All experiments with the SS2 were performed in biosafety cabinets in biosecurity level 3 laboratory. All animal studies were conducted in strict accordance with the animal welfare guidelines of the World Organization for Animal Health. The animal study protocol was approved by the Ethics Committee of Huazhong Agricultural University (Wuhan, China) and conducted in accordance with the Hubei Province Laboratory Animal Management Regulations of 2005. All efforts were made to minimize animal suffering.

The bacterial strains and plasmids used in this study are listed in Table 1. The virulent SS2 strain SC19 was isolated from a diseased pig during the 2005 epidemic outbreak in Sichuan of China (Li et al., 2009). The S. suis strains were grown in Todd-Hewitt broth (THB; Oxoid, Basingstoke, England) or on THB agar (THA; Oxoid, Basingstoke, England) plates supplemented with 5% sheep blood (Maojie, Nanjing, China) at 37°C. The arginine metabolic pathway study was performed using a chemically defined medium (van de Rijn and Kessler, 1980). Erythromycin (90 μg/mL) was added to screen for the mutant strain, while erythromycin (90 μg/mL), and spectinomycin (100 μg/mL) were added to select for the complemented strain. The E. coli DH5α strains were grown in LB broth (Difco Laboratories, Franklin Lakes, NJ, USA) or on LB agar plates at 37°C. If necessary, kanamycin (25 μg/mL) was added.

The mnmE deletion strain was obtained using an homologous recombination method (Takamatsu et al., 2001). Primers used in this study were designed according to the genome sequence of S. suis strain 05ZYH33 (GenBank accession number: CP000407) and are listed in Table 2. Primers Mup-F/Mup-R and Mdown-F/Mdown-R were used to amplify the upstream and downstream regions of mnmE. Moreover, the fragments were cloned into pSET4s, respectively. Finally, the erm^r expression cassette was amplified from pAT18 using the primers Erm-F/Erm-R and inserted between the upstream and downstream homologous arms of the recombinant pSET4s to achieve the mnmE-knockout vector pSET4s-M. The pSET4s-M was then electroporated into SC-19. The ΔmnmE mutant strain was screened on THB plates for sensitivity to spectinomycin and resistance to erythromycin. To confirm the mutant strain, the mnmE gene, the upstream, and downstream genes were amplified by PCR using the primer pairs mnmE-F/mnmE-R, 1453-F/1453-R, and 1455-F/1455-R, respectively. Moreover, Primers Test-F/Test-R were used to amplify the upstream to downstream regions of mnmE in SC19 and ΔmnmE, and the resulting DNA fragments were confirmed by DNA sequencing.

To construct a complemented strain, a DNA fragment containing the entire mnmE coding sequence and its promoter and terminator was amplified by using primers CM-F/CM-R. The promoter and coding sequences of the mnmE gene were amplified by PCR using the specific primer pair CM-F/CM-R. The amplicon was subsequently cloned into pSET2 to obtain the recombinant plasmid pSET2-CM, which was electroporated into strain ΔmnmE. The complemented strain CΔmnmE was screened on THB plates for resistance to erythromycin and spectinomycin. To confirm the complemented strain, the mnmE gene, the upstream and downstream genes were amplified by PCR using three pairs of primers mnmE-F/mnmE-R, 1453-F/1453-R, and 1455-F/1455-R.

To further confirm the mutant strain ΔmnmE and the complementary strain CΔmnmE, we also performed RT-PCR (Tan et al., 2015). Briefly, RNA was isolated using the Bacterial RNA Kit (Omega Bio-Tek, Inc., Norcross, GA, USA) in accordance with the manufacturer's instructions. In addition, cDNA was synthesized using the HiScript® II Q Select RT SuperMix for qPCR kit (Vazyme Biotech Co., Ltd., Nanjing, China) in accordance with the manufacturer's instructions. The primers mnmE-F/mnmE-R were used to confirm the deletion of mnmE gene, while, the primers 1453-F/1453-R (for upstream gene), and 1455-F/1455-R (for downstream gene) (Table 2) were used to confirm whether the upstream and downstream genes of mnmE were unaffected and functioning normally.

