extfieldsofstudy: Biology Medicine

text:
A gacS Deletion in Pseudomonas aeruginosa Cystic Fibrosis Isolate CHA Shapes Its Virulence

Pseudomonas aeruginosa, a human opportunistic pathogen, is capable of provoking acute and chronic infections that are associated with defined sets of virulence factors. During chronic infections, the bacterium accumulates mutations that silence some and activate other genes. Here we show that the cystic fibrosis isolate CHA exhibits a unique virulence phenotype featuring a mucoid morphology, an active Type III Secretion System (T3SS, hallmark of acute infections), and no Type VI Secretion System (H1-T6SS). This virulence profile is due to a 426 bp deletion in the 3′ end of the gacS gene encoding an essential regulatory protein. The absence of GacS disturbs the Gac/Rsm pathway leading to depletion of the small regulatory RNAs RsmY/RsmZ and, in consequence, to expression of T3SS, while switching off the expression of H1-T6SS and Pel polysaccharides. The CHA isolate also exhibits full ability to swim and twitch, due to active flagellum and Type IVa pili. Thus, unlike the classical scheme of balance between virulence factors, clinical strains may adapt to a local niche by expressing both alginate exopolysaccharide, a hallmark of membrane stress that protects from antibiotic action, host defences and phagocytosis, and efficient T3S machinery that is considered as an aggressive virulence factor.

Introduction
Pseudomonas aeruginosa is an opportunistic Gram negative bacterium able to trigger either severe acute or chronic human infections, depending on the environmental signals it encounters. The persistence of the bacterium during decades in the lungs of individuals with cystic fibrosis (CF) is associated with massive and inefficient inflammation contributing to airway epithelium destruction, decline of lung function and death due to respiratory failure [1]. During chronic infection, P. aeruginosa has to survive and adapt to the stressful environment encountered in the CF lungs where it is continuously exposed to antibiotics, oxidative and osmotic stresses as well as active host immune system. The CF respiratory mucus has been shown to directly impact bacterial gene transcription [2,3]. A CF mucoid strain was reported to trigger, in response to the CF niche, the synthesis of enzymes protecting the bacteria against oxidative stress and the activation of genes encoding the HSI-I Type VI Secretion System (H1-T6SS), known to play a role in bacterial competition [4,5,6]. In addition, the alginate production was unexpectedly repressed, and the expression of two small RNAs (PA2G_05393.1 and PA2G_03487.1) with putative regulatory roles was observed, pointing out the major effect of contact with CF mucus on bacterial physiology [3].
Besides these immediate adaptive responses, P. aeruginosa is also prone to accumulate point mutations and/or significant genomic rearrangements induced by extrinsic and intrinsic factors associated with CF chronic disease [1,7]. Mutations in global regulatory genes, such as lasR, vfr, rpoS, or other regulatory genes such as mucA, are frequently observed in CF isolates and these mutations profoundly affect virulence gene expression [1,8]. The most obvious phenotypic changes of P. aeruginosa observed during infection in CF disease are conversion to mucoidy, emergence of Small Colony Variants (SCV), acquisition of antibiotic multiresistance, loss of motility and shutdown of quorum sensing (QS) system [7,9]. Conversion to mucoidy is proposed to be a major survival mechanism promoting the persistence of the bacterium in CF lungs; it mostly results from the over-expression of the alginate polysaccharide by the alternative sigma factor AlgU, desequestrated from its anti-sigma partner MucA due to mutations in mucA [10,11]. More aggressive virulence factors secreted by Type II (T2SS) and Type III (T3SS) secretion machineries are switched off, thus leading to CF-adapted strains usually less virulent in animal models than the primary infecting ones [9]. However, while sharing some generally-accepted traits, adaptive evolution and resulting phenotypes of CF isolates can be extremely diverse, even within isolates from the same CF patient sputum sample [1,9,12,13].
The CHA strain is a good example of P. aeruginosa multifaceted adaptation in CF disease. It was originally isolated from a CF patient [14], 4 years after the first airways colonization by P. aeruginosa, and is thus not considered as an early infecting strain. This is an O6 serotype strain belonging to clone J, one common clonal group disseminated worldwide [15]. CHA is mucoid [16], a hallmark of chronicity, but at the same time, it is a highly efficient T3SS effector-producer fitting with acute infectious status [17,18,19,20]. Very recently, CHA has been included in the international P. aeruginosa reference panel, stressing the relevance to study its pathogenicity [21].
Using phenotypic and complementation experiments in parallel to genome analysis, we report in the present study that virulence properties of CHA result from an intrinsic genetic deletion leading to the absence of the histidine kinase (HK) GacS. GacS/GacA is a two-component regulatory system (TCS) that controls transcription of two small regulatory RNAs (sRNAs), RsmY and RsmZ. These two sRNAs prevent RsmA binding to its mRNA targets and consequently modulate, directly or indirectly, approximately 500 genes belonging to the RsmA regulon [22,23,24]. Activity of GacS is regulated in an opposite manner by two inner membrane sensors, RetS and LadS, in response to still unidentified stimuli [25,26]. The RetS/LadS/Gac/Rsm cascade is known to be a master regulator of the virulence factors of P. aeruginosa, controlling the switch in their expression during the acute/chronic phase transition (reviewed in [27]). Hence, as illustrated here, mutations in this regulatory pathway profoundly impact the global virulence traits.

Animals
All protocols in this study were conducted in strict accordance with the French guidelines for the care and use of laboratory animals. The protocol for mouse infection was approved by the animal research committee of the institute (CETEA: Comité d' É Thique en Expérimentation Animale). Pathogen-free BALB/c male mice (8-10 weeks) were obtained from Harlan Laboratories and housed in the CEA animal care facilities.

Genetic constructions
Deletion of rsmA and retS -Fused uspstream and downstream flanking regions of rsmA and retS were amplified by Splicing by Overlap Extension-Polymerase Chain Reaction (SOE-PCR) procedure using appropriate primer pairs (Table S1). The resulting fragments of 846 bp and 834 bp, respectively, were cloned into pCR-Blunt II-TOPO vector, sequenced and then subcloned into the SmaI site of the suicide plasmid pEX100-T. The resulting pEXDRsmA and pEXDRetS plasmids carry the counter-selectable sacB marker from Bacillus subtilis, which confers sensitivity to sucrose. Both plasmids were mobilized into P. aeruginosa strain by triparental mating, using the conjugative properties of the helper plasmid pRK2013. Co-integration events were selected on PIA plates containing carbenicillin. Single colonies were then plated on PIA medium containing 5% (w/v) sucrose to select for the loss of plasmid: the resulting sucrose-resistant strains were checked for carbenicillin sensitivity and for rsmA or retS (wild-type or deleted gene) genotype by PCR.
Fusion of VSV-G epitope to GacS -The VSV-G-Tag-coding sequence was fused to the 39 end of chromosomal PAO1 gacS or to that of gacS*, the CHA naturally truncated gacS gene, using SOE-PCR strategy. Upstream and downstream flanking regions of gacS and gacS* were PCR amplified using PAO1 or CHA genomic DNA, respectively, and appropriate primer pairs (see Table S1). The resulting fragments of 813 bp and 752 bp, respectively, were cloned into pCR-Blunt II-TOPO vector, sequenced, then inserted into the SmaI site of pEX100-T, leading to pEX-GacS-VSV-G and pEX-GacS*-VSV-G plasmids. Allelic replacement was performed as described above. The presence of VSV-G-encoding sequences in the PAO1 and CHA genomes was assessed by PCR and confirmed by sequencing of PCR fragments.
Complementation -The rsmA and gacS genes were PCR amplified from PAO1 genomic DNA using appropriate primer pairs (Table S1). The PCR products were cloned into pCR-Blunt II-TOPO and sequenced. The 0.23-kb XbaI-HindIII restriction fragment containing rsmA was cloned in pVLT31 plasmid under control of the IPTG-inducible ptac promoter. The 2.85-kb SmaI-XbaI restriction fragment containing gacS was cloned into pJN105 plasmid under the arabinose-inducible pBAD promoter. The two expression vectors were introduced in P. aeruginosa by transformation [39].
An integrative plasmid was also constructed to complement gacS mutation with one copy of the gene driven by its own promoter. A region comprising the 245 bp sequence upstream from gacS and the entire gene was PCR amplified from PAO1 genomic DNA using appropriate primers (Table S1). After sequencing, the SmaI-HindIII restriction fragment was cloned into pUC18-miniTn7T-Gm-LacZ20. The resulting plasmid was electroporated into the CHA strain along with the pTNS2, as described [34], leading to the CHA-GacS strain.
b-Galactosidase assays b-Galactosidase activity was assayed as already described [40].

T3SS-dependent cytotoxicity assay
Bacteria were grown in LB to an A 600 1.0 and added to the macrophage cell line J774 (J774A.1, catalog No. TIB-67, ATCC) at a multiplicity of infection (MOI) of 5. Cell death was assessed at 3 h post-infection by using a cytotoxicity detection kit (lactate dehydrogenase LDH; Roche) as described [17].

