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1.
The Norway spruce genome provides key insights into the evolution of plant genomes, leading to testable new hypotheses about conifer, gymnosperm, and vascular plant evolution.In the past year a burst of plant genome sequences have been published, providing enhanced phylogenetic coverage of green plants (Figure (Figure1)1) and inclusion of new agricultural, ecological, and evolutionary models. Collectively, these sequences are revealing some extraordinary structural and evolutionary attributes in plant genomes. Perhaps most surprising is the exceptionally high frequency of whole-genome duplication (WGD): nearly every genome that has been analyzed has borne the signature of one or more WGDs, with particularly notable events having occurred in the common ancestors of seed plants, of angiosperms, and of core eudicots (the latter ''WGD'' represents two WGDs in close succession) [1,2]. Given this tendency for plant genomes to duplicate and then return to an essentially diploid genetic system (an example is the cotton genomes, which have accumulated the effects of perhaps 15 WGDs [3]), the conservation of genomes in terms of gene number, chromosomal organization, and gene content is astonishing. From the publication of the first plant genome, Arabidopsis thaliana [4], the number of inferred genes has been between 25,000 and 30,000, with many gene families shared across all land plants, although the number of members and patterns of expansion and contraction vary. Furthermore, conserved synteny has been detected across the genomes of diverse angiosperms, despite WGDs, diploidization, and millions of years of evolution.Open in a separate windowFigure 1Simplified phylogeny of land plants, showing major clades and their component lineages. Asterisks indicate species (or lineage) for which whole-genome sequence (or sequences) is (are) available. Increases and decreases in genome size are shown by arrows.Despite the proliferation of genome sequences available for angiosperms, genome-level data for both ferns (and their relatives, collectively termed monilophytes; Figure Figure1)1) and gymnosperms have been conspicuously lacking - until recently, with the publication of the genome sequence of the gymnosperm Norway spruce (Picea abies) [5]. The large genome sizes for both monilophytes and gymnosperms have discouraged attempts at genome sequencing and assembly, whereas the smaller genome size of angiosperms has resulted in more genome sequences being available (Table (Table1)1) [6]. Because of this limited phylogenetic sample, our understanding of the timing and phylogenetic positions of WGDs, the core number of plant genes, possible conserved syntenic regions, and patterns of expansion and contraction of gene families across both tracheophytes (vascular plants) and across all land plants is imperfect. This sampling problem is particularly acute in analyses of the genes and genomes of seed plants; many hundreds of genes are present in angiosperms that are not present in mosses or lycophytes, but whether these genes arose in the common ancestor of seed plants or of angiosperms cannot be determined without a gymnosperm genome sequence. The Norway spruce genome therefore offers tremendous power, not only for understanding the structure and evolution of conifer genomes, but also as a reference for interpreting gene and genome evolution in angiosperms.
Open in a separate windown/a, not applicable. Data based on [6]. 相似文献
Table 1
Genome sizes in land plantsLineage | Range (1C; pg) | Mean |
---|---|---|
Gymnosperms | ||
Conifers | ||
Pinaceae | 9.5-36.0 | 23.7 |
Cupressaceae | 8.3-32.1 | 12.8 |
Sciadopitys | 20.8 | n/a |
Gnetales | ||
Ephedraceae | 8.9-15.7 | 8.9 |
Gnetaceae | 2.3-4.0 | 2.3 |
Cycadaceae | 12.6-14.8 | 13.4 |
Ginkgo biloba | 11.75 | n/a |
Monilophytes | ||
Ophioglossaceae | 10.2-65.6 | 31.05 |
Equisetaceae | 12.9-304 | 22.0 |
Psilotum | 72.7 | n/a |
Leptosporangiate ferns | ||
Polypodiaceae | 7.5-19.7 | 7.5 |
Aspleniaceae | 4.1-9.1 | 6.2 |
Athyriaceae | 6.3-9.3 | 7.6 |
Dryopteridaceae | 6.8-23.6 | 11.7 |
Water ferns | ||
Azolla | 0.77 | n/a |
Angiosperms | ||
Oryza sativa | 0.50 | n/a |
Amborella trichopoda | 0.89 | n/a |
Arabidopsis thaliana | 0.16 | n/a |
Zea mays | 2.73 | n/a |
2.
José Carlos Quintela Francisco García-del Portillo Ernst Pittenauer Günter Allmaier Miguel A. de Pedro 《Journal of bacteriology》1999,181(1):334-337
Peptidoglycan from Deinococcus radiodurans was analyzed by high-performance liquid chromatography and mass spectrometry. The monomeric subunit was: N-acetylglucosamine–N-acetylmuramic acid–l-Ala–d-Glu-(γ)–l-Orn-[(δ)Gly-Gly]–d-Ala–d-Ala. Cross-linkage was mediated by (Gly)2 bridges, and glycan strands were terminated in (1→6)anhydro-muramic acid residues. Structural relations with the phylogenetically close Thermus thermophilus are discussed.The gram-positive bacterium Deinococcus radiodurans is remarkable because of its extreme resistance to ionizing radiation (14). Phylogenetically the closest relatives of Deinococcus are the extreme thermophiles of the genus Thermus (4, 11). In 16S rRNA phylogenetic trees, the genera Thermus and Deinococcus group together as one of the older branches in bacterial evolution (11). Both microorganisms have complex cell envelopes with outer membranes, S-layers, and ornithine-Gly-containing mureins (7, 12, 19, 20, 22, 23). However, Deinococcus and Thermus differ in their response to the Gram reaction, having positive and negative reactions, respectively (4, 14). The murein structure for Thermus thermophilus HB8 has been recently elucidated (19). Here we report the murein structure of Deinococcus radiodurans with similar detail.D. radiodurans Sark (23) was used in the present study. Cultures were grown in Luria-Bertani medium (13) at 30°C with aeration. Murein was purified and subjected to amino acid and high-performance liquid chromatography (HPLC) analyses as previously described (6, 9, 10, 19). For further analysis muropeptides were purified, lyophilized, and desalted as reported elsewhere (6, 19). Purified muropeptides were subjected to plasma desorption linear time-of-flight mass spectrometry (PDMS) as described previously (1, 5, 16, 19). Positive and negative ion mass spectra were obtained on a short linear 252californium time-of-flight instrument (BioIon AB, Uppsala, Sweden). The acceleration voltage was between 17 and 19 kV, and spectra were accumulated for 1 to 10 million fission events. Calibration of the mass spectra was done in the positive ion mode with H+ and Na+ ions and in the negative ion mode with H− and CN− ions. Calculated m/z values are based on average masses.Amino acid analysis of muramidase (Cellosyl; Hoechst, Frankfurt am Main, Germany)-digested sacculi (50 μg) revealed Glu, Orn, Ala, and Gly as the only amino acids in the muramidase-solubilized material. Less than 3% of the total Orn remained in the muramidase-insoluble fraction, indicating an essentially complete solubilization of murein.Muramidase-digested murein samples (200 μg) were analyzed by HPLC as described in reference 19. The muropeptide pattern (Fig. (Fig.1)1) was relatively simple, with five dominating components (DR5 and DR10 to DR13 [Fig. 1]). The muropeptides resolved by HPLC were collected, desalted, and subjected to PDMS. The results are presented in Table Table11 compared with the m/z values calculated for best-matching muropeptides made up of N-acetylglucosamine (GlucNAc), N-acetylmuramic acid (MurNAc), and the amino acids detected in the murein. The more likely structures are shown in Fig. Fig.1.1. According to the m/z values, muropeptides DR1 to DR7 and DR9 were monomers; DR8, DR10, and DR11 were dimers; and DR12 and DR13 were trimers. The best-fitting structures for DR3 to DR8, DR11, and DR13 coincided with muropeptides previously characterized in T. thermophilus HB8 (19) and had identical retention times in comparative HPLC runs. The minor muropeptide DR7 (Fig. (Fig.1)1) was the only one detected with a d-Ala–d-Ala dipeptide and most likely represents the basic monomeric subunit. The composition of the major cross-linked species DR11 and DR13 confirmed that cross-linking is mediated by (Gly)2 bridges, as proposed previously (20). Open in a separate windowFIG. 1HPLC muropeptide elution patterns of murein purified from D. radiodurans. Muramidase-digested murein samples were subjected to HPLC analysis, and the A204 of the eluate was recorded. The most likely structures for each muroeptide as deduced by PDMS are shown. The position of residues in brackets is the most likely one as deduced from the structures of other muropeptides but could not be formally demonstrated. R = GlucNac–MurNac–l-Ala–d-Glu-(γ)→.
Open in a separate windowaDR5 and DR10 to DR13 were analyzed in both the positive and negative ion modes. Muropeptides DR1 to DR4 and DR6 to DR9 were analyzed in the positive mode only due to the small amounts of sample available. bMass difference between measured and calculated quasimolecular ion values. c[(Measured mass−calculated mass)/calculated mass] × 100. dN-Acetylglucosamine. eN-Acetylmuramitol. f(1→6)Anhydro-N-acetylmuramic acid. Structural assignments of muropeptides DR1, DR2, DR8 to DR10, and DR12 deserve special comments. The low m/z value measured for DR1 (700.1) fitted very well with the value calculated for GlucNAc–MurNAc–l-Ala–d-Glu (699.69). Even smaller was the mass deduced for DR9 from the m/z value of the molecular ion of the sodium adduct (702.1) (Fig. (Fig.2).2). The mass difference between DR1 and DR9 (19.9 mass units) was very close indeed to the calculated difference between N-acetylmuramitol and the (1→6)anhydro form of MurNAc (20.04 mass units). Therefore, DR9 was identified as GlucNAc–(1→6)anhydro-MurNAc–l-Ala–d-Glu (Fig. (Fig.1).1). Muropeptides with (1→6)anhydro muramic acid have been identified in mureins from diverse origins (10, 15, 17, 19), indicating that it might be a common feature among peptidoglycan-containing microorganisms. Open in a separate windowFIG. 