Growth rates of the SC19, ΔmnmE, and CΔmnmE were detected through the measurement of the density changes represented by OD600 nm values and CFU counts of the cultures. Different strains were grown to the mid-exponential phase (OD₆₀₀ of 0.6–0.8) in THB medium, then the cells were collected and washed in phosphate-buffered saline (PBS) three times, and the initial OD600 nm of all subcultures were adjusted to 0.07. Each of the subcultures was incubated at 37°C with rotating at 180 rpm and OD600 nm values were read every hour until the growth process entered the stationary phase. Meanwhile, a 100 μL aliquot of the bacterial culture was diluted and the number of viable bacteria was calculated every hour.

To probe the possible role of the MnmE in S. suis virulence, 30 female specific-pathogen-free (SPF) Kun-Ming mice (6-week-old) weighing 26 to 30 g were randomly allocated to three groups (10 mice per group). Groups 1 and 2 were inoculated by intraperitoneal injection with 3 × 10⁹ CFU of either SC19 or ΔmnmE. Saline was administered to group 3 as a negative control. Clinical signs and survival time were recorded. The mice were observed for 7 days to obtain steady survival curves.

To better evaluate the pathogenicity of ΔmnmE, we performed a determination of viable bacteria in organs assay as described previously (Tan et al., 2017). Fifteen female SPF Kun-Ming mice (6-week-old) weighing 26 to 30 g were inoculated by intraperitoneal injection with 1 × 10⁸ CFU of a 1:1 mixture of mid-log-phase SC19 or ΔmnmE. Saline was applied to five mice as a negative control. At 12 h, 1 day, and 3 days post infection (dpi), brain, lung, spleen, and blood were obtained from five mice from each group. The samples were homogenized after weighing, and serial dilutions were plated onto THA. In order to count the colonies, we used 20 μg/mL streptomycin for SC19 and ΔmnmE, while 20 μg/mL streptomycin, and 90 μg/mL erythromycin were used for ΔmnmE.

Adhesion and invasion assays were conducted in accordance with previously described methods (Ferrando et al., 2014). For the adhesion assay, HEp-2 cells were infected with mid-log-phase SS2 to reach a multiplicity of infection (MOI) of 100:1 (bacteria:cells) and then incubated for 30 min at 37°C. Unbound bacteria were removed by washing the cells gently with PBS three times and then cells were lysed in 1 mL of sterile distilled water. Adherent bacteria (cell-associated bacteria) were determined by plating a serial dilution of the lysates on THA plate. For the invasion assay, cells were incubated with bacteria for 2 h to allow for invasion and then incubated in medium containing penicillin (100 μg/mL) for 2 h to kill extracellular and surface-adherent bacteria. The numbers of invading bacteria were determined the same as the adhesion assay.

AD activity was determined by measuring the production of L-citrulline from L-arginine and evaluated according to the protocol described previously (Winterhoff et al., 2002; Xiong et al., 2014). Briefly, bacteria were grown in chemically defined medium to the mid-log phase and harvested by centrifugation. Then, cultures of test strains were lysed and, respectively, incubated for 2 h in 0.1 M potassium phosphate buffer containing 10 mM L-arginine at 37°C. Afterward, the enzymatic reaction was stopped by adding 250 μL of a 1:3 (v/v) mixture of 95% H₂SO₄ and 85% H₃PO₄, and 250 μL of 3% diacetylmonooxime solution were added to the samples incubating for 15 min at 100°C. Production of citrulline was determined colorimetrically at OD₄₅₀. A citrulline standard and the uninoculated reagents were used as positive and blank controls, respectively. Results were expressed as nanomoles of citrulline produced per h per mg of whole cell protein.

Ammonia production in the culture supernatant of wild-type, mutant, and complementary strains was quantified using an Ammonia Assay kit (Sigma-Aldrich Corporation, St. Louis, MO, USA) in accordance with the manufacturer's instructions.

