The nematode Heterorhabditis bacteriophora TH01, harboring the TT01 wild-type strain, was collected in Trinidad in 1993 43. The nematode was maintained in the laboratory and multiplied by infestation in the Lepidopteran Galleria mellonella 44. In 1998, a further bacterial isolate was taken from this nematode. During the course of a genetic study of the type III secretion system, we discovered that the bacterium isolated in 1998 is a genomic variant. It differs from the TT01 wild-type strain by a 250 base pair deletion at the 5' end of the gene lopT1 (Additional data file 1). This gene encodes a type III secretion system effector that appears to be involved in the depression of the insect innate immune system 45. Both TT01 wild-type and the lopT1 genomic variant produced many of the phenotypes associated with primary PVs, including bioluminescence, lipase activity, antibiotic production, and presence of cytoplasmic crystal (Table 1). Therefore, both were primary PVs. To distinguish between them, the TT01 wild-type strain was named TT01_/I and the lopT1 genomic variant, TT01α_/I (Figure 1).

We cultured TT01_/I and TT01α_/I in liquid broth and selected primary and secondary PVs on NBTA (nutrient agar supplemented with bromothymol blue and triphenyl 2,3,5 tetrazolium chloride) plates. TT01_/II secondary PV was derived from TT01_/I (TT01 lineage; Figure 1). TT01α_/II and TT01α'_/II secondary PVs were obtained from TT01α_/I (TT01α lineage; Figure 1). TT01_/II, TT01α_/II, and TT01α'_/II had classic secondary PV traits (Table 1).

We developed a new agar medium, the TreGNO (nutrient agar with trehalose and and bromothymol blue) medium, for color discrimination of TT01 PVs (see Materials and methods [below] for details). PVs produce green, convex, and mucoid colonies whereas secondary PVs produce yellow, flat, and nonmucoid colonies on this medium. TT01_/II and TT01α_/II colonies were homogeneous and had the colonial traits of secondary PVs. However, TT01α'_/II was composed of three CVs (TT01α' lineage; Figure 1). The first was a primary colonial form (green, convex, and mucoid colonies), named REV because it resembled a revertant colony, exhibiting primary PV traits (although bioluminescence, pigmentation, and crystal production were not completely restored; Table 1). The second was a secondary colonial form (yellow, flat, and nonmucoid colonies), named VAR because of its secondary PV traits (Table 1). The third form had small, green, convex, and mucoid colonies, and was named INT because of its intermediate traits or traits from both the primary and secondary PVs (Table 1). These CVs are unstable because each individual TT01α'_/II colony grown in liquid broth gives rise to a mixture of the three colonial forms on TreGNO medium. We generated a stable secondary PV from the VAR colonial variant by plating a liquid subculture from an individual VAR colony on nutrient agar and picking another VAR colony for a new cycle of liquid/plate culture. We continued this enrichment process until the liquid subculture generated 95% of VAR colonies on TreGNO plates. The stable population was named VAR* (Figure 1).

We PCR-amplified the lopT1 5' region from TT01_/II, TT01α_/II, TT01α'_/II, VAR*, and REV (Additional data file 1). The lopT1 deletion was only present in the TT01α and TT01α' lineages.

We injected TT01_/I, TT01_/II, TT01α_/I, TT01α_/II, and VAR* into Spodoptera littoralis larvae to evaluate the pathogenicity of these variants in insect larvae. TT01_/II, TT01α_/I, and TT01α_/II had the same level of pathogenicity as TT01_/I; 50% mortality (LT₅₀) was reached between 28 and 32 hours after injection for the TT01 wild-type strain and these three variants. By contrast, VAR* had a delayed LT₅₀ of 53 hours, although 100% mortality was reached at 3 days after infection (Figure 2).

