g., [7, 14] ). In addition, our observation that UL44 sumoylation peaks at late times during virus replication, once the viral DNA replication has been accomplished (Fig. 5) , suggests that SUMO-conjugation might enable UL44 to fulfill some role(s) in HCMV replication other than that of conferring processivity to the viral DNA polymerase. A role of UL44 in late gene expression has been previously suggested [71, 72] ; however, no conclusive demonstration has been provided yet. |
Intriguingly, higher molecular mass forms of the Epstein-Barr virus DNA polymerase processivity factor BMRF1 compatible with sumoylated products have been observed [73] , suggesting that such post-translational modification could be a general feature of the DNA polymerase accessory subunit in herpesviruses. In addition, sumoylation of Vaccinia Virus G8R protein has been recently predicted on the basis of structural similarities to PCNA [74] , but not yet experimentally demonstrated. Thus, our findings could stimulate further studies on sumoylation of DNA polymerase subunits in other herpesviruses or, more in general, in other viral systems. Figure S5 Sumoylation of UL44 in E. coli. The pRSET44 plasmid, encoding 6His-tagged UL44, was introduced into E. coli together with the pTE1E2S1 plasmid, which expresses E1 and E2 sumoylation enzymes and SUMO-1. As a control, bacteria were also transformed only with pRSET44. The 6His-tagged UL44 was purified from bacterial cultures expressing UL44 alone or in combination with the SUMO conjugation system and analyzed by western blotting with anti-UL44 and anti-SUMO-1 antibodies. (TIF) Figure S6 Mass spectrometry analysis of sumoylation sites of UL44. MSMS analysis of tryptic peptides conjugated to a tryptic peptide of SUMO-1 (ELGMEEEDVIEVYQEQTGG or IADNHTPKELGMEEEDVIEVYQEQTGG, 1 tryptic miscleavage) derived from E. coli-expressed, sumoylated UL44. Sequence of the conjugate is listed in each table above the spectra. Y-type and b-type fragment ions are assigned in the spectra. The table below each spectrum summarizes the database search. Highlighted are the m/z values that match the fragment ions obtained from in silico fragmentation of the conjugate. Conjugated lysine residues to SUMO-1 are highlighted in red and the position in UL44 is listed. Modifications, measured (observed) m/z values, the actual mass (in Da), the charge state, and the mass error (ppm, parts per million) are listed as well in the table above each spectrum. The different colors represent the various measured and calculated fragment ions of each conjugate in the spectrum and table underneath, respectively. The question marks in some of the spectra indicate fragment ions that do not match to any calculated y-and b-type ions of the conjugate. (PDF) Figure S7 Sumoylation in vitro of a GST-UL44 fusion is stimulated by DNA. E. coli-expressed, purified GST-tagged UL44 was incubated with purified sumoylation enzymes in the absence or presence of SUMO-1 and/or DNA. The samples were analyzed by western blotting with an anti-UL44 antibody. (TIF) Figure S8 UL44 is sumoylated in HCMV-infected U373 cells. (A) U373-Neo and U373-SUMO-1 cells were either mockinfected or infected with HCMV at MOI of 5 PFU/cell for 72 h. Cell lysates were then analyzed by western blotting with an anti-UL44 antibody. The arrowhead indicates the unmodified form of UL44 and the asterisks indicate the sumoylated UL44 forms. (B) Control U373-Neo cells, and U373-Neo cells transduced with lentiviral particles expressing either a Ubc9-silencing shRNA (U373-Neo shUbc9) or a non-silencing shRNA sequence (U373-Neo NS) were mock-infected or infected with HCMV at an MOI of 5 or 1 PFU/cell. At 72 h p.i., cells were fixed and stained with a primary antibody against UL44 and successively with a secondary fluorescein-conjugated antibody (green) which contained Evans Blue to counterstain cells (red). Cell samples were then analyzed by CLSM. |
(TIF) Fas-deficient mice have impaired alveolar neutrophil recruitment and decreased expression of anti-KC autoantibody:KC complexes in a model of acute lung injury BACKGROUND: Exposure to mechanical ventilation enhances lung injury in response to various stimuli, such as bacterial endotoxin (LPS). The Fas/FasL system is a receptor ligand system that has dual pro-apoptotic and pro-inflammatory functions and has been implicated in the pathogenesis of lung injury. In this study we test the hypothesis that a functioning Fas/FasL system is required for the development of lung injury in mechanically ventilated mice. METHODS: C57BL/6 (B6) and Fas-deficient lpr mice were exposed to either intra-tracheal PBS followed by spontaneous breathing or intra-tracheal LPS followed by four hours mechanical ventilation with tidal volumes of 10 mL/kg, respiratory rate of 150 breaths per minute, inspired oxygen 0.21 and positive end expiratory pressure (PEEP) of 3 cm of water. RESULTS: Compared with the B6 mice, the lpr mice showed attenuation of the neutrophilic response as measured by decreased numbers of BAL neutrophils and lung myeloperoxidase activity. Interestingly, the B6 and lpr mice had similar concentrations of pro-inflammatory cytokines, including CXCL1 (KC), and similar measurements of permeability and apoptosis. However, the B6 mice showed greater deposition of anti-KC:KC immune complexes in the lungs, as compared with the lpr mice. CONCLUSIONS: We conclude that a functioning Fas/FasL system is required for full neutrophilic response to LPS in mechanically ventilated mice. Acute lung injury (ALI) and its more severe form, the acute respiratory distress syndrome (ARDS) remain important clinical problems in the United States, with an incidence rate of 38.3 cases per 100,000 person-years and a mortality rate of 45% [1] . ALI/ARDS is characterized clinically by sudden respiratory failure with impaired oxygenation and non-cardiogenic pulmonary edema [2] . Pathologically ALI/ARDS is associated with an early inflammatory phase with neutrophilic alveolitis and destruction of the alveolar/capillary permeability barrier, followed by a late fibroproliferative phase with abnormal repair and collagen deposition. There are no specific treatments for ARDS and the main supportive treatment, mechanical ventilation, can be harmful to the lungs when delivered at high tidal volumes [3] . |