SC19 and ΔmnmE cells at mid-log phase were cultured in THB as described above. Three independent biological replicates of bacterial pellets were then treated with SDT buffer (4% SDS,100 mM Tris-HCl, 1 mM DTT, pH 7.6) and heated for 15 min at 100°C. The cell suspensions were sonicated for 5 min (10 s of sonication with 15 s intervals) on ice and the protein concentration in the supernatants was determined using the Bradford protein assay. Each sample (200 μg) were digested with 3 μg of trypsin (Sigma-Aldrich Corporation) at 37°C for 16 h. The resulting tryptic peptides were labeled according to the protocol of TMT Reagent Kit (Thermo Fisher Scientific, Waltham, MA, USA). The labeled peptides were combined and fractionated by strong cation exchange (SCX) chromatography.

After separation by SCX chromatography, equal amounts of digested protein were loaded into a Thermo Scientific EASY column (2 cm*100 μm 5 μm-C18) and then washed with solvent A (99% H2O, and 0.1% formic acid). By applying solvent B (84% acetonitrile, 16% H₂O, and 0.1% formic acid), the peptides were eluted from the trapping column over an EASY-Column™ (75 μm × 100 mm, 3 μm, C18–42; Thermo Fisher Scientific) with a gradient (0–45% solvent B for 100 min, 35–100% solvent B for 8 min, 100% solvent B for 12 min at 300 nL/min) using the Easy nLC system (Thermo Fisher Scientific). For MS analysis, peptides were analyzed in positive ion mode. MS/MS was carried out with a Q-Exactive mass spectrometer (Thermo Finnigan LLC, San Jose, CA, USA) in the positive ion mode and a data-dependent manner choosing the most abundant precursor ions with a full MS scan from 300 to 1,800 m/z and resolution of 70,000 at m/z of 200. Target value determination was based on automatic gain control and dynamic exclusion duration was 60 s. MS/MS scans were acquired at a resolution of 35,000 at m/z of 200. Normalized collision energy was 30 eV and the underfill ratio was set at 0.1%.

The data files produced by 15 fractions MS/MS were processed by Proteome Discoverer 1.4 and searched by Mascot 2.2 (Matrix Science, MA) against 83,725 S. suis protein-coding sequences deposited in the Uniprot database (downloaded on July 30, 2018). The search was conducted with trypsin applied as a specific enzyme and parameters used for normal peptides as follows: peptide mass tolerance, 20 ppm; fragment mass tolerance, 0.1 Da; max missed cleavages, 2; fixed modifications, carbamidomethyl (C), TMT 6/10plex (N-term), TMT 6/10plex (K); variable modifications: oxidation (M), TMT 6/10plex (Y); database pattern, decoy; and false-discovery rate ≤0.01 (Sandberg et al., 2012). Each of the identified proteins involved at least two unique peptides. Protein quantification was accomplished by correlating the relative intensities of reporter ions extracted from tandem mass spectra to that of the peptides selected for MS/MS fragmentation. To evaluate the DEPs between different strains, a fold change (ΔmnmE/SC19) > 1.2 and < 0.83, and a p-value of <0.05 were considered to represent up- or down-regulation, respectively. The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD012716.

Statistical analyses were performed via unpaired Student's t-tests in GraphPad Prism 5 software (GraphPad Software, Inc., La Jolla, CA, USA) all experiments were performed in triplicate at least three times. All data are expressed as the mean ± the standard error of the mean. A p-value of <0.05 was considered as the threshold for significance.

Colonies sensitive to spectinomycin and resistant to erythromycin were selected as candidates of mnmE-deletion mutants, and confirmed by PCR and RT-PCR (Figures 1A,B). At the gene and transcript levels of mnmE was expressed in strains SC19 and CΔmnmE, but not in strain ΔmnmE. RT-PCR analysis also showed that the transcripts of the genes upstream and downstream from mnmE were not affected by the mnmE deletion, which could exclude associated polarity effects.