We examined the whole genome architecture of each variant using I-CeuI genomic macrorestriction and pulsed field gel electrophoresis (PFGE) in order to detect large rearrangement such as deletions and amplifications by recombination between rrn or deletions, amplifications, and translocations inside I-CeuI fragments. I-CeuI is an intron-encoded enzyme that specifically cleaves a 26-base-pair site in the bacterial 23S rRNA gene. The PFGE pattern obtained for the TT01_/I strain matched the pattern of I-CeuI fragments predicted from the complete TT01_/I genome sequence (Figure 3a, b; also see Additional data file 2 for the details of the gels). Using the TT01_/I pattern used as a reference, we observed large genomic rearrangements in TT01α_/I, TT01α_/II, TT01α'_II, VAR*, and REV. PFGE patterns revealed identical profiles for primary and secondary PVs within both TT01 and TT01α lineages (Figure 3b and Additional data file 2). Therefore, PV status (primary versus secondary) in these variant lineages is independent from global genomic architecture.

Cluster analysis of the seven observed I-CeuI patterns reveals that variant lineages are in fact genomic lineages (Figure 3c). The TT01 and TT01α lineages exhibit genomic homogeneity. The TT01α' lineage shared common genomic features with the TT01α lineage, but exhibited a more polymorphic genomic pattern than TT01 and TT01α lineages.

The PFGE patterns of TT01α and the TT01α' lineages only reveal six apparent I-CeuI fragments, instead of seven fragments in the TT01_/I reference chromosome; however, the intensity of the 295-kb band suggests that it may represent two different fragments. We used Southern blot analysis to confirm that the seven rrn copies are present in all the variants (Additional data file 3). Therefore, variation in I-CeuI PFGE patterns among the TT01 variants appeared to be unrelated to deletion or amplifications mediated by recombination between rrn operons.

Additionally, the 465 kb faint band in the TT01α'_/II pattern (white star in Additional data file 2) corresponded to a fragment in the REV pattern, suggesting the existence of a 'REV-like' chromosome subpopulation in TT01α'_/II.

Large genomic rearrangements were present in the TT01α and TT01α' lineages. We further evaluated the nature of these rearrangements by comparing gene content between representative variants of each lineage, TT01_/I, TT01α_/I and VAR*, using genomic DNA hybridization on a P. luminescens TT01_/I microarray.

Totals of 159 and 162 genes were absent from TT01α_/I and VAR*, respectively (see Additional data file 4). We located these genes on a circular map of the TT01_/I chromosome (Figure 4); they mostly clustered into eight regions absent from both the TT01α_/I and VAR* genomes (regions A, C, D, E, F, G, I, and J) and one region specifically absent from the VAR* genome (region H). The deleted regions were located throughout the chromosome, with no particular symmetry around the replication origin or termination site. Several regions displayed a GC bias inversion (C, D, E, G, I, and J). Three overlapped with phagic regions (C, G, and I), suggesting that prophage excision occurred during the TT01_/I to TT01α_/I transition (Table 2). As well as phagic genes, the deleted regions encompass putative mobile and recombination-mediating elements such as insertion sequences and recombination hotspot (Rhs) elements (region A), and plasmid-related protein-encoding genes (region J)(Table 2). The regions C, D, E, and F potentially encode peptide synthetases involved in antimicrobial compound synthesis (Table 2). However, we did not observe any significant difference in antimicrobial activity between TT01_/I and TT01α_/I tested for 14 indicator strains (data not shown).

A more thorough analysis of hybridization ratios revealed that 122 genes had a ratio higher than 1.4 in the VAR* genome (Additional data file 5). In contrast, comparison of the TT01_/I and TT01α_/I genomes revealed only four genes with a ratio higher than 1.4. These findings suggest that numerous genes are amplified in the VAR* genome. Among these potentially amplified genes, 112 are clustered in a unique and large 275-kb region, named B. This region encompasses 4.8% of the TT01_/I genome (from plu0769 = mrfA to plu0980 = hpaA; Figure 4). Region B is located within the first quarter of the TT01_/I chromosome and is not delimited by obvious repeat elements. According to TT01_/I genome annotations, the region B may be involved in numerous and different functions (Table 2): basal cellular functions involving the DNA polymerase III ε chain (plu0943 = dnaQ), enolase (plu0913 = eno), and proteins involved in tryptophan metabolism (plu0799 = tnaA; plu0800 = mtr); and environment and/or host interactions, involving the major fimbrial biosynthesis locus (plu0769-0778 = the mrfABCDEFGHJ operon), insecticidal toxin proteins (plu0805 = tccA3; plu0806 = tccB3; plu0960 = tcc2; plu0961 = tcdB1; plu0962 = tcdA1; plu0964 = tccC5; plu0965 = tcdA4; plu0970 = tcdB2; plu0971 = tcdA2), and proteins similar to pyocins (plu0884; plu0886-0888; plu0892; plu0894).