A growing body of experimental evidence suggests that in addition to the injury caused by high tidal volumes, even moderate or low tidal volumes markedly enhance injury when the lungs are exposed to pathogens or their products [4] [5] [6] . For example, mechanical ventilation synergistically enhances lung injury in response to low doses of bacterial lipopolysaccharide (LPS) and this is associated with expression of specific sets of genes that aren't expressed with LPS or ventilation alone [4, 5, 7] . A computational analysis has mapped the biological processes that are activated by the combination of mechanical ventilation and LPS [8] . One of these biological processes is apoptosis, and a known mediator of apoptosis in injured lungs is the Fas/FasL system. |
The Fas/FasL system is composed of the membrane surface receptor Fas and its cognate ligand, FasL. FasL exists in a membrane-bound form, and also in a soluble form that is present in the lungs during acute lung injury [9] . Binding of Fas to sFasL in alveolar epithelial cells independently activates apoptotic and inflammatory pathways that result in death of the cells but also in release of pro-inflammatory cytokines [10] . Although the Fas/ FasL system is better known for its pro-apoptotic function, the pro-inflammatory function is also important in the development of lung injury, and mice deficient in Fas (lpr) have an impaired neutrophil recruitment in response to LPS installation and bacterial infections [11] . |
Several lines of evidence suggest that the Fas/FasL system plays a role in ALI/ARDS. First, bioactive sFasL is present in the lungs of patients with ARDS, and this is associated with increased mortality [9, 12] . Second, genetic variations in the Fas gene are associated with increased risk for ALI/ARDS in humans [13] . Third, activation of the Fas/FasL system in mammals leads to acute inflammatory injury followed by fibrosis [14] [15] [16] [17] . And finally, mice lacking Fas are protected in a number of models of lung injury [11, [18] [19] [20] . Thus, the apoptotic and inflammatory responses induced by Fas activation in the lungs are one important factor in the development of ALI/ARDS. |
One of the most important neutrophil chemoattractants in mice is the chemokine KC (CXCL1), which is the murine functional homologue of the human chemokine IL8 (CXCL8). Activation of Fas in alveolar epithelial cells is followed by a marked increase in KC expression [10] . Interestingly, recent studies suggest that in injured lungs, IL8 in humans and KC in mice form immune complexes with autoantibodies, and these immune complexes can in turn bind Fc receptors such as FcγRIIa and FcγRIII that amplify the inflammatory response [21, 22] . These studies suggest that, in addition to the absolute levels of chemokines, the formation of autoimmune complexes and their association with Fc receptors is important for the development of inflammatory responses in the lungs. |
In this study, we ask whether the Fas/FasL system plays a role in the amplification of inflammatory responses that occur early in the course of mechanical ventilation. The specific hypothesis to be tested is that the Fas/FasL system is required for the development of lung injury in mechanically ventilated mice exposed to LPS. To test this hypothesis, we investigate whether the combination of a non-injurious mechanical ventilation strategy with a minimal dose of intratracheal LPS results in an acute lung injury, and whether this injury is attenuated in Fas-deficient lpr mice. We further investigate whether the development of injury is associated with formation and deposition of anti-KC:KC immune complexes. |
All of the animal experiments were approved by the institutional animal research committee of the University of Washington. Mice were housed in a pathogen-free environment according to University of Washington animal use guidelines. Male C57BL/6 mice ("B6") and mice carrying a spontaneous mutation in the Fas gene that impairs Fas signaling (B6. MRL-Fas lpr /J, "lpr") were obtained from the Jackson Laboratories and studied at 7-13 weeks of age. Briefly, the mice were anesthetized with 5% inhaled isoflurane and then intubated endotracheally with a 20-gauge angiocatheter. Placement of the catheter in the trachea was verified by visualizing the movement of a 100 μl bubble of water in response to respiratory efforts. After confirming intubation, the trachea was instilled with either E. coli LPS, 15 ng/kg or PBS. The instillate was suspended in 2.5% colloidal carbon to allow later confirmation of the extent and distribution of the instillation macro and microscopically. After the installations some of the mice were extubated, returned to their cages, and allowed free access to food and water; other mice where kept intubated and subjected to mechanical ventilation with the following settings: tidal volume (Tv) 10 ml/kg; respiratory rate (RR) 150 breaths/minute; fraction of inspired oxygen (FiO 2 ) 0.21; and positive end-expiratory pressure (PEEP) of 3 cm H 2 O. The heart