2Positive-ion linear PDMS of muropeptide DR9. Muropeptide DR9 was purified, desalted by HPLC, and subjected to PDMS to determine the molecular mass. The masses for the dominant molecular ions are indicated.The measured m/z value for the [M+Na]+ ion of DR8 was 1,521.6, very close to the mass calculated for a cross-linked dimer without one disaccharide moiety (1,520.53) (Fig. (Fig.1;1; Table Table1).1). Such muropeptides, also identified in T. thermophilus HB8 and other bacteria (18, 19), are most likely generated by the enzymatic clevage of MurNAc–l-Ala amide bonds in murein by an N-acetylmuramyl–l-alanine amidase (21). In particular, DR8 could derive from DR11. The difference between measured m/z values for DR8 and DR11 was 478.7, which fits with the mass contribution of a disaccharide moiety (480.5) within the mass accuracy of the instrument.The m/z values for muropeptides DR2, DR10, and DR12 supported the argument for structures in which the two d-Ala residues from the d-Ala–d-Ala C-terminal dipeptide were lost, leaving Orn as the C-terminal amino acid.The position of one Gly residue in muropeptides DR2, DR8, and DR10 to DR13 could not be formally demonstrated. One of the Gly residues could be at either the N- or the C-terminal positions. However, the N-terminal position seems more likely. The structure of the basic muropeptide (DR7), with a (Gly)2 acylating the δ-NH2 group of Orn, suggests that major muropeptides should present a (Gly)2 dipeptide. The scarcity of DR3 and DR6, which unambiguously have Gly as the C-terminal amino acid (Fig. (Fig.1),1), supports our assumption.Molar proportions for each muropeptide were calculated as proposed by Glauner et al. (10) and are shown in Table Table1.1. For calculations the structures of DR10 to DR13 were assumed to be those shown in Fig. Fig.1.1. The degree of cross-linkage calculated was 47.2%. Trimeric muropeptides were rather abundant (8 mol%) and made a substantial contribution to total cross-linkage. However, higher-order oligomers were not detected, in contrast with other gram-positive bacteria, such as Staphylococcus aureus, which is rich in such oligomers (8). The proportion of muropeptides with (1→6)anhydro-muramic acid (5 mol%) corresponded to a mean glycan strand length of 20 disaccharide units, which is in the range of values published for other bacteria (10, 17).The results of our study indicate that mureins from D. radiodurans and T. thermophilus HB8 (19) are certainly related in their basic structures but have distinct muropeptide compositions. In accordance with the phylogenetic proximity of Thermus and Deinococcus (11), both mureins are built up from the same basic monomeric subunit (DR7 in Fig. Fig.1),1), are cross-linked by (Gly)2 bridges, and have (1→6)anhydro-muramic acid at the termini of glycan strands. Most interestingly, Deinococcus and Thermus are the only microorganisms identified at present with the murein chemotype A3β as defined by Schleifer and Kandler (20). Nevertheless, the differences in muropeptide composition were substantial. Murein from D. radiodurans was poor in d-Ala–d-Ala- and d-Ala–Gly-terminated muropeptides (2.2 and 2.4 mol%, respectively) but abundant in Orn-terminated muropeptides (23.8 mol%) and in muropeptides with a peptide chain reduced to the dipeptide l-Ala–d-Glu (18 mol%). In contrast, neither Orn- nor Glu-terminated muropeptides have been detected in T. thermophilus HB8 murein, which is highly enriched in muropeptides with d-Ala–d-Ala and d-Ala–Gly (19). Furthermore, no traces of phenyl acetate-containing muropeptides, a landmark for T. thermophilus HB8 murein (19), were found in D. radiodurans. Cross-linkage was definitely higher in D. radiodurans than in T. thermophilus HB8 (47.4 and 27%, respectively), largely due to the higher proportion of trimers in the former.The similarity in murein basic structure suggests that the difference between D. radiodurans and T. thermophilus HB8 with respect to the Gram reaction may simply be a consequence of the difference in the thickness of cell walls (2, 3, 23). Interestingly, D. radiodurans murein turned out to be relatively simple for a gram-positive organism, possibly reflecting the primitive nature of this genus as deduced from phylogenetic trees (11). Our results illustrate the phylogenetic proximity between Deinococcus and Thermus at the cell wall level but also point out the structural divergences originated by the evolutionary history of each genus. 相似文献
TABLE 1
Calculated and measured m/z values for the molecular ions of the major muropeptides from D. radioduransMuropeptidea | Ion | m/z
| Δmb | Error (%)c | Muropeptide composition
| Muropeptide abundance (mol%) | ||||||
---|---|---|---|---|---|---|---|---|---|---|---|---|
Calculated | Measured | NAGd | NAMe | Glu | Orn | Ala | Gly | |||||
DR1 | [M+H]+ | 699.69 | 700.1 | 0.41 | 0.06 | 1 | 1 | 1 | 0 | 1 | 0 | 12.0 |
DR2 | [M+H]+ | 927.94 | 928.3 | 0.36 | 0.04 | 1 | 1 | 1 | 1 | 1 | 2 | 5.7 |
DR3 | [M+Na]+ | 1,006.97 | 1,007.5 | 0.53 | 0.05 | 1 | 1 | 1 | 1 | 1 | 3 | 3.0 |
DR4 | [M+Na]+ | 963.95 | 964.6 | 0.65 | 0.07 | 1 | 1 | 1 | 1 | 2 | 1 | 2.5 |
DR5 | [M+H]+ | 999.02 | 999.8 | 0.78 | 0.08 | 1 | 1 | 1 | 1 | 2 | 2 | 27.7 |
[M−H]− | 997.00 | 997.3 | 0.30 | 0.03 | ||||||||
DR6 | [M+Na]+ | 1,078.5 | 1,078.8 | 0.75 | 0.07 | 1 | 1 | 1 | 1 | 2 | 3 | 2.4 |
DR7 | [M+H]+ | 1,070.09 | 1,071.0 | 0.90 | 0.08 | 1 | 1 | 1 | 1 | 3 | 2 | 2.2 |
DR8 | [M+Na]+ | 1,520.53 | 1,521.6 | 1.08 | 0.07 | 1 | 1 | 2 | 2 | 4 | 4 | 2.2 |
DR9 | [M+Na]+ | 701.64 | 702.1 | 0.46 | 0.03 | 1 | 1f | 1 | 0 | 1 | 0 | 5.0 |
DR10 | [M+H]+ | 1,907.94 | 1,907.8 | 0.14 | 0.01 | 2 | 2 | 2 | 2 | 3 | 4 | 10.1 |
[M−H]− | 1,905.92 | 1,906.6 | 0.68 | 0.04 | ||||||||
DR11 | [M+H]+ | 1,979.01 | 1,979.1 | 0.09 | 0.01 | 2 | 2 | 2 | 2 | 4 | 4 | 19.1 |
[M−H]− | 1,977.00 | 1,977.3 | 0.30 | 0.02 | ||||||||
DR12 | [M+H]+ | 2,887.93 | 2,886.5 | −1.43 | −0.05 | 3 | 3 | 3 | 3 | 5 | 6 | 4.4 |
[M−H]− | 2,885.91 | 2,885.8 | −0.11 | −0.01 | ||||||||
DR13 | [M+H]+ | 2,959.00 | 2,957.8 | −1.20 | −0.04 | 3 | 3 | 3 | 3 | 6 | 6 | 3.6 |
[M−H]− | 2,956.99 | 2,955.9 | −1.09 | −0.04 |
3.