Sample preparation, antibodies and immunoblot analysis
H1-T6SS Hcp1 (for haemolysin-coregulated protein 1) production and secretion were assessed as described [41]. For GacS-VSV-G and GacS*-VSV-G analysis, 100 ml of cultures at A 600 of 2.8-3.0 were harvested and analyzed as the total bacterial fraction. Membrane fractions were recovered after lysozyme treatment, sonication and ultracentrifugation. The samples were submitted to SDS-PAGE and immunoblotting analysis.

Motility assays
Motilities were assayed on media as described [42]. All plates were inoculated with bacteria from overnight cultures on LB agar using sterile toothpicks.

Air-liquid biofilm
The P. aeruginosa adherence assay was performed in individual glass tubes as previously described [43].

RT-PCR and RT-qPCR
The strains were grown at 37uC under agitation in LB, that was supplemented with 5 mM EGTA and 20 mM MgCl 2 (conditions of in vitro T3SS induction) for Reverse Transcriptase (RT)-qPCR analysis. Total RNA was either extracted with the TRIzol Plus RNA Purification Kit (Invitrogen) then treated with DNase I (Amplification Grade, Invitrogen), or the PureYield RNA Mid-iprep System (Promega), cleaned up and concentrated using the RNeasy kit (Qiagen). Yield, purity and integrity of RNA were further evaluated on Nanodrop and by agarose gel migration. Complementary DNA synthesis was carried out with SuperScript III First-Strand Synthesis System (Invitrogen) in presence or not of the SuperScript III RT enzyme to assess the absence of genomic DNA. For RT-PCR experiments, performed to assess the presence of GacS mRNA, PCR amplifications were performed using TaqPCRx DNA Polymerase (Invitrogen) and following the Basic PCR protocol described by the manufacturer. Calibration of the PCR amplification steps was done by varying number of cycles with primers targeting either gacS or 16S rRNA as a reference transcript. The RT-qPCR runs, used to quantify the effect of GacS on its target genes, were carried out on a CFX96 Real-Time System (Bio-Rad). Cycling parameters of the real time PCR were 98 uC for 2 min, following by 45 cycles of 98uC for 5 s and 60uC for 10 s, ending with a melting curve from 65 uC to 95 uC to assess the specificity of the amplification. To determine the amplification kinetics of each product, the fluorescence derived from the incorporation of EvaGreen into the double-stranded PCR pRK2013 Kn r ; helper plasmid with conjugative properties [33] pUC18-mini-Tn7T-Gm r ; translational fusion vector lacZ20-Gm [34] pUC18-mini-Tn7T-Promoter and entire gacS sequence of PAO1 cloned into pGacS the integrative vector This study pCTX-PA0081-lacZ Tc r ; pfha1-lacZ transcriptional fusion in mini-CTX-lacZ [23] pJN105 Gm r ; pBAD transcriptional fusion vector [35] pJN-GacS pBAD-gacS transcriptional fusion in pJN105 This study pVLT31 Tc r ; plac transcriptional fusion vector [36] pVLT-RsmA plac-rsmA transcriptional fusion in pVLT31 This study pMP220 Tc r ; lacZ transcriptional fusion vector [37] pMP220-rsmY-lacZ prsmY-lacZ transcriptional fusion in pMP220 [38] pMP220-rsmZ-lacZ prsmY-lacZ transcriptional fusion in pMP220 [38] doi:10.1371/journal.pone.0095936.t001 products was measured at the end of each cycle using the SsoFast EvaGreen Supermix 2X Kit (Biorad). The results were analyzed using the Bio-Rad CFX Manager Software 3.0 (Bio-Rad). The relative mRNA quantity of each gene under GacS production as compared to absence of production was analyzed using the Relative Expression Software Tool REST2009 (Qiagen) with a pair wise fixed reallocation randomization test [44] coupled to a standard error (SE) calculated via a Taylor algorithm. The 16S rRNA was used as reference for normalization. The sequences of all primers are given in Table S1.

H1-T6SS expression is off in CF strain CHA
The mucoid CF isolate CHA has been previously reported as harboring T3SS-dependent cytotoxicity in vitro [45,46]. As CF strains usually exhibit a decreased toxicity compared to strains triggering acute infection [9], we compared the virulence of CHA in mice to that of the PAO1 reference strain, which is a wound isolate. The CHA strain was clearly more virulent than the PAO1 strain in a murine acute model of lung infection ( Figure S1A); this was associated with an increased dissemination of the bacteria both in blood and spleen compared to PAO1 ( Figure S1B). The CHA T3SS-deficient strain exhibited a reduced toxicity in the model (data not shown) which is in agreement with a previous report pointing to T3SS being a major actor in CHA virulence [18].
We then examined the expression of H1-T6SS (HSI-I), a machinery expressed during chronic infection [4,5] and secreting bacterial toxins used for competing with other species in biofilm communities. Surprisingly, H1-T6SS production was undetectable in CHA as compared to other strains, notably CF strains TB, LES400, KK1 and CF6, or reference strains PAO1, PA7 and PAK, as assessed by the production of the Hcp1 component of the H1-T6SS apparatus ( Figure 1A). The transcriptional fusion fha1-lacZ (PA0081) was expressed in CHA, albeit 2.4 fold less than in PAO1, while the corresponding translational fusion exhibited no reporter activity in CHA ( Figure 1B). These results strongly suggested that CHA is incapable of H1-T6SS synthesis due to a default of mRNA stability and/or protein translation rather than an absence of HSI-I gene transcription.

Levels of RsmA and RsmY/Z are perturbed in CHA
T3SS and the H1-T6SS are known to be modulated in an opposite manner by the translational regulator RsmA, whose activity is antagonized by the two sRNAs, RsmY and RsmZ [23,25]. Thus, we examined the rsmY and rsmZ expression in CHA. The rsmY gene was not expressed in CHA, and only a slight expression of rsmZ was observed, mainly in overnight culture (Figure 2), while in PAO1, the two sRNAs were efficiently transcribed, with higher level of rsmY expression compared to that of rsmZ (Figure 2), as already reported [3,24]. These data strongly suggested that, in CHA strain, amounts of RsmY and RsmZ sRNAs were not sufficient to efficiently titrate RsmA and to relieve its effect on its target mRNAs. Indeed, high amounts of Hcp1 were produced in the supernatant and whole cell extracts of the CHADrsmA mutant, a phenotype corrected by introduction of the rsmA wild-type gene in trans ( Figure 3A). Additionally T3SS activity, assessed either by in vitro calcium-induced secretion of translocator proteins (data not shown) or T3SS-dependent cytotoxicity on macrophages ( Figure 3B), was completely abolished by rsmA deletion and further re-induced by rsmA complementation.
Taken together, the absence of H1-T6SS synthesis in the CHA strain, resulting from a constant translational inhibition exerted by RsmA on H1-T6SS mRNAs, and the highly active T3SS, due to RsmA-mediated positive post-transcriptional regulation of ExsA expression [47], are both related to a defective rsmY/Z gene expression.

A 426 bp chromosomal deletion led to a truncated and unstable GacS
We then investigated other regulatory players that could be involved in this deregulation by examining the recently sequenced genome of the CHA strain [48]. Indeed, a 426 bp deletion was found in the chromosome that affects the 39 end (last 146 nucleotides) of gacS gene and the 59 end (first 278 nucleotides) of the downstream gene ldhA, encoding a lactate dehydrogenase ( Figure 4A) [48]. This gacS deletion generates a putative truncated protein that we named GacS*, possessing 19 unrelated residues in place of the 48 C-terminal residues of the PAO1 GacS protein ( Figure 4B). The three phosphorylation sites (residues His-293, Asp-715 and His-859) are conserved ( Figure 4C) but the fifth helix of the alternative transmitter domain (H2), also called Histidine PhosphoTransfer (Hpt) domain, is missing ( Figure 4D).
To test whether this deletion impacts the stability and/or functionality of GacS*, we first provided in trans the Hpt domain (H2 domain) of PAO1 GacS protein [49], which did not complement the GacS* function (not shown). Thus we assessed by immunoblot the presence of the GacS and GacS* proteins in PAO1 and CHA, respectively. To do so, we first added at the 39 end of the endogenous genes a sequence encoding the VSV-G Tag, generating GacS-VSV-G and GacS*-VSV-G proteins (see Materials and Methods). GacS*-VSV-G could not be detected in CHA, neither in the membranes nor in the whole cells, although GacS-VSV-G was clearly present in the membranes of the PAO1 strain ( Figure 4E). We also ruled out the possibility that the absence of GacS* was a consequence of a transcription defect or an effect of the genomic deletion on mRNA stability by comparing the level of gacS transcripts in both PAO1 and CHA strains using RT-PCR ( Figure 4F). Altogether, these results strongly suggested that truncation of GacS C-terminus in CHA led to an unstable protein.
We next provided the PAO1 gacS gene cloned under the control of the pBAD promoter to the CHA strain. Introduction of the functional gacS gene increased the amounts of rsmY and rsmZ transcripts in CHA as assessed by RT-qPCR ( Figure 5), data corroborated by the restoration of rsmY/Z promoter activity ( Figure 2). It also increased quantity of PA0094 transcripts, PA0094 being a part of the H1-T6SS-encoding cluster ( Figure 5), as well as the synthesis of Hcp1 ( Figure 3A). In addition, providing gacS gene to CHA strain significantly reduced the level of exoS transcription, thus affecting the T3SS-dependent cytotoxicity on J774 macrophages ( Figure 3B). Furthermore, even though retS deletion in CHA did not affect T3SS activity (due to absence of a functional GacS protein), introducing gacS in the CHADretS background abolished it completely ( Figure 3B). These features indicate that absence of a functional GacS may allow CHA to escape from the negative control exerted by GacS/GacA TCS on T3SS during chronic infection.
Taken together, these results demonstrate that the truncation of GacS in CHA is responsible for defect in rsmY/Z gene expression as well as for alterations observed in T3SS and H1-T6SS synthesis.