rate, airway pressures, rectal temperatures and EtCO 2 were monitored continuously using a computerized monitoring system (Chart 4, AD Instruments, Colorado Springs, CO). The RR was adjusted to maintain the EtCO 2 between 30 -40 mmHg. The body temperature was maintained between 37 and 38°C with external heating. The mice were hydrated with a continuous intraperitoneal infusion of lactated ringer solution at 500 μl/hour. Muscle relaxation was attained with pancuronium bromide, 1 μg/g i.p followed by 0.5 μg/g i.p. every hour. After four hours of mechanical ventilation the mice were euthanized with 0.30 ml/kg i.p. of Beuthanasia-D (Schering-Plough Animal Health, Union, NJ). The thorax was rapidly opened and the mouse was exsanguinated by direct cardiac puncture. The left lung was removed and flash-frozen in liquid nitrogen. The right lung was lavaged with 0.6 mM EDTA in PBS; an aliquot of the bronchoalveolar lavage fluid (BALF) was removed for cell counts and differentials, the remaining fluid was spun at 1200 x g, and the supernatants stored at -80°C in individual aliquots. Following the BAL, the right lung was fixed in 4% buffered paraformaldehyde at an inflation pressure of 15 cm H 2 O for histological evaluation. |
We studied four groups of mice. Two groups consisted of B6 and lpr mice instilled with PBS and allowed to breathe spontaneously ("SB") (n = 7 for B6 mice, 4 for lpr). The other two groups consisted of B6 and lpr mice instilled with LPS and exposed mechanical ventilation (MV + LPS) (n = 10 for B6 mice, 6 for lpr). The main experimental comparison was between the B6 and lpr mice instilled with LPS and exposed to ventilation. |
Total cell counts in the BALF were performed with a hemacytometer. Differential counts were performed on cytospin preparations using the Diff-quick method (Fisher Scientific Company L.L.C., Kalamazoo, MI). BALF total protein was measured with the bicinchoninic acid method (BCA assay, Pierce, Rockford, IL). BALF IgM (Bethyl Laboratories, Montgomery, TX) and α-macroglobulin (Life Diagnostics, West Chester, PA) were measured with immunoassays. Lung homogenate TNF-α, KC, IL1β, and IL6 were measured using a multiplex fluorescent bead assay (R&D Systems, Minneapolis, MN). |
As a measurement of the total content of PMN in the lungs we measured myeloperoxidase (MPO) activity in lung homogenates prepared in 50 mM K 2 HPO 4 , pH 6.0 with 5% CH 3 (CH 2 ) 15 N(Br)(CH 3 ) 3 , 5 mM EDTA. Active caspase-3 and Poly ADP ribose polymerase (PARP) activity were measured in lung homogenates prepared on a 1:20 ratio of a lysis buffer (0.5% Triton-X-100, 150 mM NaCl, 15 mM Tris, 1 mM CaCl, and 1 mM MgCl, pH7.4). The lung homogenate was spun at 10,000 x g for 20 min at 4°C and the supernatant used for measurements of active caspase-3 and PARP using the CPP32/ Caspase-3-Fluorometric Protease Assay Kit (BioVision, Mountain View, CA) and a PARP activity kit (Cell Signaling, Boston MA). Serum Creatinine, ALT and bilirubin were measured at a commercial laboratory. Anti-KC autoantibody:KC immune complexes were measured in BAL fluids using an ELISA assay according to a previously described protocol [21] . Briefly, 96-well microtiter plates were coated with antibody against KC (Peprotech). After blocking, the plates were incubated with BAL fluid samples obtained from mice. Then, the plates were washed and incubated with biotinylated horse antibody against mouse immunoglobulins (Vector Laboratories) followed by HRP-conjugated streptavidin and the substrate tetramethyl benzidine (Sigma). |
C57BL/6 and lpr mice were euthanized by exposure to CO 2 followed by cervical dislocation. The femur and tibia of both hind legs were isolated and freed of all soft tissue, and then the ends of both bones were removed. The femur and tibia were placed proximal end down in a 0.6 mL Eppendorf tube, which had been punctured at its lower tip with an 18-gauge needle and placed inside a 1.5 mL Eppendorf tube. The tubes were spun at 2000 X g for 30 seconds and neutrophils were isolated as previously described [23] . After isolation, neutrophils were labeled with calcein-AM (5 μg/ml; Molecular Probes, Eugene, OR) for 30 minutes at 37°C, washed two times in PBS and resuspended at a concentration of 1 x 10 6 /mL. |
Neutrophil chemotaxis was assessed using the Neuro Probe ChemoTx W Disposable Chemotaxis system (Neuro Probe Inc. Gaithersburg, MD). The wells of the 96-well plate were filled with various concentrations of KC. A polycarbonate filter (8 μm pores) with a hydrophobic ring around the area over each well was placed on the 96-well plate and calcein-labelled neutrophils were added to each ring. The chemotaxis chamber, consisting of the polycarbonate filter and 96-well plate, was incubated for 30 min at 37°C in 5% CO 2 and then non-migrating neutrophils were removed from the upper side of the filter. The chemotaxis chamber was placed in a multi-well fluorescent plate reader (Synergy 4, BioTek, Winooski, VT) and the migrated cells were measured using the calcein fluorescence signal (excitation -485 nm, emission -530 nm). Neutrophil migration was expressed as a percent of the total number of neutrophils that were placed on the topside of the filter (% Total). |
Lung sections were embedded in paraffin, cut into 4 μm sections, and stained with hematoxylin and eosin. |