John Chen Nuria Carpena Nuria Quiles-Puchalt Geeta Ram Richard P Novick José R Penadés 《The ISME journal》2015,9(5):1260-1263
Bacteriophage-mediated horizontal gene transfer is one of the primary driving forces of bacterial evolution. The pac-type phages are generally thought to facilitate most of the phage-mediated gene transfer between closely related bacteria, including that of mobile genetic elements-encoded virulence genes. In this study, we report that staphylococcal cos-type phages transferred the Staphylococcus aureus pathogenicity island SaPIbov5 to non-aureus staphylococcal species and also to different genera. Our results describe the first intra- and intergeneric transfer of a pathogenicity island by a cos phage, and highlight a gene transfer mechanism that may have important implications for pathogen evolution.Classically, transducing phages use the pac site-headful system for DNA packaging. Packaging is initiated on concatemeric post-replicative DNA by terminase cleavage at the sequence-specific pac site, a genome slightly longer than unit length is packaged, and packaging is completed by non-sequence-specific cleavage (reviewed in Rao and Feiss, 2008). Generalized transduction results from the initiation of packaging at pac site homologs in host chromosomal or plasmid DNA, and typically represents ∼1% of the total number of phage particles. In the alternative cos site mechanism packaging is also initiated on concatemeric post-replicative DNA by terminase cleavage at a sequence-specific (cos) site. Here, however, packaging is completed by terminase cleavage at the next cos site, generating a precise monomer with the cohesive termini used for subsequent circularization (Rao and Feiss, 2008). Although cos site homologs may exist in host DNA, it is exceedingly rare that two such sites would be appropriately spaced. Consequently, cos phages, of which lambda is the prototype, do not engage in generalized transduction. For this reason, cos-site phages have been preferred for possible phage therapy, since they would not introduce adventitious host DNA into target organisms.The Staphylococcus aureus pathogenicity islands (SaPIs) are the best-characterized members of the phage-inducible chromosomal island family of mobile genetic elements (MGEs; Novick et al., 2010). SaPIs are ∼15 kb mobile elements that encode virulence factors and are parasitic on specific temperate (helper) phages. Helper phage proteins are required to lift their repression (Tormo-Más et al., 2010, 2013), thereby initiating their excision, circularization and replication. Phage-induced lysis releases vast numbers of infectious SaPI particles, resulting in high frequencies of transfer. Most SaPI helper phages identified to date are pac phages, and many well-studied SaPIs are packaged by the headful mechanism (Ruzin et al., 2001; Ubeda et al., 2007). Recently, we have reported that some SaPIs, of which the prototype is SaPIbov5 (Viana et al., 2010), carry phage cos sequences in their genomes, and can be efficiently packaged and transferred by cos phages to S. aureus strains at high frequencies (Quiles-Puchalt et al., 2014). Here we show that this transfer extends to non-aureus staphylococci and to Listeria monocytogenes.Since the pac phages transfer SaPIs to non-aureus staphylococci and to the Gram-positive pathogen Listeria monocytogenes (Maiques et al., 2007; Chen and Novick, 2009), we reasoned that cos phages might also be capable of intra- and intergeneric transfer. We tested this with SaPIbov5, into which we had previously inserted a tetracycline resistance (tetM) marker to enable selection, and with lysogens of two helper cos phages, φ12 and φSLT, carrying SaPIbov5 (strains JP11010 and JP11194, respectively; Supplementary Table 1). The prophages in these strains were induced with mitomycin C, and the resulting lysates were adjusted to 1 μg ml−1 DNase I and RNase A, filter sterilized (0.2 μm pore), and tested for SaPI transfer with tetracycline selection, as previously described (Ubeda et al., 2008). To test for trans-specific or trans-generic transduction, coagulase-negative staphylococci species and L. monocytogenes strains were used as recipients for SaPIbov5 transfer, respectively, as previously described (Maiques et al., 2007; Chen and Novick, 2009). As shown in Figure 1 and Supplementary Table 2). In contrast, deletion of the SaPIbov5 cos site (strains JP11229 and JP11230) did not affect SaPI replication (Supplementary Figure 1), but completely eliminated SaPIbov5 transfer (Supplementary Table 2). The TerS protein is essential for φ12 and SaPIbov5 DNA packaging, but not for phage-mediated lysis (Quiles-Puchalt et al., 2014). As expected, this mutation abolished SaPIbov5 transfer (Open in a separate windowFigure 1(a) Map of SaPIbov5. Arrows represent the localization and orientation of ORFs greater than 50 amino acids in length. Rectangles represent the position of the ori (in purple) or cos (in red) sites. Positions of different primers described in the text are shown. (b) Amplimers generated for detection of SaPIbov5 in the different recipient strains. Supplementary Table 2 lists the sequence of the different primers used. The element was detected in S. epidermidis JP829 (Se-1), S. epidermidis JP830 (Se-2), L. monocytogenes SK1351 (Lm-1), L. monocytogenes EGDe (Lm-2), S. xylosus C2a (Sx) and S. aureus JP4226 (Sa).
Open in a separate windowAbbreviation: SAPI, Staphylococcus aureus pathogenicity island.aThe means of results from three independent experiments are shown. Variation was within ±5% in all cases.bNo. of transductants per ml induced culture.Because plaque formation is commonly used to determine phage host range, we next determined the ability of phages φ12 and φSLT to parasitize and form plaques on S. xylosus, S. epidermidis and L. monocytogenes strains. As shown in Supplementary Figure 2, phages φ12 and φSLT can parasitize and form plaques on their normal S. aureus hosts, but are completely unable to lyse the non-aureus strains. Therefore, as previously observed with pac phages (Chen and Novick, 2009), these results indicate that the overall host range of a cos phage may also be much wider if it includes infection without plaque formation.Previous studies have demonstrated pac phage-mediated transfer of MGEs between S. aureus and other bacterial species (Maiques et al., 2007; Chen and Novick, 2009; Uchiyama et al., 2014); however, no previous studies have described the natural intra- or intergeneric transfer of pathogenicity islands by cos phages. As bacterial pathogens become increasingly antibiotic resistant, lytic and poorly transducing phages, such as cos phages, have been proposed for phage therapy, on the grounds that they would not introduce adventitious host DNA into target organisms and that the phages are so restricted in host range that the resulting progeny are harmless and will not result in dysbiosis of