The physiological absence of GacS shapes the phenotype of the clinical isolate
To get an overview of the impact of GacS absence in CHA, we investigated some of the major phenotypic traits of the bacterium.
At first we studied its swimming and swarming motilities [42], as RsmA positively affects genes involved in formation of Type IVa pili and function of the flagellum [22]. These extracellular appendages are fundamental for motility, biofilm formation, and efficient injection of T3SS toxins into eukaryotic cells as this process requires cell adherence [50,51,52,53]. CHA clearly possesses functional flagellum and Type IVa pili as it is able to swim comparably to PAO1 and exhibits even better twitching motility ( Figure 6A). As expected, rsmA mutation in CHA resulted in a decrease in the two motilities. Introduction of a functional copy of gacS in the chromosome of CHA did not substantially change its swimming motility, and only twitching was affected and down-regulated in our laboratory conditions ( Figure 6A). Thus, unlike frequently reported for CF-adapted strains [7], CHA is a  motile CF isolate, with functional flagellum and Type IVa pili that may contribute to its cytotoxic phenotype.
Since synthesis of Pel and Psl exopolysaccharides is known to be under the control of the Gac/Rsm post-transcriptional cascade and to mediate biofilm formation [25,53], we examined the ability of CHA and different mutants to form biofilm and, besides constitutive alginate production, synthesize Pel exopolysaccharide. CHA did not form biofilm rings in static growth conditions and inactivation of rsmA gene in CHA led to a hyperbiofilm phenotype ( Figure 6B). Introduction of a wild-type gacS gene in trans restored the ability of CHA to produce biofilm and triggered an 8.3 increase in pelA transcript level ( Figure 5), while it further enhanced biofilm production in a CHADretS strain ( Figure 6B).
Interestingly, different from the blue-green color of PAO1, the CHA strain exhibits a reddish color on PIA plate, likely resulting from a diffusible molecule ( Figure 6C). This phenotype was not due to pyomelanin production, since inactivation of the hpd gene encoding the enzyme synthesizing HGA pyomelanin precursor [54] did not affect pigmentation (not shown). The production of this pigment is clearly under control of the Gac/Rsm pathway, since inactivation of rsmA abolished the red pigmentation to a level comparable to the one observed when wild-type gacS was introduced in a CHADretS background ( Figure 6C). Complementation of CHA with gacS gene affected the phenotype slightly. These data indicated that production of this red pigmentation, similar to T3SS, should escape from the control of Gac/RsmYZ during chronic infection. It is known that RsmA controls synthesis of pigments like pyocyanin [22,55], and P. aeruginosa is able to synthesize other phenazines, as two red pigments called aeruginosins A and B, whose biological relevance is still not known [56]. Therefore the red color may rely on specific combination and quantities of these pigments. It is important to point out that phenazines are pigments with redox properties that contribute to bacterial pathogenicity in both acute and chronic models of infection. As identity and amount of phenazines are crucial for toxicity as well as for biofilm formation ( [57] and references herein), as illustrated for the precursor of pyocyanin called 5MPCA which is more efficient for yeast killing than pyocyanin itself [56], identifying the nature of the Gac/Rsm pathwayregulated pigment(s) produced by CHA could give new insights in the physiological impact of this pigmentation.
All these data indicated that a deletion in gacS gene shapes pathogenic ability of the CHA clinical isolate by blocking the Gac/ Rsm pathway. Indeed, CHA is a mucoid and motile strain expressing acute virulence factors that has lost the ability to switch from acute to chronic lifestyle.

Discussion
Particular environmental conditions found in chronic infection, such as in CF, force P. aeruginosa to adapt in order to survive. In this work, by combining genomic and extensive phenotypic analyses, we found that an intrinsic genomic deletion in the CF strain CHA equips bacteria with virulence factors that are considered to be more aggressive and are in general associated  with acute infections. Indeed, one of the key regulatory pathways, the RetS/LadS/Gac/Rsm pathway, proposed to control the transition between acute and chronic infection-associated phenotypes [25,26], is impaired by a genomic deletion that affects the gacS gene region leading to an unstable truncated form of GacS. The same signalling pathway was found affected by mutations either in ladS [58] or in retS [59] in two other clinical strains isolated from acute and chronic infection, respectively. However, inactivation of GacS, the master regulator of this pathway, impacts more strongly the read-outs than that of LadS and RetS, by blocking the switch orchestrated by the Gac/Rsm pathway.
Spontaneous mutations in gacS and/or gacA represent the most important mechanism responsible for phenotypic variation of many pseudomonads, as observed in rhizosphere-associated Pseudomonas [60]. Pseudomonas sp. strain PCL1171 undergoes phase variation which is characterized by different production of secondary metabolites and exoenzymes. This phenotypic variation was reported to be caused by spontaneous and reversible mutations in gacS and gacA genes, probably randomly generated by an inefficient MutS-dependent repair of replication-related mismatches [60,61]. One of these gacS mutations, a 307 bp deletion event, implied beforehand a spontaneous mutation that created a perfect 10 bp-direct tandem repeat allowing the recombination rearrangement [61]. A similar mechanism might have occurred in CHA genome to generate the deletion between the two 11-tandem imperfect repeats (59-CGGCCTGCCA/GG) flanking the 39 end of the gacS gene and the 59 end of the ldhA gene in PAO1 genome ( Figure 5A).
Numerous P. aeruginosa phenotypic variants are retrieved from the CF lungs, as the genome constantly accumulates mutations [1,7] and even short-term growth in biofilm was shown to generate high genetic diversity in P. aeruginosa communities [62]. Interestingly, gacS mutants in PA14 are prone to generate stable SCVs when growing in biofilm or exposed to stresses [63] as well as in vivo [64]. When CHA was grown in static conditions in vitro, we also observed the emergence of stable SCV-like colonies (not shown). These observations suggest that the absence of GacS might have conferred to the CHA bacterium further advantage for persistence in CF lung by providing it with the capacity to convert to stress-tolerant SCVs.
Clone CHA strains were isolated worldwide from rivers in Germany, soil in Japan, and from several CF patients in Central Europe [15]. In addition to the original strain used in this study, two other clone CHA strains, one isolated from the environment and another from the sputum of a CF patient with normal lung function, were sequenced [48]. Interestingly, only the herecharacterized CHA strain harbors a gacS deletion that affects so strongly its virulence properties. Genome examination did not highlight any other genetic event that could explain such deregulation [48]. This indicates that strains of the same clonal group isolated from diverse environments do not necessarily share the same infective properties [48]. Analysis of numerous CF strains with the same T6SS (absence)/T3SS (presence) pattern of expression will help to determine whether this genetic mechanism of turning a bacterium to hypervirulence is an isolated phenomenon or if it is a widespread alternative among CF lung isolates.
The CHA strain over-expresses alg genes and overproduces alginate [16]; this mucoid phenotype is due to one mutation in mucA leading to replacement of Ala-5 residue by Gly in the antisigma MucA protein [48] (Figure S2). Several reports indicated that mucA mutation negatively affects flagellum motility [11,65], leads to a reduced expression of T3SS [66] as well as of other traits of acute virulence like elastase production (T2SS) [67], suggesting that mucoidy correlates with reduced virulence [68,69]. However, despite a mucoid phenotype, CHA is endowed with efficient T3SS and also motility appendages that may contribute to its aggressive phenotype. Two mechanisms coupling mucA mutation to reduced T3SS gene expression have been reported, one dependent and the other independent of the regulator Vfr [47,68]. Indeed, the cAMP/Vfr-dependent signalling (CVS) pathway, known to activate synthesis of the virulence factors associated with acute disease such as the T3SS, was shown to be turned-off in mucAmutant strains by a mechanism involving AlgU and AlgR; this pathway thus activates alginate synthesis and reduces T3SSdependent virulence [68]. However, for a still unknown reason, inactivation of vfr does not affect T3SS in CHA (B. Toussaint, personal communication); this is surprising in regard to the high induction of CHA T3SS expression in response to calcium depletion, known to induce cAMP synthesis and, consequently, the CVS pathway [70]. Hence, neither the Gac/Rsm pathway nor the CVS pathway modulates the highly active T3SS in this isolate. The second mechanism linking mucoidy to T3SS expression implies a Vfr-independent pathway, in which mucA inactivation triggers RsmYZ transcription through activation of AlgZ/AlgR TCS, in a mechanism requiring GacS/GacA TCS [47]. In CHA strain, in which both mucA and gacS genes are inactive, this particular regulatory pathway can not control T3SS gene expression. Besides these pathways, other elements influence expression of virulence factors involved in acute and persistent infections, such as the second messenger c-di-GMP, which has been reported to inversely control T6SS and T3SS expression [71]. Interestingly, the c-di-GMP dependent-switch is linked, at an unknown level, to the RetS/Gac/Rsm pathway and requires the regulatory sRNAs. As no efficient amounts of sRNAs are synthesized in CHA, the role of c-di-GMP in the regulation of virulence factors of CHA requires additional analyses. Finally, a recent finding has added another level of complexity to the RetS/ Gac/Rsm pathway. Indeed, a new member of the RsmA/CsrA family, called RsmF/RsmN, has been identified in P. aeruginosa [72,73]. This new post-transcriptional regulator controls some of RsmA targets, notably T3SS and H1-T6SS mRNA, and its expression is negatively controlled by RsmA. Assessing its role in the virulence pattern of the CHA strain would be another challenge.
To conclude, these data clearly establish that adaptation in CF lungs generates clones with profound deregulation in the intertwined pathways controlling pathogenicity. Their study can allow identifying the different molecular links connecting the essential regulatory cascades and, thus, new therapeutic targets. Figure S1 Survival rates and bacterial dissemination in CHA-infected mice. Acute pneumonia was provoked in mice by nasal instillation of a bacterial suspension (5610 6 CFU) of either CHA or PAO1. The reference PAO1 strain was provided by A. Rietsch as PAO1F (A) Kaplan-Meyer survival curves were established from 10 infected mice per strain. Statistical differences were calculated with LogRank test. (B) Mice were euthanized 15 hours post-infection; blood and spleen were withdrawn and P. aeruginosa CFU were determined in each tissue. Data represent the mean CFU + SEM calculated for total tissue (n = 5 mice per strain). Statistical differences between strain dissemination: p = 0.009 (*) in blood and spleen as established by Mann-Whitney test.