Lung tissue sections were deparaffinized, washed in xylene, rehydrated, permeabilized with proteinase K, and incubated with the TUNEL reaction mixture according to manufacturer instructions (In Situ Cell Death Detection kit, AP, Roche Applied Science, Indianapolis, IN). Negative controls were treated with labeling solution without terminal transferase. Immediately after TUNEL labeling the slides were washed three times in PBS, blocked with Protein Block (Dako, Carpinteria CA) and incubated for 2 hr in the dark at 37°C with the mouse monoclonal pan-cytokeratin antibody C11 (Abcam, Cambridge, UK) previously labeled with Alexa Fluor 555 (Invitrogen, Eugene, OR). Negative control slides were incubated with an isotype control mouse IgG1k labeled with Alexa Fluor 555 (BD Pharmingen, San Diego, CA). The slides were washed and immediately visualized with a fluorescent microscope. |
Lung tissue sections from B6 and lpr mice exposed to MV + LPS were processed as previously described [21] . The sections were incubated with anti-KC antibody (Peprotech, Rocky Hill, NJ) followed by chicken antirabbit secondary antibody (Alexa 647, pseudocolor red) (Invitrogen), and then with anti-FcγRIII antibody (R&D Systems) followed by chicken anti-rat secondary antibody (Alexa 488, green), and finally with biotynylated anti-Ly-6 G antibody (eBioscience, San Diego, CA), used as a neutrophil marker, followed by streptavidin (Alexa 568, pseudocolor magenta). Lung tissues were counterstained with Hoechst 33342 (Calbiochem, Gibbstown, NJ). The slides were evaluated using a PerkinElmer Ultra VIEW LCI confocal imaging system with Nikon TE2000-S fluorescence microscope using PlanApox20 objective and PlanApox60 or x100 immersion oil objective (numerical aperture [NA] 1.4) at room temperature. Ultra VIEW Imaging Suite software (version 5.5.0.4) was used for image processing. |
Statistical analysis was performed using two-factor ANOVA followed by the Bonferroni post-hoc analysis. One factor was "treatment" (SB or MV + LPS) and the other factor was "strain" (B6 or lpr). The analysis was designed to determine the overall effect of each of the factors, the presence of an interaction effect, and the comparison of "strain" for each level of "treatment". Data was generated using GraphPad Prism. A p value of less than 0.05 was considered significant. |
The neutrophil response to MV + LPS was attenuated in the lpr mice There were significantly less PMN in the BAL of lpr mice exposed to LPS + MV than in the B6 mice (2.2 x 0.5 x 10 3 vs 1.3 ± 0.2 x 10 4 cells, p < 0.05) (Figure 1-A) . A similar pattern was seen for lung MPO activity (Figure 1-B) . However, the lung concentrations of the PMN chemoattractants KC and MIP2 (CXCL2), although increased in response to MV + LPS, were similar in the lpr and B6 mice; this was also true for the pro-inflammatory cytokine TNFα (Figure 1 , C-F). Interleukin 6, GM-CSF, and VEGF were not affected by the administration of MV + LPS and were similar in the B6 and lpr mice (data not shown). These data suggest that both lung PMN recruitment and airspace PMN migration were impaired in the absence of functional Fas, but the difference in neutrophil recruitment seen in this model cannot be explained by differences in KC or MIP-2 release. |
Other parameters of lung injury were similar in B6 and lpr mice Permeability response |
We and others have postulated that the Fas/FasL system leads to lung injury by inducing apoptosis of pneumocytes, resulting in disruption of the alveolar-epithelial barrier and non-cardiogenic pulmonary edema. However, in the present study the BAL concentration of total protein and of the serum protein α-macroglobulin were similar in the lpr and B6 mice, despite the difference in PMN numbers ( Figure 2 ). The lack of difference may be due to a low level of injury, as supported by the lack of overall change in the MV + LPS group as compared with the SB group; indeed the model was designed so that the ventilatory pattern and the LPS dose would cause minimal or no injury by themselves, so as to determine the very first components of the injury response. The data suggest that neutrophil changes precede permeability changes in ventilated mice exposed to LPS, and that the role of the Fas/FasL system on neutrophil recruitment precedes its disruptive effect on the alveolar-capillary barrier. |
Interestingly, the activity of caspase-3, often used as a surrogate for apoptosis, was increased in the spontaneously breathing lpr mice, and this increase reached significance in the MV + LPS group (185 ± 24 vs 480 ± 137, p < 0.05) (Figure 3-A) . PARP, a downstream target of active caspase-3, showed similar activity in the lpr and B6 mice (Figure 3-B) . Double labeling of tissue sections for TUNEL and cytokeratin revealed that in both the lpr and the B6 mice, the apoptotic cells were located primarily in the alveolar walls, but were cytokeratin-negative (Figure 3-C) . Thus, contrary to our expectations, there was increased caspase-3 activity in the lpr mice exposed to MV + LPS, and the apoptotic cells were mostly cytokeratin-negative cells localized to the alveolar walls. |
The lungs of spontaneously breathing B6 and lpr mice showed normal architecture (Figure 4 ). The lungs of B6 mice exposed to MV + LPS showed thickening of the alveolar walls, intra-alveolar neutrophilic infiltrates, and intra-alveolar fibrin strand deposition. In contrast, the lungs of lpr mice exposed to MV + LPS retained their normal architecture. The LPS was administered with inert colloidal carbon, which on the tissue sections was taken up by macrophages and appears as black granulate material in the cytoplasm. This indicates that the Figure 1 Inflammatory response. C57BL/6 (B6) and Fas-deficient lpr mice received intratracheal installations of either PBS followed by four hours of spontaneous breathing (SB), or E. coli LPS, 15 ng/kg, followed by 4 hrs of mechanical ventilation (MV) with tidal volumes of 10 mL per kilogram, FiO 2 of 0.21, PEEP = 3 and respiratory rate = 150 breaths per minute. In response to the combination of MV and LPS, the B6 mice showed significantly more total neutrophils (PMN) in the BAL fluid than the lpr mice (A). A similar pattern was seen for the lung MPO activity, which is a measure of the total neutrophil content in the lung (B). MV + LPS was associated with increases in the lung homogenate concentrations of the cytokinesTNF-α, MIP-2, and KC, and these similar in the B6 and the lpr mice; MV + LPS had no effect on IL-1β (C-F). n = at least 6/group. sections depicted were all exposed to instillate -an important point as intratracheal instillations are patchy and normal tissue can simply reflect a non-instilled area. |