human bacterial flora. Because plaque formation was once thought to determine the host range of a phage, the evolutionary impact of phages on bacterial strains they can transduce, but are unable to parasitize, has remained an unrecognized aspect of phage biology and pathogen evolution. Our results add to the recently recognized concept of ‘silent transfer'' of pathogenicity factors carried by MGEs (Maiques et al., 2007; Chen and Novick, 2009) by phages that cannot grow on the target organism. They extend this capability to cos phages, which have hitherto been unrecognized as mediators of natural genetic transfer.The potential for gene transfer of MGEs by this mechanism is limited by the ability of cos phages to adsorb and inject DNA into recipient strains, and also by the presence of suitable attachment sites in recipient genomes. However, since different bacterial genera express wall teichoic acid with similar structures, which can act as bacteriophage receptors governing the routes of horizontal gene transfer between major bacterial pathogens, horizontal gene transfer even across long phylogenetic distances is possible (Winstel et al., 2013). In addition, our previous results also demonstrated that the SaPI integrases have much lower sequence specificity than other typical integrases, and SaPIs readily integrate into alternative sites in the absence of the cognate attC site, such that any bacterium that can adsorb SaPI helper phage is a potential recipient (Chen and Novick, 2009). Thus, we anticipate that cos phages can have an important role in spreading MGEs carrying virulence and resistance genes. We also predict that cos sites will be found on many other MGEs, enabling cos phage-mediated transfer of any such element that can generate post-replicative concatemeric DNA. 相似文献
Table 1
Intra- and intergeneric SaPIbov5 transferaDonor strain | |||
---|---|---|---|
Phage | SaPI | Recipient strain | SaPI titreb |
φ12 | SaPIbov5 | S. aureus JP4226 | 8.3 × 104 |
S. epidermidis JP829 | 2.4 × 104 | ||
S. epidermidis JP830 | 4.7 × 104 | ||
L. monocytogenes SK1351 | 6.6 × 103 | ||
L. monocytogenes EGDe | 2.1 × 104 | ||
S. xylosus C2a | 7.1 × 104 | ||
φ12 | SaPIbov5 Δcos | S. aureus JP4226 | <10 |
S. epidermidis JP829 | <10 | ||
S. epidermidis JP830 | <10 | ||
L. monocytogenes SK1351 | <10 | ||
L. monocytogenes EGDe | <10 | ||
S. xylosus C2a | <10 | ||
φ12 ΔterS | SaPIbov5 | S. aureus JP4226 | <10 |
S. epidermidis JP829 | <10 | ||
S. epidermidis JP830 | <10 | ||
L. monocytogenes SK1351 | <10 | ||
L. monocytogenes EGDe | <10 | ||
S. xylosus C2a | <10 | ||
φSLT | SaPIbov5 | S. aureus JP4226 | 4.1 × 103 |
S. epidermidis JP829 | 1.1 × 103 | ||
S. epidermidis JP830 | 2.1 × 103 | ||
L. monocytogenes SK1351 | 3.6 × 102 | ||
L. monocytogenes EGDe | 3.1 × 103 | ||
S. xylosus C2a | 4.0 × 103 | ||
φSLT | SaPIbov5 Δcos | S. aureus JP4226 | <10 |
S. epidermidis JP829 | <10 | ||
S. epidermidis JP830 | <10 | ||
L. monocytogenes SK1351 | <10 | ||
L. monocytogenes EGDe | <10 | ||
S. xylosus C2a | <10 |
4.
5.
6.
Takehiko Kenzaka Masao Nasu Katsuji Tani 《Applied and environmental microbiology》2010,76(4):1274-1277
The transfer range of phage genes was investigated at the single-cell level by using an in situ DNA amplification technique. After absorption of phages, a phage T4 gene was maintained in the genomes of non-plaque-forming bacteria at frequencies of 10−2 gene copies per cell. The gene transfer decreased the mutation frequencies in nonhost recipients.Recently, whole-genome analyses have revealed that many bacterial genomes contain foreign genes, especially phage genes (9). The phage genes in bacterial genomes include genes for virulence or fitness factors such as extracellular toxins, superantigens, lipopolysaccharide-modifying enzymes, and proteins conferring serum resistance, etc. (1). These findings suggest that the horizontal transfer of phage genes has contributed significantly to the acquisition of new genetic traits and to the genetic diversity of bacteria (1, 9, 10). To truly appreciate the mechanisms behind phage-associated evolution, it is important to understand the frequency and range of transfer of phage genes.Most phage genomes consist of many genes derived from different origins (5, 8). Some genes are similar to those of other phages with phylogenetically different hosts or are found in the genomes of bacteria that are not the phage hosts. The mosaic nature of phage genomes has been known for some time, and a body of molecular genetic studies of phages to explain the mechanisms that drive this feature have been attempted previously (1, 5). More importantly, the horizontal transfer of phage genes has emerged as a major factor in the evolution of the phage genome. Since recombination between phage and phage/prophage can occur when these elements coexist in the same cell, coinfection with multiple phage species may result in the production of hybrid phage genomes (5). The pathways by which phages exchange genetic material vary dramatically in concert with host ranges. However, conventional plaque assays have shown that the host ranges of the phages studied are narrow. We hypothesized that phage genes can be transferred to more diverse species than previously thought.In order to accurately quantify DNA movement, gene targeting that does not require cultivation or gene expression is necessary (7). In situ DNA amplification methods allow the visualization of specific DNA sequences inside bacterial cells. In this study, we employed cycling primed in situ amplification-fluorescent in situ hybridization (CPRINS-FISH) to examine the possible range and frequency of the transfer of phage genes. CPRINS uses one primer and results in linear amplification of the target DNA inside cells, and multiply labeled fluorescent probe sets are applied for detection of the amplicons to improve the specificity and sensitivity of CPRINS (3). Previously, CPRINS-FISH did clarify the movement of DNA of a specific gene among Escherichia coli cells at the single-cell level (4).Enterobacterial phages P1 and T4 infect E. coli and have been well studied. P1 can exist as circular DNA within the bacterial cell as if it were a plasmid. Phage T4 is capable of undergoing only a lytic life cycle and not the lysogenic life cycle. Conventional methods using plaque assays have shown that the host of P1 and T4 is E. coli, but orthologous phage genes have been found in bacteria other than E. coli (6, 8). In the present study, strains of Enterobacteriaceae were allowed to grow on agar medium after the phage was adsorbed, and the maintenance of the transferred phage gene in the bacterial genomes was examined at the community level by quantitative real-time PCR and at the single-cell level by CPRINS-FISH.The following bacterial strains were used for maintenance experiments: Citrobacter freundii IFO 12681, Enterobacter aerogenes BM 2688, E. coli NBRC 12713, a Proteus