Supporting Information
(TIF) Figure S2 The mucoid phenotype of CHA is complemented by a functional mucA copy. CHA strains containing either pJN105 (empty vector) or pJN-MucA were plated on PIA plates containing gentamycin (400 mg/ml) and 0.2% arabinose, as indicated, for 16 h at 37uC. (TIF) Materials and Methods S1 Used for generating Figures S1 and S2. (DOC)


s2fieldsofstudy:
Biology
Agricultural And Food Sciences

text:
Molecular characterization of African swine fever virus from domestic pigs in northern Tanzania during an outbreak in 2013

African swine fever (ASF) is an acute, highly contagious and deadly viral hemorrhagic fever of domestic pigs caused by African swine fever virus (ASFV), a double-stranded DNA virus of the family Asfarviridae. In this study, molecular diagnosis and characterization of outbreak ASFV in northern Tanzania, was performed on spleen, lymph node, kidney, and heart samples collected in June and July 2013 from domestic pigs that died during a hemorrhagic disease outbreak. Confirmatory diagnosis of ASF was performed using polymerase chain reaction (PCR) by partial amplification of B646L gene of ASFV encoding the major capsid protein p72 using PPA1/PPA2 primers. PCR using PPA1/PPA2 primers produced an expected PCR product size, confirming ASF outbreak in northern Tanzania. In addition, nucleotide amplification and sequencing, and phylogenetic reconstruction of the variable 3′-end of the B646L gene and complete E183L gene encoding the inner envelope transmembrane protein p54 showed that the 2013 outbreak ASFV from northern Tanzania were 100 % identical and clustered into ASFV B646L (p72) and E183L (p54) genotype X. Furthermore, the tetrameric amino acid repeats within the central variable region (CVR) of the B602L gene coding for the J9L protein had the signature BNBA(BN)5NA with a single novel tetramer NVDI (repeat code N). The results of the present study confirm an ASF outbreak in northern Tanzania in the year 2013 and show that the present outbreak ASFV is closely related to other ASFV from ticks, warthogs, and domestic pigs previously reported from Tanzania.

Introduction
African swine fever (ASF) is an acute, highly contagious and deadly viral hemorrhagic fever of domestic pigs caused by African swine fever virus (ASFV) (Costard et al. 2013). ASFV is an enveloped double stranded DNA virus classified into the family Asfarviridae (Dixon et al. 2005). Depending on the ASFV strain, ASF morbidities and mortalities can reach 100 %, making ASF the most serious constraint for domestic pig production, food security, and livelihood (Penrith 2009). Although ASF was first described in Africa and is still endemic in many African countries, devastating transcontinental spread to Asian, European, and South American countries has occurred (Costard et al. 2013;Rowlands et al. 2008).
In eastern and southern African countries, ASF is naturally maintained in a sylvatic cycle involving warthog Phacochoerus africanus and the soft argasid tick Ornithodoros moubata, and transmission to domestic pigs leading to outbreaks is either through infected tick bites, feeding contaminated warthog carcasses or contact with warthog feces (Costard et al. 2009(Costard et al. , 2013. Because of the involvement of this sylvatic cycle in the emergence of ASF, most outbreaks have been reported to start in the vicinity of National Parks (Penrith et al. 2007;Costard et al. 2013;Okoth et al. 2013). Transboundary spread of ASF during outbreaks within eastern Africa is mainly attributed to horizontal transmission between pigs due to uncontrolled pig and pig products movements, swill feeding, and lack of biosecurity measures (Wambura et al. 2006;Misinzo et al. 2011Misinzo et al. , 2012b.
After the isolation of ASFV in O. moubata ticks in the Serengeti National Park by Plowright et al. (1969), sporadic ASF outbreaks have been reported to occur at irregular intervals spanning several years (Wilson and Swai 2013). Since 2000, the frequency of reported ASF outbreaks in Tanzania has increased. Sporadic ASF outbreaks were reported in different regions including Dar es Salaam and Mbeya in 2001, Arusha in 2003, Kigoma in 2004, Mwanza in 2005and Morogoro, Coast, and Dar es Salaam in 2008and Arusha in 2009(Wambura et al. 2006Misinzo et al. 2011Misinzo et al. , 2012a. In 2010, an ASF outbreak was reported in Kyela of the Southern highland zone region of Mbeya that later on spread to other Southern highland zone regions of Rukwa and Iringa and Morogoro and Dar es Salaam regions of eastern Tanzania (Misinzo et al. 2012b). The 2010 ASF outbreak persisted until 2013 decimating pig stocks in the Southern highland zone, indicating the shift from sporadic to endemic occurrence of ASF in Tanzania (Sikombe 2013). The presence of asymptomatic pigs infected by virulent ASFV, possibly due to breedrelated resistance (Uttenthal et al. 2013), may be one of the factors that contribute to the observed persistence of ASF in the Southern highland zone. The course of ASF may vary from acute, subacute, or chronic forms depending on the virulence of ASFV, host factors, and immunological status of pigs (Gómez-Villamandos et al. 2013). All ASF outbreaks in Tanzania reported between 2000 and 2013 are related to highly virulent ASFV strains that cause the acute form of ASF accompanied with high mortality rates (Wambura et al. 2006;Misinzo et al. 2011Misinzo et al. , 2012b. Several virological tests are currently available for the diagnosis of ASF by detecting live ASFV, antigen and genome, including virus isolation, enzyme-linked immunosorbent assay (ELISA), fluorescent antibody assays, polymerase chain reaction (PCR), and isothermal amplification assays (Oura et al. 2013). Molecular differentiation between ASFV primarily relies on PCR amplification and nucleotide sequencing of the variable 3′-end of the B646L gene encoding the major capsid protein p72 (Bastos et al. 2003;Boshoff et al. 2007;Lubisi et al. 2007). Twenty two different genotypes (I to XXII) of ASFV have been identified based on nucleotide sequencing of the B646L (p72) gene (Bastos et al. 2003;Boshoff et al. 2007). The 22 ASFV the B646L (p72) genotypes can also be distinguished using microarray (Leblanc et al. 2013). Further subtyping of ASFV B646L (p72) genotypes into subgroups is achieved by nucleotide sequencing of the complete E183L gene encoding the inner envelope transmembrane protein p54 and analysis of the tetramer amino acid repeats within the hypervariable central variable region (CVR) of the B602L gene coding for the J9L protein (Irusta et al. 1996;Phologane et al. 2005;Nix et al. 2006;Lubisi et al. 2007;). Genotype I ASFV is confined to Europe, South America, the Caribbean, and West Africa whereas viruses belonging to all the 22 known ASFV genotypes have been restricted to southern and eastern Africa (Costard et al. 2013). However, genotype IX ASFV has been described in western Africa (Gallardo et al. 2011a) and genotype II ASFV have spread to Madagascar, the Caucasus region, and Russian Federation Costard et al. 2013). In East and Southern Africa, some ASFV genotypes are countryspecific, while others have transboundary distributions (Costard et al. 2009). Tanzanian outbreak ASFV belong into B646L (p72) genotypes II, IX, X, XV, and XVI (Lubisi et al. 2005;Misinzo et al. 2011Misinzo et al. , 2012aUttenthal et al. 2013).
In the present study, we report a 2013 deadly outbreak of ASF in domestic pigs in northern regions of Arusha and Kilimanjaro, Tanzania. Furthermore, we report the diagnosis and molecular characterization of 2013 outbreak ASFV in northern Tanzania based on partial amplification of the p72 gene using PCR and nucleotide sequencing of the variable 3′end of the B646L gene, the complete E183L gene, and analysis of the tetramer amino acid repeats within the CVR. Our results indicate that the 2013 northern Tanzania outbreak ASFV belongs to B646L (p72) and E183L (p54) genotype X, has a unique CVR signature BNBA(BN) 5 NA including a single novel tetramer NVDI (repeat code N) and is very closely related to other ASFV previously described from ticks, warthogs, and domestic pigs in Tanzania.