We have previously reported that the combination of MV + LPS for 6 hours (2 hr longer than the present study) is associated with distal organ injury, in particular biochemical and histologic evidence of kidney and liver damage [24] . In this study, we also noticed an increase in serum creatinine in the mice exposed to MV + LPS, but this increase was strain-independent ( Figure 5-A) . Interestingly, the lpr mice had increased serum AST concentrations at baseline, and this was not further increased by the addition of MV + LPS (Figure 5-B) . Total bilirubin was similar in lpr and B6 mice, and was not affected by MV + LPS (Figure 5-C) . Thus, at this early time the evidence of distal organ injury was limited to increased creatinine, and this increase was not dependent on the Fas/FasL system. |
The differences in BAL neutrophils are not explained by differences in B6 and lpr neutrophil chemotaxis An impairment in lpr neutrophil chemotaxis towards KC could explain the attenuation of BAL neutrophils seen in the lpr mice exposed to MV + LPS. Therefore, we compared the chemotactic ability of neutrophils isolated from B6 and lpr mice to KC in vitro. However, we noticed no impairment of chemotaxis in the lpr neutrophils; if anything, chemotaxis was slightly increased in the lpr neutrophils as compared with the B6 neutrophils ( Figure 6 ). |
A new line of research suggests that mammals form autoantibodies against chemokines such as IL8 in humans and KC in mice, and immune complexes derived from these autoantibodies can induce inflammation by binding to Fcγ receptors on leukocytes, endothelial cells and other cells. Measurement of anti-KC:KC complexes in the BALF revealed an increase in B6 mice treated with MV + LPS, which was not seen in the lpr mice (Figure 7) We also used immunofluorescence staining and confocal microscopy to investigate if there were differences in the deposition of anti-KC:KC complexes in ventilated B6 and lpr mice. The alveolar walls of the ventilated B6 mice contained abundant leukocytes, identified based on Gr-1 positivity ( Figure 8 ). The leukocytes expressed the receptor FcγRIII on their membrane surface. KC co-localized with the FcγRIII receptors. In contrast, there were, in general, Caspase-3 activity was significantly higher in the lpr mice exposed to MV + LPS. n = at least 6/group. Double-labeling for TUNEL (green) and cytokeratin (red) reveals that the TUNEL positive cells are located in the alveolar wall, but most of them are cytokeratin negative. |
less Gr-1 positive cells in the lungs of the lpr mice, and there was also decreased deposition of KC in the tissue. Thus, the formation of anti-KC:KC IC was attenuated in lpr mice, and this was associated with decreased KC deposition in the tissues. |
To confirm the specificity of the anti-KC:KC labeling, mouse PMN were purified and incubated with either anti-KC-KC immune complexes (Figure 9 , first two columns) or with KC alone (Figure 9 , third column). Then, the cells were labelled with either the Peprotech antibody, which detected the immune complexes, or an R&D antibody, which detected only KC. |
Finally, we tested expression of FcγRIII in WT and lpr mice treated with MV + LPS, and found it to be lower in the lpr mice ( Figure 10 ). |
The goal of this study was to determine whether Fas deficiency is protective in mechanically ventilated mice exposed to intratracheal LPS. The main finding was that Fas-deficient lpr mice showed decreased neutrophil recruitment in response to combined mechanical ventilation and LPS, even though the cytokine, permeability, and apoptotic responses were similar to those of B6 mice. Interestingly, despite the presence of similar concentrations of KC in the BAL of lpr and B6 mice, only the B6 mice showed extensive KC deposition in the lungs, and this was associated with the presence of anti-KC:KC complexes in the BALF and lung tissue. |
In this study, we focused on the initial events of the injury response by studying mice four hours after the initiation of injury, which was induced by combining a very low dose of endotoxin with mechanical ventilation. The main finding was that a functional Fas/FasL system was required for neutrophil migration into the airspaces of the lungs of mechanically ventilated mice exposed to LPS. The changes in BAL neutrophils seen in this study were associated with changes in lung myeloperoxidase (MPO) activity, and because lung MPO activity is a measurement of the total content of neutrophils in the lungs, the data suggests that the Fas/FasL system is required for both neutrophil migration into the airspaces and for neutrophil recruitment into the lungs. This observation confirms other studies that suggest a key role for the Fas/FasL system in neutrophil migration into the airspaces; for example, activation of Fas in the lungs of mice and rabbits results in a neutrophilic alveolitis (1, 2) , whereas pharmacologic blockade of the Fas/FasL system attenuates the neutrophilic response to bacteria and bacterial products (3) (4) (5) (6) . Tissue respons. C57BL/6 or Fas-deficient lpr mice were exposed to either intratracheal installations of PBS followed by spontaneous breathing or to intratracheal installation of LPS, 15 ng/kg, followed by four hours of mechanical ventilation (MV) with tidal volumes of 10 mL per kilogram. Lung tissue sections were stained with hematoxylin and eosin to demonstrate the pattern of injury. The instillate contained 2.5% colloidal carbon to identify the instilled areas. Spontaneously breathing mice instilled with PBS showed normal lung architecture, regardless of strain (A, C). The B6 mice exposed to MV and LPS showed intra-alveolar and interstitial neutrophilic infiltrates, as well as thickening of the alveolar walls and occasional fibrin deposition (B). |