mirabilis clinical isolate, Salmonella enterica serovar Enteritidis IID 640, and Yersinia enterocolitica IID 981. The bacterial strains were grown in Luria-Bertani (LB) medium (1% tryptone, 0.5% yeast extract, 0.5% NaCl; Nacalai Tesque Inc., Kyoto, Japan) at 37°C overnight.Stationary-phase cultures of 500 μl were incubated with 500 μl of SM buffer (50 mmol liter−1 Tris-HCl [pH 7.5], 100 mmol liter−1 NaCl, 8 mmol liter−1 MgSO4, 0.01% gelatin) containing the phage P1kc NBRC 20008 (2) or T4GT7 (11) at 37°C for 10 min at a multiplicity of infection of 1:1 (ratio of PFU of the phage to CFU of the recipient bacterium). The concentration of bacterial cells was adjusted to 109 cells ml−1. After 10 min of incubation, the diluted cell suspension (105 cells) was filtered through a polycarbonate filter (Advantec, Tokyo, Japan) with a pore size of 0.2 μm and a diameter of 25 mm. Cells trapped on the filter were cultured on LB agar medium at 37°C for 24 h. The filter was transferred into a microtube, and cells on the filter were suspended in 1 ml of sterile deionized water. The numbers of cells in the suspension and cells remaining on the filter were determined by using an epifluorescence microscope (see below) after staining of the samples with 1 μg ml−1 of 4′,6-diamidino-2-phenylindole (DAPI; Sigma-Aldrich Japan, Tokyo). The level of recovery of cells from the filter into sterile deionized water was about 99%. The cultured cells were subjected to real-time PCR and CPRINS-FISH.For real-time PCR, bacterial DNA was extracted using a QIAamp DNA isolation kit (Qiagen, Tokyo, Japan). The cell suspension was mixed with 10 mg ml−1 of lysozyme solution and incubated at 37°C for 1 h. DNA extraction was then performed according to the kit manufacturer''s instructions. Table Table11 lists the oligonucleotide primers for PCR and CPRINS and the polynucleotide probes used in the present study. Tail fiber genes from phages P1kc and T4GT7 were quantified by real-time PCR with a LightCycler system (Roche Diagnostics, Tokyo, Japan). LightCycler FastStart DNA master SYBR green I (Roche Diagnostics) was used with 5 mmol liter−1 Mg2+ and 0.5 μmol liter−1 (each) primers targeting the tail fiber genes of P1kc (P1-tail931f and P1-tail1148r) and T4GT7 (T4-tail2770f and T4-tail2983r). After a hot start for 10 min at 95°C, 40 cycles of PCR were run with denaturation at 94°C for 15 s, annealing at 60°C for 10 s, extension at 72°C for 10 s, and fluorescence detection at 83°C for 5 s. The known amounts of PCR products from the phage DNA (101 to 107 copies per reaction) were used for the standard curves to quantify the target DNA. To confirm the specificity of the reaction after real-time PCR, the PCR mixture was collected in a glass capillary and subjected to agarose gel electrophoresis in addition to a melting-curve analysis with the LightCycler system. The maintenance frequencies determined by real-time PCR were recorded as the copy number of the phage tail fiber gene per bacterial genome detected by staining with PicoGreen (Invitrogen, Tokyo, Japan) after cultivation of cells on LB agar medium for 24 h as described above. The frequencies were determined in triplicate for each sample. The increase in the phage gene copy number was determined by comparing the copy numbers in cells on the filter before and after cultivation. The phage gene copy number in cells on the filter was determined by the following formula: (total number of cells determined by DAPI staining) × (phage tail fiber gene copy number determined by real-time PCR)/(bacterial genome copy number determined by PicoGreen staining).
Open in a separate windowCPRINS-FISH targeting the tail fiber gene of phage T4GT7 was performed as described by Kenzaka et al. (3, 4), except for the probe/primer sequences and thermal conditions. After cell wall permeabilization by lysozyme treatment (3), the CPRINS reaction was performed under the following conditions: a hot start at 95°C for 9 min, denaturation at 94°C for 1 min, annealing at 60°C for 30 s, and extension at 72°C for 1.5 min for primer T4-tail2983r. Amplification was repeated for 30 cycles by using a thermal cycler (PTC-200; Bio-Rad Laboratories, Inc.). After amplification, filters were rinsed with 0.1% Nonidet P-40 and sterile deionized water, dehydrated in 99% ethanol, and vacuum dried. Hybridization with Alexa Fluor 546-labeled polynucleotide probes (T4-tail2664, T4-tail2720, T4-tail2769, T4-tail2818, T4-tail2869, and T4-tail2922), washing, and DAPI staining were performed as described in a previous study (4). In order to exclude the possibility of nonspecific probe binding to cell structures other than the target DNA in the target cells, FISH using laboratory strains without amplification of target DNA and CPRINS-FISH targeting the tail fiber genes in E. coli strains that did not carry the genes were performed.In order to examine the infection ranges of phages, plaque assays and direct counting of phages were performed. Plaque assays were performed with LB soft agar (0.8% agar) as described by Kenzaka et al. (4). For the direct counting, phages were stained with 5× SYBR gold (Invitrogen, Tokyo, Japan) and trapped onto an Anodisc filter (Whatman Japan, Tokyo) with a pore size of 0.02 μm and a diameter of 25 mm.The cells or phage particles on the filters were observed under an epifluorescence microscope (E-400; Nikon, Tokyo, Japan) with the Nikon filter sets UV-2A (EX300-350, DM400, and BA420) for DAPI, B-2A (EX450-490, DM505, and BA520) for SYBR gold, and HQ-CY3 (G535/50, FT565, and BP610/75) for Alexa Fluor 546. Images were acquired using a Retiga 2000R cooled charge-coupled device camera (QImaging, Surrey, BC, Canada), and at least 2,000 DAPI- or SYBR gold-stained objects per sample were counted. The maintenance frequencies determined by CPRINS-FISH were recorded as the number of CPRINS-FISH-positive cells divided by the total direct count of recipient cells after cultivation as described above. The frequencies were determined in triplicate for each sample.After cultivation on LB agar medium for 24 h, the total number of cells on the filter as determined by DAPI staining increased by 8.7 × 102- to 1.1 × 104-fold (Table (Table2).2). Real-time PCR showed that the phage P1kc gene copy number increased only in plaque-forming strains (E. coli and E. aerogenes) and not in non-plaque-forming strains (Table (Table2).2). In contrast, the phage T4GT7 gene copy number increased in both plaque-forming and non-plaque-forming strains by 7.6 × 101- to 7.0 × 104-fold. The maintenance frequencies were more than 10−2 gene copies per bacterial genome (Table (Table2).2). Direct observation via epifluorescence microscopy showed that progeny phages were not produced in the non-plaque-forming strains (Table (Table2),2), and thus, fragments of phage genes were thought to integrate into the genomes of non-plaque-forming strains and replicate with the bacterial genomes.