Study area and sampling
This study was conducted in the northern Tanzania regions of Arusha and Kilimanjaro after a reported hemorrhagic disease outbreak in domestic pigs with clinical and gross morphological presentations suggestive of ASF. Spleen, lymph node, kidney, and heart were collected from dead pigs from different farms in Moshi city (the ASFV strain is identified as TAN/13/ Moshi), Rombo town (TAN/13/Rombo), Nshara village of Machame ward in Hai district (TAN/13/Machame), and Arusha city (TAN/13/Arusha) during postmortem examinations ( Fig. 1). Tissues were collected and placed in separate containers from a total of 13 pigs from Moshi city (n=2), Rombo town (n=4), Machame ward (n=3), and Arusha city (n=4). Samples were transported on ice and reached the laboratory in Morogoro within 24 h. Upon arrival, 1 g from each of the spleen, lymph node, kidney, and heart belonging to the same pig were pooled into sterile petri dishes and chopped using sterile scalpel blades after addition of 5:1 (v/v) sterile phosphate buffer saline (PBS) filtered through a 0.22-μm Minisart syringe filter (Sartorius Stedim Biotech, Goettingen, Germany). Afterwards, homogenized tissue samples were centrifuged at 6,000g for 5 min and the supernatants aliquoted into cryovials before being stored at −80°C.

DNA extraction and amplification
Frozen aliquots of pooled spleen, lymph node, kidney, and heart tissue homogenates were thawed and DNA extraction performed using a QiaAmp nucleic extraction kits (Qiagen, Hilden, Germany), following the manufacturer's instructions. During nucleic acid extraction and PCR, the homogenized tissue suspension containing the TAN/12/Ifakara (Misinzo et al. 2012b) was used as positive control and PBS as negative control. PCR detection of ASFV was performed using primers PPA1 and PPA2 that partially amplify the B646L gene encoding the major capsid protein p72, as previously described by Aguero et al. (2003). The expected size of the PCR product using primers PPA1 and PPA2 primers is 257 bp. PCR for the molecular characterization of ASFV was performed to amplify the (i) variable 3′-end of the B646L gene encoding the major capsid protein p72 using primers p72U and p72D, (ii) complete E183L gene encoding the inner envelope transmembrane protein p54 using primers PPA89 and PPA722, and (iii) the tetramer amino acid repeats within the hypervariable CVR of the B602L gene using ORF9L-F and ORF9L-R primers, as previously described by . All PCR amplifications were performed using AccuPower PCR premix (Bioneer, Daejeon, Republic of Korea) on a GeneAmp PCR systems 9700 (Applied Biosystems, Foster City, CA). Afterwards, PCR products were electrophoresed in a 2 % agarose gel mixed with GelRed nucleic acid stain (Phenix Research Products, Candler, NC) before visualization and imaging using a BioDoc-It imaging system (UVP, Upland, CA).
Nucleotide sequencing PCR products from B646L (p72), E183L (p54), and CVR were purified from agarose gels using a NucleoSpin gel and PCR clean-up kit (Macherey-Nagel, Düren, Germany) and subjected to dideoxynucleotide cycle sequencing by using Big Dye Terminator Cycle Sequencing Kit Version 3.1 (Applied Biosystems, Foster City, CA). Products from dideoxynucleotide cycle sequencing reaction were purified by ethanol precipitation and separated on a 3500 Genetic Analyzer (Applied Biosystems, Foster City, CA). Chromatograms for both the forward and the reverse primer reactions were read using Sequence Scanner v1.0 software (Applied Biosystems, Foster City, CA). The sequence from the forward primer and the reverse complement sequence of the reverse primer were manually overlapped using a text editor to obtain a single consensus sequence delimited by the forward and reverse primers.
Molecular characterization of ASFV using the B646L (p72), E183L (p54), and CVR approach The nucleotide sequence of B646L (p72), E183L (p54), and CVR from the 2013 ASF outbreak in northern Tanzania were submitted to GenBank and given accession numbers. The similarity search of the obtained nucleotide sequences against other ASFV sequences at GenBank database was performed using BLASTn (version 2.2.29). BLASTn compares nucleotide sequences to sequence databases and calculates the statistical significance of matches. The nucleotide sequences of the B646L (p72), E183L (p54), and CVR genes of northern Tanzanian 2013 outbreak ASFV were aligned with other Tanzanian ASFV nucleotide sequences available at GenBank (Table 1) using ClustalW algorithm in BioEdit (Ibis Biosciences, Carlsbad, CA). The clustering pattern of ASFV was determined by neighbor-joining method using the Kimura-2-parameter option implemented within MEGA 5 (Tamura et al. 2011). Phylogeny was inferred following 1,000 bootstrap replications. The CVR nucleotide sequences of the northern Tanzanian 2013 outbreak ASFV were translated and coded to obtain signatures based on previously reported codes (Nix et al. 2006;Boshoff et al. 2007;Misinzo et al. 2011). A similarity search against other ASFV amino acid sequences was performed using BLASTp (v2.2.2.29).

Epidemiology of ASF in Arusha and Kilimanjaro
ASF was first reported in Tarakea (Rombo district), located at the border between Tanzania and Kenya in May 2013 (Fig. 1). Afterwards, ASF spread from Tarakea to neighboring Useri and Mashati wards of Rombo district before it was reported in Arusha city on 06 June 2013 in villages surrounding the Arusha dampo, including Muriet and Engosengiu of Sokoni One ward. On 18 June 2013, ASF was reported in a number of districts including Moshi Rural, Moshi Urban (Mbokomu ward), Arusha, and Arumeru (Sekei and Olga Lai wards) districts (Fig. 1). Between July and August 2013, ASF had spread to Korogwe and Hai (Machame ward) districts. The number of recorded domestic pig death due to ASF is shown in Table 2. By September 2013, there were no reports on new cases. In Rombo and Arusha districts, ASF spread as a result of feeding pork leftovers from hotels, restaurants, and homes.
The spread between the different districts is attributed to introductions of infected animals bought at a cheaper price from ASF areas under quarantine due to ineffective law enforcement. The main clinical signs presented by the sick domestic pigs in the visited farms included hind leg weakness, recumbence, dyspnea, anorexia, and erythema and cyanosis of the skin (Fig. 2). Domestic pigs of all ages were affected although severe clinical signs were observed in adults than piglets. Abortion was frequently observed in pregnant sows. At postmortem, strawcolored or blood-tinged fluid was observed in the pleura, pericardium, and peritoneal cavities. In addition, splenomegaly, petechiations of the heart and kidney, and hemorrhagic heart, kidneys, liver, intestines, and lymph nodes especially the mesenteric and the hepatogastric lymph nodes were observed (Fig. 2).

Confirmatory diagnosis of ASFV using PCR
An ASF diagnostic PCR previously reported by Aguero et al. (2003) was used to confirm the presence of ASF in samples collected from dead pigs in Moshi, Rombo, Machame, and Arusha. With the exception of tissues from one pig in Arusha, all pooled tissues belonging to 12 pigs from Moshi city (n=2), Rombo town (n=4), Machame ward (n=3), and Arusha city (n=3) were positive. The PCR products of ASFV from Moshi, Rombo, Machame, and Arusha obtained after performing diagnostic PCR using PPA1/2 primers were 257 nucleotides long (Misinzo et al. 2014).
Molecular characterization of ASF based on the B646L (p72), E183L (p54), and CVR approach Nucleotide amplification and sequencing of the variable 3′end of the B646L gene encoding the major capsid protein p72, complete E183L gene encoding the inner envelope transmembrane protein p54, and the hypervariable CVR of the   (Misinzo et al. 2012a). When phylogenetic analysis of the Tanzanian ASFV B646L (p72) nucleotide sequences was performed together with the nucleotide sequences obtained from this study, the 2013 ASFV from Moshi, Rombo, Machame, and Arusha clustered together with ASFV belonging to genotype X (Fig. 3). The 2013 ASFV from Moshi, Rombo, Machame, and Arusha also clustered into genotype X when phylogenetic analysis was performed using the E183L (p54) nucleotide sequences (data not shown).
The CVR nucleotide sequences of the northern Tanzanian 2013 outbreak ASFV were translated and coded to obtain their signatures. ASFV tetrameric amino acid repeats within CVR that have previously been reported in ASFV include CAST/CASI/CVST/CTST (repeat code A), CADT/CTDT  while an ear with marked cutaneous congestion (indicated with an arrow head) in pigs with ASF is shown in (d). In addition, pigs with ASF showed hemorrhages (indicated by an arrow head) of the intestines (e), kidneys (f), spleen (g), and mesenteric lymph nodes (h). g Enlargement of the spleen (splenomegaly) was observed in pigs with ASF