One potential explanation for the decreased migration of neutrophils into the airspaces of the Fas-deficient mice is that Fas ligation induces release of neutrophilic cytokines such as KC in macrophages and in lung epithelial cells in vitro (7-9). Thus, we had expected to see lower concentrations of KC in the mechanically ventilated lpr mice exposed to LPS, as compared with the B6 animals. However, this was not the case, and the difference in neutrophil migration was not due to differences in soluble KC concentrations. |
Another potential mechanism that would explain differences in neutrophil migration with similar concentrations of soluble KC is a difference in neutrophil chemotaxis of lpr and B6 neutrophils. However, our chemotaxis studied showed that, if anything, neutrophils from lpr mice have slightly increased chemotaxis to KC, strongly suggesting that differences in chemotaxis are not the reason for the attenuation in the neutrophilic response seen in the lpr mice. Neutrophil recruitment and migration appears to be partly dependent on the formation of anti-KC:KC immune complexes, which can bind Fcγ receptors in local leukocytes, enhancing the inflammatory response [22] . For example, the generation of anti-KC:KC immune complexes in the lungs of mice is followed by acute inflammatory lung injury, and this injury requires the presence of Fcγ receptors [21] . Furthermore, mice lacking γ chains show attenuated injury in response to LPS, suggesting that this process is relevant for inflammation secondary to bacterial products [21] . The human equivalent of anti-KC:KC complexes are anti-IL8:IL8 complexes, and these are present in the lungs of patients with ARDS [25, 26] . In our study, we found that there are less anti-KC:KC complexes in the lungs of the lpr mice, even though the concentrations of KC were similar to those in the B6 mice. Thus, it is possible that the lower numbers of neutrophils seen in the lpr mice were due to decreased formation and deposition of anti-KC:KC complexes in the lungs. Additional studies are necessary to confirm this hypothesis, and it remains possible that differences in other unmeasured chemotactic agents account for the differences in neutrophil recruitment. The mechanism linking the Fas/FasL system with impaired formation and deposition of immune complexes remains unclear. To date, lpr mice, particularly the MRL/ lpr strain are known to generate autoantibodies that can result in a lupus-like syndrome [27] . |
A surprising finding in the present study was that there was no increase in the markers of permeability or apoptosis in the B6 mice exposed to mechanical ventilation and LPS, as compared to the lpr mice; instead, the injury response was limited to neutrophilic inflammation and cytokine release. One explanation is that neutrophil recruitment precedes the development of tissue injury in our model, in which the mice were studied relatively early, four hours after the onset of ventilation. Less clear is the finding that the lpr mice actually had increased caspase-3 activity and TUNEL (See figure on previous page.) Figure 8 Anti-KC autoantibody:KC complex deposition in the lungs. FcγRIII (green) and KC (red) in tissue sections from B6 (A) and lpr (B) mice treated with intratracheal E. coli LPS, 15 ng/kg, and exposed to 4 hours of mechanical ventilation (MV) at Tv = 10 cc/kg, FiO2 = 0.21, PEEP = 3. FcγRIII is shown in green, KC is shown in red, and Gr1 (staining leukocytes) is shown in pink. Nuclei are shown in blue. There is an increase signal for both KC and FcγRIII in the B6 mice as compared with the lpr mice. Note the colocalization of KC and FcγRIII. Figure 9 The detection of anti-KC:KC immune complex is specific. Mouse PMN were purified and incubated with either anti-KC-KC immune complexes (first two columns) or with KC alone (third column). Note that the Peprotech antibody, detected only anti-KC-KC immune complexes, whereas the R&D antibody detected only KC. The FcγRIII was labeled green and KC was labeled red. |
positive cells compared with the B6 mice. We do not have a clear explanation for this finding, but they seem to be specific to the LPS + MV model, because in another model using mechanical ventilation in which WT and lpr mice were exposed to pneumonia virus of mice (PVM) prior to four hours of MV, we found a decrease in caspase-3 activity in the lpr mice [28] . |
It is important to emphasize that the decrease in neutrophils seen in the lpr mice does not imply "protection". It is unclear whether the lung neutrophilic response is directly associated with negative outcomes in ALI/ARDS or not. Most studies of lung injury presume that neutrophilia or increased concentrations of cytokine are deleterious, but all of these studies are limited in that they do not effectively reproduce the series of events seen in the clinical setting, where patients with multiple comorbidities are intubated for prolonged periods of time. Our study should be interpreted as knowledge on the mechanisms of that initiate lung injury, but not extrapolated to the ultimate results of that injury. |