First, the growth kinetics of the mutant strain were characterized. The SC19, ΔmnmE, and CΔmnmE strains were grown to the mid-log phase and then inoculated in fresh THB at 37°C while rotating at 180 rpm. The growth rate of the mnmE-deficient cells was reduced as compared to that of strain SC19. Meanwhile, the CFU counts also showed that strain ΔmnmE grew much slower than strain SC19 (Figures 1C,D). Indeed, complementation of mnmE (CΔmnmE) almost restored the growth defect. Notably, the colonies of ΔmnmE appeared smaller than those of SC19 when cultured on THB plates overnight. Based on these observations, MnmE appears to be critical for S. suis growth.

Mice were experimentally infected to estimate differences in viability between strains ΔmnmE and SC19 in vivo. All 10 SC19-infected mice developed severe clinical symptoms, such as septicemia and meningitis, and died within the first day of infection. By contrast, the symptoms of the ΔmnmE-infected mice were milder and the mortality rate was relatively low (2/10) during the 7-day observational period (Figure 2A), indicating that the pathogenicity of ΔmnmE was markedly attenuated.

To further evaluate the pathogenicity of ΔmnmE, a colonization experiment was performed using the intraperitoneal route of inoculation. First, the optical density at 600 nm (OD₆₀₀) and CFU counts showed that in vitro, strain ΔmnmE grew much slower than strain SC19, and the colonies of ΔmnmE appeared smaller as compared to strain SC19. Hence, we investigated whether the characteristics would be the same under in vivo conditions. As expected, the in vivo adaptability of ΔmnmE was decreased as compared to that of SC19. Bacteria were recovered from brain, lung, spleen, and blood samples at different time points post infection. The bacterial loads of ΔmnmE in these tissues were lower than those of SC19 from 12 h to 3 dpi, and the mutant strain was partly cleared at 3 dpi (Figures 3A–D).

Next, the role of mnmE in the adhesion to and invasion in host cells was investigated. The efficiencies of SC19 and its derivatives to adhere to and invade in HEp-2 cells were calculated. The binding and invasion rates of SC19 to the HEp-2 cells were 2- and 2.5-fold greater than that of ΔmnmE (Figures 2B,C), indicating that the inactivation of mnmE impaired the capacity of S. suis to adhere to and invade in epithelial cells.

To understand the underlying molecular mechanisms of the phenotype of ΔmnmE, TMT-based MS profiles of proteins produced by strains SC19 and ΔmnmE were applied. In this study, a total of 1,619 proteins were identified and quantified via TMT proteomics, of which 365 were DEPs, including 174 that were up-regulated and 191 down-regulated (Table S1).

According to gene ontology analysis, the 365 DEPs were classified into biological processes, cellular components, and molecular functions (Figure 5). The most prevalent biological process were metabolic processes (149, 40.8%) and cellular processes (139, 38.1%), which are the most important responses of S. suis to environmental stressors. The top two molecular functions were catalytic activity (235, 64.4%) and binding (122, 47.1%). The most common cellular components were the cell (104, 28.5%), followed by cell parts (102, 27.9%), the membrane (68, 19.1%), and membrane parts (62, 17.0%).

Inactivation of S. suis MnmE affects the bacterial phenotype, including the size of the colonies. There are two reasons for this phenotype: (i) reduced growth, as verified by CFU counts and absorbance at OD₆₀₀; and (ii) the smaller size of strain ΔmnmE, as compared to strain SC19. Analysis of cell growth kinetics and transmission electron microscopy were both performed in this study. CFU counts and OD₆₀₀ readings revealed that strain ΔmnmE grew much slower than strain SC19 and there was no obvious difference in the cell size between strains, as observed via transmission electron microscopy (Figure S1).