Discussion
In this study, the molecular diagnosis and characterization of ASFV from domestic pigs that died of a hemorrhagic disease outbreak between May and August 2013 in northern Tanzania was performed. The results obtained from the present study confirm ASF outbreak in Arusha and Kilimanjaro regions located in the northern Tanzania. ASFV in domestic pigs was confirmed by nucleotide amplification, sequencing, and phylogenetic reconstruction of ASFV B646L (p72) and E183L (p54) genes, and translation of the B602L (CVR) gene. The obtained B646L (p72), E183L (p54), and B602L (CVR) nucleotide sequences of 2013 outbreak ASFV from northern Tanzania were 100 % identical and clustered into ASFV B646L (p72) and E183L (p54) genotype X. Furthermore, the tetrameric amino acid repeats within the CVR of the B602L gene had the signature BNBA(BN) 5 NA including a single novel tetramer NVDI (repeat code N).
The present ASF outbreak started in May 2013 in Kilimanjaro region before spreading to Arusha region in June 2013. By September 2013, no new cases of ASF were recorded in Kilimanjaro and Arusha. The sporadic nature and short longevity of the 2013 ASF outbreak is comparable to previous outbreaks of 2001, 2003, 2004, 2005, 2008, and 2009 that lasted for a few months before resolving (Wambura et al. 2006;Misinzo et al. 2011Misinzo et al. , 2012a. In Tanzania, the only recorded outbreak that persisted longer was the ASF outbreak between 2010 and 2013 that started in Mbeya region and later on spread to Iringa, Rukwa, Dar es Salaam, and Morogoro regions (Misinzo et al. 2012b;Sikombe 2013). The 2010 to 2013 ASF outbreak was caused by a B646L (p72) genotype II ASFV that had previously not been described in Tanzania and eastern Africa (Misinzo et al. 2012b;Sikombe 2013). The B646L (p72) and E183L (p54) genes of this Tanzanian genotype II ASFV are identical to the Georgia 2007/1 virus and clusters into this genotype with other ASFV from Mozambique, Madagascar, and Mauritius (Misinzo et al. 2012b;Uttenthal et al. 2013). Before the genotype II introduction, Tanzanian outbreak ASFV clustered into B646L (p72) genotypes IX in 2005, X in 2004, XV in 2001, and XVI in 2003(Lubisi et al. 2005Wambura et al. 2006;Misinzo et al. 2011Misinzo et al. , 2012a. The ASFV identified in this study clusters into B646L (p72) genotype X, similar to other viruses that have caused ASF outbreak in Tanzania in 2009 in Longido (northern Tanzania), 2004 in Kigoma (western Tanzania), and other ASFV recovered from ticks and warthogs in Serengeti National Park (northern Tanzania) (Lubisi et al. 2005;Wambura et al. 2006;Misinzo et al. 2012a). Genotype X is a sylvatic cycle-associated genotype that comprises ASFV recovered from domestic pigs, warthogs and ticks in Burundi, Tanzania, and Kenya (Bastos et al. 2003;Lubisi et al. 2005;Lubisi et al. 2007;Gallardo et al. 2011b).
The Tanzanian 2013 ASFV reported in the present study originated from Tarakea, a town bordering Kenya and very close to the Kilimanjaro National Park. In addition, the Tanzanian 2013 ASFV is closely identical to the 2009 outbreak ASFV (TAN/09/Longido) from another town located within northern Tanzania. Alignment of 444 nucleotide long sequence of the variable 3′-end of the B646L (p72) gene of Tanzanian 2013 with 2009 ASFV show only a single nucleotide substitution (A→T). The close nucleotide identity between 2013 and 2009 outbreak ASFV indicates the presence of closely related viruses in northern Tanzania. Northern Tanzania has several wildlife protected areas (Fig. 1). ASFV diversity is generated during the sylvatic cycle of the virus (Jori et al. 2013). There is a possibility that viruses evolving in these wildlife protected areas spill over to the Fig. 3 Neighbor-joining tree depicting partial B646L (p72) gene relationships of African swine fever viruses from Tanzania. Phylogeny was inferred following 1,000 bootstrap replications, and the node values show percentage bootstrap support. Scale bar indicates nucleotide substitutions per site. The Genbank accession numbers for the different B646L (p72) genes are indicated in parenthesis domestic pig population causing outbreaks such as the 2009 and 2013 outbreaks. It will be important to sequence the E183L (p54) and B602L (CVR) genes of TAN/09/Longido ASFV in order to understand the similarity between this virus and the 2013 viruses at these genomic locations. In addition, it will be important to perform the surveillance and genetic characterization of ASFV from warthogs and ticks found in Tanzanian National Parks in order to understand the relationship between sylvatic and outbreak ASFV.
To date, the CVR region has only been analyzed for the 2008 Tanzanian outbreak ASFV (Misinzo et al. 2011). The tetrameric amino acid repeats within the CVR of the B602L gene of the 2013 Tanzanian ASFV reported in this study had the signature BNBA(BN) 5 NA, which is distinct from the 2008 Tanzanian ASFV that had the signature AVUA VUVAVVUAVUVAVUVAVVUAVVUUUXV (Misinzo et al. 2011). BLASTp (v2.2.29) analysis using the deduced amino acid sequence showed 75 % overall amino acid identity with the CVR of Mozambican viruses SPEC/265 and Moz 94/ 1, that belong to genotype VI (Bastos et al. 2004;Nix et al. 2 0 0 6 ) . T h e C V R s i g n a t u r e f o r S P E C / 2 6 5 i s AABABNABABNBABMA while that of Moz94/1 is AAAABABNABABNBTBA, indicating that the 2013 Tanzanian ASFV differ from Mozambican viruses.
Over the past two decades, the frequency of ASF outbreaks in Tanzania has increased. When the ASF outbreak in northern Tanzania (Kilimanjaro and Arusha) was reported, there was an ongoing outbreak of ASF in southern Tanzania (Mbeya, Iringa and Rukwa) (Misinzo et al. 2012b). Northern and southern Tanzania are the major pig-producing zones in Tanzania with highest pig population in Mbeya followed with Iringa, Ruvuma, and Kilimanjaro region (URT 2012). Pig husbandry has recently increased, for instance in Rombo district (Kilimanjaro region), pig farming has expanded as source of income to many households replacing coffee farming. The ASF outbreaks in southern and northern Tanzania has decimated the pig population affecting the food security and livelihoods of poor pig farmers.
It can be concluded from the results obtained in this study that the ASFV that caused the 2013 outbreak in Arusha and Kilimanjaro regions in northern Tanzania are related to other previously reported B646L (p72) genotype X eastern African sylvatic cycle viruses and differ from the genotype II ASFV reported in southern Tanzania. Furthermore, unrestricted pig movements and swill feeding continues to contribute to the spread of ASF outbreaks within Tanzania. The impact of ASF on the livelihood of farmers resulting from higher frequency and increased durations of ASF outbreaks in Tanzania is worth investigating.


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Comparison of 2 synthetically generated recombinant prions

Prion is a protein-conformation-based infectious agent causing fatal neurodegenerative diseases in humans and animals. Our previous studies revealed that in the presence of cofactors, infectious prions can be synthetically generated in vitro with bacterially expressed recombinant prion protein (PrP). Once initiated, the recombinant prion is able to propagate indefinitely via serial protein misfolding cyclic amplification (sPMCA). In this study, we compared 2 separately initiated recombinant prions. Our results showed that these 2 recombinant prions had distinct biochemical properties and caused different patterns of spongiosis and PrP deposition in inoculated mice. Our findings indicate that various recombinant prions can be initiated in vitro and potential reasons for this variability are discussed.