In summary, we report that in B6 mice, the combination of mechanical ventilation and LPS is associated with recruitment of neutrophils to the lungs and with deposition of anti-KC:KC immune complexes. In comparison, mice deficient in Fas recruited lower numbers of neutrophils, and this was associated with less deposition of immune complexes in the lungs. We conclude that a functioning Fas/FasL system is required for a full neutrophilic response to LPS in mechanically ventilated mice. Evolutionary Dynamics of the Interferon-Induced Transmembrane Gene Family in Vertebrates Vertebrate interferon-induced transmembrane (IFITM) genes have been demonstrated to have extensive and diverse functions, playing important roles in the evolution of vertebrates. Despite observance of their functionality, the evolutionary dynamics of this gene family are complex and currently unknown. Here, we performed detailed evolutionary analyses to unravel the evolutionary history of the vertebrate IFITM family. A total of 174 IFITM orthologous genes and 112 pseudogenes were identified from 27 vertebrate genome sequences. The vertebrate IFITM family can be divided into immunity-related IFITM (IR-IFITM), IFITM5 and IFITM10 sub-families in phylogeny, implying origins from three different progenitors. In general, vertebrate IFITM genes are located in two loci, one containing the IFITM10 gene, and the other locus containing IFITM5 and various numbers of IR-IFITM genes. Conservation of evolutionary synteny was observed in these IFITM genes. Significant functional divergence was detected among the three IFITM sub-families. No gene duplication or positive selection was found in IFITM5 sub-family, implying the functional conservation of IFITM5 in vertebrate evolution, which is involved in bone formation. No IFITM5 locus was identified in the marmoset genome, suggesting a potential association with the tiny size of this monkey. The IFITM10 sub-family was divided into two groups: aquatic and terrestrial types. Functional divergence was detected between the two groups, and five IFITM10-like genes from frog were dispersed into the two groups. Both gene duplication and positive selection were observed in aquatic vertebrate IFITM10-like genes, indicating that IFITM10 might be associated with the adaptation to aquatic environments. A large number of lineage- and species-specific gene duplications were observed in IR-IFITM sub-family and positive selection was detected in IR-IFITM of primates and rodents. Because primates have experienced a long history of viral infection, such rapid expansion and positive selection suggests that the evolution of primate IR-IFITM genes is associated with broad-spectrum antiviral activity. First discovered by cDNA library screening in 1984 [1] , the interferon-induced transmembrane (IFITM) gene family plays critical roles in a variety of cellular processes and contains IFITM1, IFITM2, IFITM3, IFITM5, IFITM6, IFITM7, IFITM10 and some IFITM-like genes [2] . Except for IFITM5 that is specifically expressed in bone cells in an interferon (IFN)independent way [3, 4] , all IFITM genes can be stimulated by IFN [5, 6] , and are widely expressed in tissues and organs [2] . |
IFITM family members contain a conservative CD225 domain and two terminal hypervariable regions [2] . The CD225 domain accounts for more than half of the protein in length, containing one intact transmembrane domain (TMD), two S-palmitoylation sites regions and partial TMD in the C-terminus of the protein. The S-palmitoylation sites have been demonstrated to play important roles in post-translational processing and stability of IFITM proteins [7] . The N-terminal hypervariable region generally contains 21 amino acid residues and the C-terminal one includes a TMD (Fig. 1) [8] . |
In different vertebrates, the functions of different IFITM members diverge. IFITM1, IFITM2 and IFITM3 are involved in cell adhesion [9] , antiproliferation [9] , tumor suppression [10, 11] , and germ cell and embryonic development [12] . More recently, these genes were identified as novel types of antiviral restriction factors with a wide spectrum of antiviral activity against influenza A viruses (e.g. H1N1 viruses), West Nile virus, dengue virus, filoviruses, HIV-1, HCV, venezuelan equine encephalitis virus (VEEV), chikungunya virus (CHIKV), vesicular stomatitis virus (VSV) and even SARS-coronavirus [8, [13] [14] [15] [16] [17] [18] . The main function of IFITM5 is associated with bone development in vertebrates [4, 19, 20] . IFITM6 seems to be involved in macrophage functions in tumor suppression [20] . To date, however, there is no information about the functions of IFITM7 and IFITM10. |
Several antiviral restriction factors (e.g. APOBEC3G, Tetherin, and SAMHD1) have been demonstrated to evolve under positive selective pressure from viruses [21] [22] [23] [24] [25] [26] [27] [28] [29] . As important virus inhibitors, IFITM1, IFITM2 and IFITM3 may have also undergone a similar co-evolutionary process, such as other antiviral restriction factors do. Despite this connection, relationships between antiviral functions and adaptive evolution in IFITM family have seldom been reported and although previous reports had illustrated the phylogenetic history of IFITM family in some eukaryotic species [2, 30, 31] , there has been no detailed information about IFITM genes in vertebrates. In this study, we performed detailed evolutionary analyses not only to test whether the primate IFITM genes evolved under positive selection throughout primate evolution, but also to unravel the evolutionary history of vertebrate IFITM family. |