These results indicate that MnmE is involved in the growth regulation of S. suis. This finding is in agreement with those of previous reports on E. coli and S. enterica serovar Typhimurium. To further understand the molecular mechanism of MnmE on growth regulation, TMT-based proteomic analysis was conducted. According to the DPE analysis, many proteins involved in cell growth and division were down-regulated in the ΔmnmE mutant, which included three classes of proteins: (i) those related to DNA replication, recombination, and repair, such as DNA/RNA helicases (YjqA and PcrA), DNA topoisomerase I (TopA), DNA/RNA helicase (PcrA), site-specific recombinase (XerD), DNA mismatch repair protein (MutS), DNA-directed DNA polymerase (DnaE), and ribonuclease (RNY); (ii) cell division, including FtsK and StpK; and (iii) cell wall biosynthesis, including MurD, MurE, MurG, and PBP1A, which positively regulate cell division (Eniyan et al., 2018; Szafran et al., 2018; Zucchini et al., 2018). We previously reported that STK, a serine/threonine protein kinase, regulates cell growth and division. Phosphoproteomics revealed that the down-regulated substrates of STK were all crucial cell division-associated proteins, which included FtsA, GpsB, DivIVA, and MapZ (Zhang et al., 2017). Therefore, down-regulation of STK could contribute to the slow growth rate of the ΔmnmE mutant. As we know, cell wall peptidoglycan (PG) is vital for bacterial survival and normal cell growth, and thus is an attractive drug target in bacterial infection (Bhat et al., 2017). The cytoplasmic biosynthesis of PG is a complex process that involves six MurA-F enzymatic reactions to catalyze the formation of PG units from the primary substrate uridine diphosphate N-acetylglucosamine (Eniyan et al., 2018). In addition, PBP1A, which catalyzes the cross-linking reaction of polymeric glycan chains (Frere and Page, 2014), was also down-regulated. However, two cell growth and division-associated proteins, MurB and FtsE, were up-regulated. The MurB enzyme belongs to a family of proteins that contain FAD-binding domains and share a characteristic FAD binding fold (Eniyan et al., 2018), the same as GidA, which is the partner of MnmE. As mentioned above, GidA and MnmE form an α2β2 heterotetrameric complex that controls the addition of a cmnm group at the wobble position of tRNA molecules (Moukadiri et al., 2014). Based on these considerations, we suppose that the deletion of mnmE renders GidA more available for FAD-binding. Thus, the mutant up-regulates MurB to compete with GidA for FAD-binding. FtsE together with FtsX forms a dimer that acts as an ABC transporter. The FtsEX protein complex plays a major role in the regulation of PG hydrolases in response to signals from cell division (Sham et al., 2013). Any disruption, either through inhibition or overactivation of the hydrolases, results in the inability to maintain the function of the cell wall (Margulieux et al., 2017), therefore, up-regulation of FtsE may impair cell growth.

The deletion of mnmE in S. suis also resulted in an alteration to bacterial pathogenicity. In vivo and ex vivo studies revealed that ΔmnmE is associated with decreased mortality (Figure 2A) and bacterial loads in mice (Figure 3), as well as reduced adhesion to and invasion in epithelial cells (Figures 2B,C). This attenuation may result from the impaired growth of ΔmnmE or impaired translation efficiency of the virulence-associated proteins.

In this study, analysis of protein expression profiles indicated that several proteins associated with adherence and hemolysis were repressed in ΔmnmE, such as FBPs and SadP (Ge et al., 2004). A considerable number of FBPs of various bacterial species are crucial virulence factors (Joh et al., 1999). In S. pyogenes, FBPs were expressed in the human host and preferentially mediated adherence to human buccal epithelial cells (Courtney et al., 1996). In S. suis, FBPs play roles in the colonization of specific organs during infection (de Greeff, 2002). Our results showed that mnmE disruption decreased the abilities of S. suis to adhere to and invade in epithelial cells, which can be explained by the down-regulation of the above DEPs. As previously reported, the tRNA-modifying GTPase MnmE has been implicated in the bacterial response to stressors other than low pH. In this study, we found that the ADS of S. suis (ArcABC), which was down-regulated, is involved in the adhesion to and invasion in epithelial cells (Degnan et al., 2000; Fulde et al., 2014), as well as resistance to oxygen, nutrient starvation, and acidic environments (Gruening et al., 2006). Here, only the arginine repressor ArgR was upregulated. ArgR is an essential transcriptional regulator that specifically regulates the arcABC operon by directly interacting with its promoter (Fulde et al., 2011). Interestingly, the protein expression of ArcABC was remarkably down-regulated rather than up-regulated, indicating that the translation efficiency mediated by MnmE has a major impact on ADS expression far beyond the role of ArgR in transcriptional activation. The biological significance of tRNA modification by MnmE on these virulence-associated proteins warrants further investigations.