Introduction
Prion is a protein-conformation-based infectious agent that causes transmissible spongiform encephalopathies (TSEs, also known as prion disease) in animals and humans. 1,2 The prion hypothesis posits that the pathogenic conformer of host encoded prion protein (PrP) is able to convert host PrP to the pathogenic conformation. 1,3 It is well established that the pathogenic PrP Sc conformer is β-sheeted, aggregated, and protease-resistant, which distinctly differs from the α-helical, soluble, and protease-sensitive normal PrP C conformer. 4 The most widely used biochemical property separating these 2 PrP conformers is the sensitivity to proteinase K (PK) digestion. PrP C is sensitive to PK digestion, whereas the C-terminus of PrP Sc is highly resistant to PK digestion resulting in a characteristic C-terminal PK-resistant core. 4,5 The association of the PK-resistant PrP Sc conformer and prion infectivity was established by the development of serial protein misfolding cyclic amplification (sPMCA) technique, which consists of alternating cycles of sonication and incubation. [6][7][8] Using normal brain homogenate as substrate for sPMCA, Castilla et al. demonstrated that PrP Sc is able to seed PrP C conversion to the PK-resistant PrP Sc conformation and more importantly, the prion infectivity is propagated along with the propagation of PrP Sc conformers. 8 Recently, the same sPMCA technique has been successfully applied to propagate the pathogenic PrP Sc conformer and prion infectivity using a variety of native or recombinant PrP substrates. [9][10][11][12][13][14][15][16][17][18] Although sPMCA was originally developed as a technique to propagate the pathogenic PrP Sc conformer, 6,7 several studies showed that it also has the capability to initiate the pathogenic PrP Sc conformation de novo in unseeded reactions. 9,11,15,19 Using sPMCA with a substrate system consisting of purified bacterially-expressed recombinant murine PrP (rPrP) plus synthetic phospholipid POPG (1-palmitoyl-2-oleoylphosphatidylglycerol) and total RNA isolated from normal mouse liver as cofactors, we previously reported de novo formation of a PK-resistant and highly infectious prion in the laboratory of Ohio State University (named as rPrP-res OSU ). 9 To confirm this result, we repeated the same experiment in a newly established lab in East China Normal University that has never been exposed to any naturally occurring prion. An infectious prion with a 17 kDa PK-resistant core was generated de novo during this attempt, which was named as rPrP-res 17kDa . 11 Both rPrP-res OSU and rPrP-res 17kDa have the characteristic self-perpetuating C-terminal PK-resistant conformation Abbreviations: TSEs, transmissible spongiform encephalopathies; PrP, prion protein; PrP Sc , the pathogenic conformational isoform of prion protein; PrP C , the normal host prion protein; PK, proteinase K; sPMCA, serial protein misfolding cyclic amplification; rPrP, bacterially-expressed recombinant prion protein;POPG, 1-palmitoyl-2-oleoylphosphatidylglycerol; rPrP-res, proteinase K resistant recombinant PrP conformer; dpi, days post inoculation; FC, frontal cortex;CN, caudate nucleus;CWM, cerebellum white matter; PET blot, paraffin-embedded tissue blot; IHC, immunohistochemical; GuHCl, guanidine hydrochloride; TBST, trisbuffered saline buffer with 0.1% Tween-20; BSA, bovine serum albumin; PBS, phosphate buffered saline Prion is a protein-conformation-based infectious agent causing fatal neurodegenerative diseases in humans and animals. Our previous studies revealed that in the presence of cofactors, infectious prions can be synthetically generated in vitro with bacterially expressed recombinant prion protein (PrP). Once initiated, the recombinant prion is able to propagate indefinitely via serial protein misfolding cyclic amplification (sPMcA). In this study, we compared 2 separately initiated recombinant prions. Our results showed that these 2 recombinant prions had distinct biochemical properties and caused different patterns of spongiosis and PrP deposition in inoculated mice. Our findings indicate that various recombinant prions can be initiated in vitro and potential reasons for this variability are discussed. and cause fatal prion disease in wild-type mice with 100% attack rate. 9,11 Mice inoculated with rPrP-res OSU or rPrP-res 17kDa survived for 150 or 172 dpi (days post inoculation) respectively, 9,11 which are in the range of survival times for naturally occurring mouse prions. To determine the similarity and/or difference between these 2 separately initiated recombinant prions, we compared the neuropathology caused by these 2 recombinant prions and their biochemical properties in this study.

Results
Pathological change in prion disease is characterized by the presence of vacuoles (spongiosis) in diseased brains. Both rPrP-res OSU and rPrP-res 17kDa caused wide spread spongiosis in wild-type CD-1 mice. 9,11 Interestingly, the spongiosis profile in rPrP-res 17kDa -inoculated mice appeared to be different from that of rPrP-res OSU (Fig. 1A). The most prominent differences of spongiosis were observed in frontal cortex (FC) and caudate nucleus (CN). Severe spongiosis was observed in both FC and CN areas of rPrP-res OSU -inoculated mice, but only mild spongiosis was detected in these 2 areas of rPrP-res 17kDa -inoculated mice (Fig. 1A). The severity of spongiosis in other brain areas, such as cerebellum white matter (CWM), was very similar (Fig. 1A).
To determine whether rPrP-res 17kDa and rPrP-res OSU led to different aberrant PrP deposition patterns in the brain, we performed paraffin-embedded tissue (PET) blot and immunohistochemical (IHC) staining. The PET blot revealed a drastic difference with rPrP-res 17kDa (17kDa) or rPrP-res OsU (OsU) as indicated. P, pons; MB, middle brain; cWM, cerebellum white matter; hyp, hypothalamus; Tha, thalamus; hip, hippocampus; cN, caudate nucleus; Fc, frontal cortex. The lesion profile was based on the scores of 6 rPrP-res 17kDainoculated and 15 rPrP-res OsU -inoculated mice. Units of Y-axis are arbitrary units of lesion severity, which are described in detail in methods section. Representative images of spongiosis in frontal cortex (Fc), caudate nucleus (cN), and cerebellum white matter (cWM) are shown in (A). (B) PeT blot analysis of mice inoculated with rPrP-res 17kDa (17kDa) or rPrP-res OsU (OsU) as indicated. PeT blots were stained at the same time and using exactly the same condition. Three pairs of mice inoculated with rPrP-res 17kDa or rPrP-res OsU were subjected to PeT blot analysis and PK-resistant PrP deposition pattern was consistent within the same group. (C) Immunohistochemical staining of aberrant PrP deposit in various regions of mouse brain receiving intracerebral inoculation of rPrP-res 17kDa (17kDa) or rPrP-res OsU (OsU) as indicated. Mice receiving control inoculum (PBs + BsA) were used as controls in all panels (control). These sections were stained at the same time and using exactly the same condition. Three pairs of mice inoculated with rPrP-res 17kDa or rPrPres OsU were analyzed and the aberrant PrP deposition pattern was consistent within the same group.
in PK-resistant PrP deposition in cortex. Compared with that in rPrP-res OSU -inoculated mice, the cortical PK-resistant PrP deposition was much stronger in rPrP-res 17kDa -inoculated mice (Fig. 1B). The IHC staining confirmed the difference in PrP deposition in cortex (Fig. 1C, left panel). Moreover, IHC staining also revealed that, compared with rPrP-res OSU -inoculated mouse brains, the rPrP-res 17kDa inoculation caused a stronger PrP deposition in hippocampus, but a weaker PrP deposition in thalamus (Fig. 1C, middle and right panels). Together, neuropathological comparison revealed a clear difference between the disease pathologies caused by rPrP-res 17kDa and rPrP-res OSU .
Since the bioassays of rPrP-res 17kDa and rPrP-res OSU were performed in outbred CD-1 mice in 2 different animal facilities, the observed difference in disease pathology could be resulted from a variety of factors such as the differences in mouse genetic background. However, it is also possible that rPrP-res 17kDa and rPrP-res OSU are different rPrP conformers, which contributes to the difference in disease pathology. To test the latter possibility, we compared the biochemical properties of rPrP-res 17kDa and rPrP-res OSU .
Immunoblot analysis with POM1 antibody that recognizes an epitope at C-terminus of PrP 20,21 showed that the sizes of the PK-resistant fragments of 2 rPrP-res forms were similar, but the PK-resistant core of rPrP-res 17kDa was slightly larger than that of rPrP-res OSU ( Fig. 2A, left panel). When immunoblot analysis was performed with SAF32 antibody that recognizes the N-terminal octarepeat region, 22 a much stronger signal was detected with PK-digested rPrP-res 17kDa (+PK, Fig. 2A, right panel). The immunoblot analysis of control samples with SAF32 antibody (-PK, Fig. 2A, right panel) showed comparable intensity, indicating that there is no significant difference between the affinity of SAF32 antibody to rPrP-res 17kDa and rPrP-res OSU . Thus, a weaker signal of the PK-resistant fragment of rPrP-res OSU (+PK, Fig. 2A, right panel) suggests that rPrP-res 17kDa contains a longer PK-resistant core(s), which extends into the octarepeat region resulting in a better SAF32 antibody detection.
When the same blots were scanned by a Storm 860 PhosphorImager that holds a wider detection range than X-ray film, multiple PK-resistant bands were recognizable in both rPrP-res 17kDa and rPrP-res OSU samples (Fig. 2B). For rPrP-res OSU , 2 major PK-resistant bands were detected by POM-1 antibody and the most abundant band was the one with smallest molecular weight (Fig. 2B, indicated by an arrow), whereas for rPrP-res 17kDa , 3 major PK-resistant bands were detected by POM-1 antibody and the most abundant bands were the 2 slower migrating bands (Fig. 2B, indicated by * and ** on left panel). In addition to the band intensity, a slight difference in band size was also observed (Fig. 2B, the band with evident size difference was indicated by ** on left panel). The SAF32 antibody recognized slower migrating PK-resistant bands but not the one with smallest molecular weight (Fig. 2B, right panel), accounting for its weaker recognition of PK-resistant bands of rPrP-res OSU (Fig. 2A, right panel). Together, these results indicate that the PK cleavage sites in rPrPres are heterogeneous, which is similar to that of naturally occurring PrP Sc . 5 A possible reason for the banding pattern difference between rPrP-res 17kDa and rPrP-res OSU (Fig. 2B, left panel) could be due to an insufficient PK digestion of the rPrP-res 17kDa . To test this possibility, we performed serial PK digestion of 2 samples. We first normalized rPrP-res 17kDa and rPrP-res OSU to similar amount of total rPrP (2 controls in the lower panel of Fig. 2C); confirmed by PhosphorImager quantification (the difference between the average of raw PhosphorImager quantification scores for these 2 controls was less than 1%). After being normalized, samples were digested with increasing amounts of PK and the PK-resistant bands of rPrP-res 17kDa disappeared at PK concentration of 500 μg/mL. The rPrP-res OSU , however, showed a much higher PK-resistance and its PK-resistant fragment was clearly detectable after a 30 min 1000 μg/mL PK digestion at 37 °C (Fig. 2C). Thus, the banding pattern difference between rPrP-res 17kDa and rPrP-res OSU (Fig. 2B, left panel) is not due to insufficient PK digestion of rPrP-res 17kDa . A more plausible explanation could be that the 2 recombinant prions produce different PK-resistant bands.
Next, we performed the guanidine hydrochloride (GuHCl) denaturation assay to compare the stability of rPrP-res 17kDa and rPrP-res OSU . Figure 2D shows that the denaturation curves for rPrP-res 17kDa and rPrP-res OSU were very similar, with [GuHCl] 1/2 (the GuHCl concentration causing loss of half of the PK-resistant PrP signal) at 1.51 M for rPrP-res OSU and 1.55 M for rPrP-res 17kDa . Despite the similarity in denaturation curve, the rPrP banding patterns after GuHCl treatment were clearly different between 2 rPrP-res forms. In rPrP-res 17kDa , a smaller PK-resistant fragment became prominent after 1.5 M GuHCl treatment and remained detectable when the concentration of GuHCl was at 2 or 2.5 M (Fig. 2D, pointed by an arrow in the bottom rPrP-res 17kDa panel). In contrast, this smaller PK-resistant fragment was not clearly detected in rPrP-res OSU samples (Fig. 2D, the rPrP-res OSU panel).
Collectively, our biochemical analyses clearly showed that, despite the similar self-perpetuating PK-resistant conformation and similar capability of causing prion disease in wild-type mice, the rPrP-res 17kDa and rPrP-res OSU have distinct differences in biochemical properties.