To characterize the IFITM gene repertoires in vertebrates, we searched 27 vertebrate genome sequences with high genome coverage ($66) or representing the major evolutionary lineages in vertebrate phylogeny (such as opossum, lizard, platypus, etc.), using previously described IFITM sequences as queries. The taxa included ten non-mammalian vertebrates (five fishes: stickleback, tetraodon, medaka, fugu, zebrafish; one amphibian: frog; one nonavian reptile: lizard; three birds: chicken, turkey, zebra finch) and 17 mammals covering six primates (human, chimpanzee, gorilla, orangutan, macaque, marmoset), four glires (mouse, guinea pig, rat, and rabbit), five other mammals (tree shrew, cow, horse, dog and elephant), 1 metatherian (opossum) and 1 prototherian (platypus). We divided the newly identified genes into three types based on the following criteria: (i) functional genes, which contain full-length ORFs with intact CD225 domain and C-terminal TMDs; (ii) putative functional genes, which have CD225 domains but contain incomplete ORFs; and (iii) pseudogenes, which are sequences with pre-mature stop codons. |
We identified a total of 286 IFITM-related sequences ( Table 1 ). The number of IFITM genes varies considerably between mammals and non-mammalian vertebrates. A total of 27 functional genes were identified from 10 non-mammalian species, ranging from one gene in stickleback or tetraodon to six genes in frog. By contrast, 134 functional genes were identified from 17 mammalian species, ranging from one in platypus or rabbit to 26 in marmoset. Additionally, 14 putative functional genes including 13 in mammals and one in non-mammalian vertebrates were identified. Among the 10 non-mammalian and 17 mammalian species, 10 and 102 pseudogenes were identified, respectively. Interestingly, among 10 non-mammalian vertebrates, frog has the highest number (six functional genes and eight pseudogenes) of IFITM-related sequences. Among the 17 mammals, marmoset has the highest number (40) of IFITM-related sequences, of which 26 are functional genes. Human has the second highest number (29) of IFITM-related sequences, of which 18 are pseudogenes, the maximum in vertebrates. |
To understand the phylogenetic relationship of IFITM gene family, 160 functional IFITM genes identified from 27 vertebrate species (Table S1 ) were subjected to phylogenetic analyses using Bayesian inference, maximum likelihood (ML) and maximum parsimony (MP) methods. The Bayesian, ML and MP trees show consistent topological structures (Figs. 2A and S1). All functional IFITM genes were well divided into three clades I, II and III with Bayesian posterior probabilities of $97. These analyses indicate that the IFITM family most likely originated from three progenitors. |
The clades I, II and III contain 114, 26, and 20 IFITM genes, respectively. All IFITM10 and IFITM5 genes are clustered in clades II and III, respectively. All other functional IFITM genes, including IFITM1, IFITM2, IFITM3, IFITM6 and IFITM7, are grouped in clade I, forming the biggest sub-family in the IFITM family. Because the expression of the IFITM genes in clade I can be induced by IFN and their functions are associated with immunity [8, 13, 15] , they are defined as immunity-related IFITM (IR-IFITM) sub-family. |
In each clade, eutherian IFITM genes form a separate group from opossum and bird genes, consistent with the species phylogeny. Besides IFITM5 and IFITM10, IR-IFITM genes also have orthologs in both marsupials and eutherians, arguing against previous observation that only IFITM5 and IFITM10 orthologs could be identified in marsupials and eutherians [30] . In addition, clade II and III contain homologous IFITM sequences from teleosts and amphibians, but clade I does not ( Fig. 2A) , indicating that IR-IFITM originated later than IFITM5 and IFITM10. |
Phylogenetic Analyses of the Vertebrate Immunityrelated IFITM Sub-family IR-IFITM genes can be divided into two groups: one consisting of eutherian homologs and the other including homologs from metatheria and bird. We further constructed a sub-tree to show the phylogenetic relationship of 109 IR-IFITM genes from eutheria (Fig. 3) . The sequences from elephant are located on the basal position. All IR-IFITM genes from the primate lineages form a sub-clade, and those from rodents form another sub-clade. Four genes from tree shrew form a species-specific cluster located between the sub-clades of the primates and the rodents. Three mammal species, dog, horse and cow, form several species-specific IR-IFITM gene clusters, which further compose a sub-clade in accordance with the phylogeny of these three species. These suggest that IR-IFITM genes evolved via gene duplication after species separation. |
Interestingly, the majority of IR-IFITM genes from rodents do not form species-specific clusters as each of IFITM1, IFITM2 and IFITM3 clusters together in a lineage-specific manner. This clustering indicates that IR-IFITM genes have diverged into different IFITM isoforms prior to the split of rodents from other mammals. Additionally, in the lineage-specific clusters, more than one IR-IFITM gene was observed from certain species, indicating that gene duplication of IR-IFITM genes continued until after species separation of the rodents. Furthermore, rat IFITM7 clusters closely with rat IFITM3, indicating that they are a pair of duplicated genes. Similarly, mouseIFITM7, mouseIFITM1 and mouseIFITM-like3 (IFITMac) are another group of duplicated genes, suggesting that IFITM7 might have similar biological function to IFITM3 or IFITM1. |
Subsets and Splits