Open in a separate windowaPlaque formation on soft agar was tested.bThe production of progeny phage particles was observed via epifluorescence microscopy.cThe increase (n-fold) in the total number of cells during bacterial growth for 24 h was determined via epifluorescence microscopy.dThe increase (n-fold) in the copy number of the phage tail fiber gene during bacterial growth for 24 h was determined by real-time PCR. Values in parentheses indicate standard deviations of results for triplicate samples.eMaintenance frequencies were determined by real-time PCR and CPRINS-FISH analyses targeting the phage tail fiber gene and are shown as the phage tail fiber gene copy numbers per bacterial genome and the numbers of gene-positive cells divided by the total numbers of cells, respectively. Values in parentheses indicate standard deviations of results for triplicate samples. ND, not determined.Real-time PCR provided a copy number for the target phage gene in the whole population, but the location of the target phage gene and the frequency of cells carrying the target gene were unclear. In addition, bacterial genomic DNA, which was measured using PicoGreen, included phage DNA, and thus the frequencies measured by dividing by the amount of bacterial genomic DNA were probably less accurate than those measured as described below. In order to confirm that the phage gene was located inside bacterial cells and determine a more accurate maintenance frequency for total cells, CPRINS-FISH targeting the tail fiber gene of phage T4GT7 was performed. CPRINS-FISH visualized the target phage gene in individual cells under an epifluorescence microscope (Fig. (Fig.1).1). It showed that the frequencies of maintenance of the tail fiber gene, expressed as the number of gene-positive cells divided by the total number of cells, were 2.1 × 10−1 to 4.0 × 10−1 for plaque-forming strains after growth on LB medium for 24 h (Table (Table2).2). Since phage T4GT7 is capable of undergoing only a lytic life cycle, CPRINS-FISH would detect cells in which the phage gene was replicating. For non-plaque-forming strains, the maintenance frequencies were 2.2 × 10−2 to 8.8 × 10−2 (Table (Table2).2). If the gene was amplified by the CPRINS reaction outside bacterial cells, the amplicon would not accumulate inside bacterial cells and they would not exhibit bright fluorescence. Therefore, CPRINS-FISH proved that a part of the phage T4GT7 gene was located inside cells of non-plaque-forming strains. The tail fiber gene is responsible for the phage tail structure. The DNA sequences of the phage genes responsible for phage morphology have been found in many bacterial genomes (1, 5).Open in a separate windowFIG. 1.Visualization of E. coli cells carrying the tail fiber gene transferred by phage T4GT7. (A) After being mixed with phages for 10 min, E. coli NBRC 12713 cells were cultured for 24 h and subjected to CPRINS-FISH targeting the phage gene. Only cells having amplified tail fiber gene products emitted the fluorescence of the Alexa Fluor 546-labeled probe under green excitation (exposure, 0.5 s). (B) All DAPI-stained bacterial cells were visualized under UV excitation (exposure time, 0.1 s).In order to explore the effect of integration of the phage gene into the bacterial genome on bacterial heredity, we determined the mutation frequency for a C. freundii strain that acquired the phage T4GT7 gene. Two colonies which acquired the phage T4GT7 gene were screened by colony PCR with T4-tail2770f and T4-tail2983r primers and designated Cik8-1 and Cik8-4. Mutation frequencies were determined with LB medium containing 150 μg ml−1of rifampin (rifampicin) or 10 μg ml−1of nalidixic acid. The mutation frequencies associated with nalidixic acid resistance decreased by 12- to 240-fold and the frequencies associated with rifampin resistance decreased by 40- to 83-fold compared to those for the parent strains (Fig. (Fig.2).2). Mutation increases genetic variation. The decreased mutation frequency would contribute to the genetic stability of the genome in individual cells but not to genetic variation in the population. Our results show that phage T4GT7 was capable of affecting the genomic properties of C. freundii, which was thought previously not to be the host, although the mechanism by which mutation frequencies decreased remains unknown. Further experiments are required to clarify the molecular mechanism by which mutation frequencies altered after gene transfer.Open in a separate windowFIG. 2.Mutation frequencies for T4GT7-infected C. freundii strains. Mutation frequencies were determined with LB agar medium containing nalidixic acid or rifampin. Cik8-1 and Cik8-4 were strains which acquired a phage gene transferred from phage T4GT7. Cik1 and Cik2 were the parent strains.In summary, during growth on agar medium after the phage was allowed to be adsorbed by strains of Enterobacteriaceae, the phage P1kc gene was not maintained in non-plaque-forming strains but the phage T4GT7 gene was maintained in more diverse species than previously expected. The transfer of foreign DNA molecules (DNA entry) into a bacterium is an important first step in genetic diversification through horizontal gene transfer. A previous study reported that phage P1kc is capable of injecting DNA into non-plaque-forming E. coli cells (4), but the phage P1kc gene was not maintained during bacterial growth in the present study. The results showing the difference in maintenance between phage P1kc and T4GT7 genes suggest that the maintenance of transferred phage genes depends on phage gene sequences or other phage factors. When maintained, the phage gene could alter the mutation frequency for bacteria that acquired the gene, affecting the genomic variability at the population level. Conventionally, phage-bacterium interaction has been studied with certain models consisting of a phage and a bacterium in which the phage can multiply (12, 13). Our results indicate the importance of the dynamic of phage genes among diverse bacteria that were previously thought not to be hosts and the hereditary impact of phage gene transfer on such bacteria. 相似文献
TABLE 1.