Discussion
Our 2 successful attempts to generate de novo recombinant prions by sPMCA, particularly the latter attempt conducted in a lab that has never exposed to any naturally occurring prions, support that an infectious recombinant prion can be synthetically generated in vitro. The current comparative study of separately initiated rPrP-res OSU and rPrP-res 17kDa suggests that the 2 recombinant prions are differently misfolded rPrPs, with distinct biochemical properties and causing different neuropathological changes. The sPMCA substrate mixtures for de novo formation of rPrP-res OSU and rPrP-res 17kDa contained the same 3 components: recombinant murine PrP, POPG, and total RNA isolated from normal mouse liver. Despite the relatively simple components in the substrate mixture, many variations may lead to the differences in de novo generated, self-perpetuating PK-resistant rPrP-res forms.
First, depending on the metabolic status of the mouse, the total RNA isolated from each individual mouse liver may contain different RNA species with different three-dimensional RNA structures. Moreover, purification of total RNA is always accompanied with co-purified molecules, which may also vary according to the metabolic status of individual mouse. Therefore, despite the same reagent and protocol, the isolated total RNA may vary from mouse to mouse. Our recent study has shown that variations in RNA species or co-purified molecules are not essential for generating the rPrP-res conformation and prion infectivity. 10 However, it does not preclude the potential difference in modulating rPrP-res conformation by different RNA species or co-purified molecules.
Second, the differences in rPrP-res forms may result from different ratios of rPrP and cofactors. Thus far, it remains unclear whether cofactors are in the final rPrP-res complex. 23,24 Even if cofactors play a chaperone-like role to facilitate rPrP-res formation, different rPrP:cofactors ratios may guide rPrP to different misfolding stage and result in rPrP-res conformational differences. If cofactors are part of the final rPrP-res complex, variation in rPrP:cofactor ratios will most likely alter the final rPrP threedimensional structure. If cofactors are not essential for the selfperpetuating capability, then the final rPrP-res complex can be with or without cofactors. In this case, the presence or absence of cofactors may produce different rPrP-res conformations leading to different biological effects. Thus, despite the relatively simple sPMCA substrate system for de novo rPrP-res formation, the possible variations in rPrP:cofactors ratios are enormous, which may contribute to the formation of different rPrP-res form.
Third, the variations in rPrP-res conformation may also result from the sonication step of sPMCA. Previous studies showed that the interaction between rPrP and cofactors (POPG and RNA) in our sPMCA substrate converts rPrP to a β-sheeted, aggregated, and PK-resistant conformation similar to rPrP-res. [24][25][26][27][28][29][30][31] When this substrate is subjected to sPMCA, sonication breaks rPrP aggregates in the substrate (and/or directly alters rPrP conformation) and new rPrP aggregates form during incubation. As we suggested previously, 11 this repeated rPrP aggregate breaking-and-formation cycles may ultimately re-package rPrP into self-perpetuating rPrPres aggregates. Because of its seeding capability, once initiated, the rPrP-res aggregates can be faithfully propagated via sPMCA. If this model is correct, the power of sonication, such as different sonicators or different tube positions in the sonicator horn, will likely influence the extent of rPrP-res aggregate breakage and/or the level of rPrP conformation alteration, both of which may lead to the variations in the initial rPrP-res conformation.
A fourth explanation for the formation of 2 different forms of rPrP-res could be simply due to the adoption of different stable rPrP-res conformers. The rPrP-res is a misfolded rPrP form and there could be multiple stable rPrP-res spots in the energy landscape. 32 In this scenario, the presence of cofactors and the sPMCA process may create an environment favoring rPrP conformational change, allowing rPrP to randomly search for the stable spots in the energy landscape. Once rPrP adopts distinct stable rPrP-res conformations, different rPrP-res forms are initiated and propagated.
Any of the above reasons, or combination of them, may contribute to the de novo formation of different rPrP-res forms, which may account for the observation of our current study and the recent reports of various de novo generated non-infectious self-perpetuating rPrP-res forms. 11,33 It is important to note that although we have shown the biochemical and neuropathological differences between rPrP-res OSU and rPrP-res 17kDa , we cannot conclude that these are 2 different prion strains. This is because the mouse bioassays were performed in outbred CD-1 mice in 2 different animal vivaria. Careful serial passage in inbred mouse line is required to clarify this question. Despite the caveats, our observation that the 2 forms of rPrP-res produce heterogeneous PK-resistant rPrP species (Fig. 2B) is consistent with the quasispecies theory, 32,34 which posits that a prion strain consists of an ensemble of PrP Sc conformers, and various prion strains may differ in certain or all PrP Sc species in the group and/or the proportion of each PrP Sc species.
Results from our comparison study clearly showed that different rPrP-res forms can be formed de novo, which leads to different disease pathologies in mice. This finding supports that not a single PrP conformer, but multiple PrP conformers are capable of supporting prion infectivity.

Materials and Methods
PET blot was performed as previously described. 9,35 Briefly, 4-μm-thick paraffin sections were collected onto 0.45-μm nitrocellulose membranes (Bio-Rad Laboratories) and incubated at 55 °C for 16 h. Membranes were dewaxed by immersion in xylene (45 °C, 20 min) and rinsed in isopropanol (2 × 10 min) followed by stepwise rehydration. After washing with tris-buffered saline buffer (TBST, 50 mM TRIS-HCl, 150 mM NaCl, pH 8.0, 0.1% Tween-20), the membrane was subjected to 250 μg/mL PK digestion in a buffer consisting of 10 mM TRIS-HCl, pH 7.8, 100 mM NaCl, 0.1% Brij 35 for 16 h at 55 °C. After washing with TBST, the membrane was treated with 4 M guanidine thiocyanate for 10 min and washed 3 times in TBST. The membrane was blocked by 2% non-fat milk in TBST for 1 h, incubated with monoclonal SAF84 anti-PrP antibody (1:2000 in blocking solution) for 90 min at room temperature, washed 3 times in TBST, and incubated with an AP conjugated goat anti-mouse IgG antibody (1:3000 in blocking solution) for 1 h at room temperature. After 3 washes with TBST, the color was developed by BCIP/ NBT, and the images were assessed using a stereomicroscope.
GuHCl denaturation assay was performed by mixing 30 μL sPMCA product with 30 μL GuHCl stock solutions to reach final GuHCl concentrations ranging from 0 to 4 M. The mixtures were incubated at room temperature for 1 h and mixed by vortexing every 15 min. Afterwards, samples were kept on ice and stock solutions were added to normalize the final salt and detergent concentrations to: 0.4 M GuHCl, 0.15% Triton X-100, 16.5 mM TRIS-HCl, pH 7.4 and 7.5 mM NaCl. Normalized samples were digested with PK (10 μg/mL) for 30 min at 37 °C. Digestion was terminated by adding 1 mM PMSF and incubation on ice for 5 min. PK-resistant proteins were precipitated by adding 4 volumes of cold methanol (-20 °C) and incubated at -20 °C for 30 min. The precipitated proteins were pelleted by centrifugation, resuspended in SDS-PAGE sample buffer, subjected to immunoblot analysis, and the results were quantified by scanning with a Storm 860 PhosphorImager.
Immunoblot analysis was performed as previously described. 37 The blot was developed with ECL-plus reagent (GE Healthcare Life Science) and the signal was detected by exposure to X-ray film or scanning with a Storm 860 PhosphorImager (Fig. 2B).

Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.