According to the KEGG pathway analysis results, seven metabolic pathways were significantly perturbed following mnmE deletion (Table 4). The fatty acid biosynthesis, arginine biosynthesis, and purine metabolic pathways were inhibited, while the valine, leucine, and isoleucine biosynthesis, cofactor, and vitamin metabolic pathways were activated. These perturbations could be related to the impaired growth and virulence of ΔmnmE.

Lipid metabolism is thought to play a key role in the pathogenicity of several bacteria (Bohl et al., 2018; Ghazaei, 2018; Rameshwaram et al., 2018). There are three main functions provided by this pathway in bacteria: (i) the activation of bacterial lipolytic enzymes that hydrolyze lipids from the host cell to release free fatty acids, which are used as an energy source; (ii) the formation of building blocks for the synthesis of the bacterial cell wall, and (iii) modulation of the host immune responses (Rameshwaram et al., 2018). Our data showed that all the DEPs in the lipid metabolic pathway were down-regulated, which could help to explain the attenuated virulence and slow growth rate of the mutant strain ΔmnmE.

Deletion of mnmE down-regulated three core enzymes associated with arginine metabolism (ArcA, ArcB, and ArcC), which participate in the ADS. As a secondary metabolic pathway, ADS catalyzes the conversion of arginine to ornithine, as well as carbon dioxide and ammonia, with the concomitant release of adenosine triphosphate (Figure 4C). There are three main functions of the enzymes involved in the ADS: (i) production of energy via arginine metabolism, (ii) provision of carbamoyl phosphatase for the biosynthesis of citrulline or pyrimidines, and (iii) protection against acidic damage by the production of NH₃. Based on the above theory, ADS involvement has been confirmed in the pathogenicity and adaptability to acidic conditions of many bacteria, such as Streptococcus gordonii, Listeria monocytogenes, S. suis, Streptococcus pyogenes, Laribacter hongkongensis, and E. coli (Liu et al., 2008; Smith et al., 2013; Fulde et al., 2014; Xiong et al., 2014; Sakanaka et al., 2015; Willenborg et al., 2016). A previous report has also stated that mnmE deficiency could affect the adaptability of E. coli to an acidic environment (Vivijs et al., 2016). This molecular mechanism can be explained by the down-regulated activity of the ADS through MnmE. Therefore, inhibition of the arginine metabolic pathway significantly contributed to reduced pathogenicity and arginine deiminase activity of the ΔmnmE mutant.

About two-thirds of the proteins (14 of 20) involved in the purine metabolic pathway were repressed in strain ΔmnmE. Previous studies have emphasized the importance of nucleotide biosynthesis in bacteria. Auxotroph mutants of nucleotide biosynthesis in S. aureus, Salmonella, and S. pneumoniae have already been associated with stunted growth (McFarland and Stocker, 1987; Polissi et al., 1998; Mei et al., 2003; Fitzsimmons et al., 2018). However, the biosynthesis of valine, leucine, and isoleucine, cofactor production, and vitamin metabolic pathways were activated, while the DEPs in these pathways were overlapped. The connection between the deletion of MnmE and activation of these pathways is unclear, thus further investigations are needed to determine exactly how MnmE regulates these pathway in S. suis.

In conclusion, our in vitro and in vivo studies clearly demonstrate the important roles of MnmE on the growth, pathogenicity, and arginine metabolism of S. suis. Consistently, TMT analysis also confirmed these results. These findings provide new insights to better understanding the function of MnmE as a central tRNA-modifying GTPase and stress the importance of this protein in the mRNA decoding process in bacterial pathogens.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.