Probes and primers designed in this studyName | Target | Type | Nucleotide sequence (5′-3′) |
---|---|---|---|
P1-tail931f | Tail fiber gene of phage P1 | Primer | AACGACCCGAATTACAGCAC |
P1-tail1148r | Tail fiber gene of phage P1 | Primer | AGTGCTGCTGCAAGCTCATA |
T4-tail2770f | Tail fiber gene of phage T4 | Primer | AGCACAAATGGTGAGCACAG |
T4-tail2983r | Tail fiber gene of phage T4 | Primer | TTGCTACCGTGTGGGTATGA |
T4-tail2664 | Tail fiber gene of phage T4 | Probe | GGCTTCAAGTACTGACTTAGGTACTAAAACCACATCAAGCTTTGACTATGGTACG |
T4-tail2720 | Tail fiber gene of phage T4 | Probe | AAGGGAACTAACAGTACGGGTGGACACACTCACTCTGGTAGTGGTTCTA |
T4-tail2769 | Tail fiber gene of phage T4 | Probe | TAGCACAAATGGTGAGCACAGCCACTACATCGAGGCATGGAATGG |
T4-tail2818 | Tail fiber gene of phage T4 | Probe | GGTGTAGGTGGTAATAAGATGTCATCATATGCCATATCATACAGGGCGGG |
T4-tail2869 | Tail fiber gene of phage T4 | Probe | GGGAGTAACACTAATGCAGCAGGGAACCACAGTCACACTTTCTCTTTTGGG |
T4-tail2922 | Tail fiber gene of phage T4 | Probe | TAGCAGTGCTGGCGACCATTCCCACTCTGTAGGTATTGGTGCTCATA |
TABLE 2.
Frequencies of maintenance of phage P1kc and T4GT7 genes in Enterobacteriaceae strainsPhage | Recipient | Result for infection range indicator: | Increase in total no. of cellsc | Increase in phage gene copy no. (SD)d | Maintenance frequency (SD) as determined bye: | ||
---|---|---|---|---|---|---|---|
Plaque formationa | Production of progenyb | Real-time PCR | CPRINS-FISH | ||||
P1kc | C. freundii | − | − | 7.0 × 103 | None | <1.5 × 10−3 | ND |
E. aerogenes | + | + | 1.7 × 103 | 7.7 × 103 (6.5 × 103) | 5.0 × 100 (4.2 × 100) | ND | |
E. coli | + | + | 7.2 × 103 | 5.5 × 103 (2.7 × 103) | 9.1 × 10−1 (0.5 × 10−1) | ND | |
P. mirabilis | − | − | 7.4 × 103 | None | <1.5 × 10−3 | ND | |
S. Enteritidis | − | − | 8.4 × 103 | None | <1.7 × 10−4 | ND | |
Y. enterocolitica | − | − | 4.6 × 103 | None | <1.8 × 10−4 | ND | |
T4GT7 | C. freundii | − | − | 1.5 × 103 | 7.5 × 103 (4.0 × 103) | 8.3 × 10−1 (4.4 × 10−1) | 8.6 × 10−2 (3.4 × 10−2) |
E. aerogenes | + | + | 8.7 × 102 | 1.2 × 103 (0.8 × 103) | 8.0 × 10−1 (5.0 × 10−1) | 4.0 × 10−1 (0.7 × 10−1) | |
E. coli | + | + | 1.1 × 104 | 7.0 × 104 (2.7 × 104) | 8.0 × 101 (3.0 × 10) | 2.1 × 10−1 (0.4 × 10−1) | |
P. mirabilis | − | − | 4.0 × 103 | 5.8 × 103 (4.2 × 103) | 3.3 × 10−1 (2.4 × 10−1) | 3.4 × 10−2 (2.2 × 10−2) | |
S. Enteritidis | − | − | 1.0 × 104 | 7.6 × 101 (5.0 × 101) | 1.0 × 10−2 (0.7 × 10−2) | 8.8 × 10−2 (2.0 × 10−2) | |
Y. enterocolitica | − | − | 3.6 × 103 | 1.6 × 104 (0.4 × 104) | 6.1 × 10−1 (1.6 × 10−1) | 2.2 × 10−2 (2.9 × 10−2) |
7.
As translational clinical researchers familiar with the risk-benefit of hematopoietic stem cell transplantation in autoimmune diseases, we are intrigued by the recent report of umbilical cord mesenchymal stem cell (UC-MSC) transplantation in treatment-refractory systemic lupus erythematosus nephritis by Wang and colleagues. They report the results of an open-label single-arm multicenter phase I/II study. This stimulated us to examine whether collective data from this group provide sufficient evidence for the feasibility, safety, dose rationale, and potential efficacy of UC-MSCs to conduct a randomized controlled trial in such patients. Results, though confounded by variable baseline prednisone and immuno-suppressive treatment, appear to indicate near-term response rates of approximately 50%, which are comparable to those seen with hematopoietic stem cell transplantation but with less morbidity and mortality.Wang and colleagues have been in the forefront of evaluating the potential for mesenchymal stem cells (MSCs) to treat systemic lupus erythematosus (SLE), based on studies in murine autoimmune disease models, demonstrating immunomodulatory properties of MSCs [1]. We have reviewed the additional reports published in peer-reviewed journals from this group [2–5], together with two protocols available on ClinicalTrials.gov ( and NCT00698191) (Table NCT01741857Authors (date) ClinicalTrials.gov protocol number Study design/duration of follow-up Number of patients MSC type/regimen Conditioning Safety: deaths/serious infection PD marker
a
Efficacy Sun et al. [2] NR Single-arm/ median of 8.25 months (range of 3 to 28 months) 16 (15 SLEN) UC, single infusion CYC 0.8 to 1.8 mg/kg intravenously, 2 to 4 days 0/0 Percentage of Treg cells increased at 3 months (P = 0.03) ‘Decreasing SLEDAI and proteinuriab in all patients’ Liang et al. [3] NCT 00698191 Single-arm/17.2 ± 9.5 months 15 SLEN BM, single infusion Included in protocol, but NR 0/0 Percentage of Treg cells increased at 1 week and 3 and 6 months (P <0.05) ‘Decreasing SLEDAI and proteinuriab in all patients’ Wang et al. [4] NCT 00698191 Unblinded-randomized, 2-arm/12 months 58 (~88% SLEN) BM, UC, single versus 2× (7 days apart) CYC 10 mg/kg per day, day 4, 3, and 2 1/NR ND CR single: 16/30 (53%); double: 8/27 (29%) Wang et al. [5] NR Single-arm/mean of 27 months 87 (84% SLEN) BM, UC, single infusion, 18 patients retreated at relapse CYC 10 mg/ kg/day, day 4, 3, and 2 5/NR ND CR in 23/83, relapse 10/83 Wang et al. [1]
c NCT 01741857 Single-arm 40 (38 SLEN) UC, 2× infusion, 7 days apart) No 3/4 ND MCR 13/PCR 11, 7 relapse