To determine whether DNA microarray experiments explain the architectural modifications observed by macrorestriction experiments, we compared the two sets of data. The observed I-CeuI macrorestriction fragments from the TT01α lineage (36 kb, 295 kb, 295 kb, 330 kb, 465 kb, 610 kb, ~3600 kb) were similar to the theoretical I-CeuI fragments calculated after size subtraction of the eight deleted regions from the TT01_/I I-CeuI fragments (36 kb, 244 kb, 266 kb, 330 kb, 462 kb, 627 kb, ~3478 kb). Therefore, large-scale deletion events appear to underlie the TT01 to TT01α lineage transition. DNA microarray experiments in the TT01α' lineage identified a 275 kb amplification of the TT01_/I genome. Duplication or triplication of region B may account for the increase in genome size (~100 kb to 650 kb) observed by macrorestriction for the TT01α to TT01α' transition. Therefore, duplication appears to be mainly responsible for the TT01α to TT01α' lineage transition.

We first examined the genomic deletions observed in the TT01α_/I and VAR* variants. We focused on region H, which, by contrast to other deleted regions, did not exhibit typical recombination-mediating elements. Probes targeting different parts of the region H were hybridized on genomic DNA of the wild-type strain and the six variants. Hybridization patterns were identical within each variant lineage and confirmed the presence of a 25 kb deletion within the region H (from plu3237 to plu3253) for the TT01α' lineage (data not shown). Southern analysis also indicated the presence of a small deletion of about 10 kb (from plu3238 to plu3248) in the TT01α lineage. To map the deletion borders accurately, primers flanking the 25-kb deletion (R-3236 and F-3254) and the 10-kb deletion (R-3238bis and F-3249) were designed (Figure 5) and used for PCR amplification in the TT01α' and TT01α lineages. Amplified fragments of 4.8 kb and 5.2 kb were observed (data not shown). These fragments were sequenced for TT01α_/I and VAR*, and the deletion was physically mapped (a genetic map of the region H is presented in Figure 5). The deletions in TT01α_/I and VAR* were 12,820 bases (from coordinates 3,833,904 to 3,846,723) and 25,140 bases long (from coordinates 3,830,001 to 3,855,140), respectively.

We used Nosferatu, software that can detect approximate repeats in large DNA sequences 46. The region H is rich in pairs of repetition units (RPT) larger than 1 kb (Figure 5). Each deletion began at the right-hand extremity of the first repetition and finished at the right-hand extremity of the corresponding second repetition (RPT179385 repetitions for the 10-kb deletion and RPT179383 repetitions for the 25-kb deletion). Therefore, successive deletions mediated by homologous recombination between RPT are likely to have occurred in the region H during the TT01_/I to TT01α_/I to VAR* transition, leading to genomic reduction.

In a second set of analyses, we focused on the gene amplification observed in region B, occurring in the TT01α_/I to VAR transition. Quantitative PCR was performed for two genes in region B, mrfA (plu0769) and dnaQ (plu0943). Comparison of VAR and TT01α_/I data confirmed that these two genes were duplicated in the VAR* genome (Figure 6a).

In order to determine whether region B is duplicated specifically in the VAR* variant or in all variants of the TT01α' lineage, a probe covering the entire region B (the probe B) was prepared and hybridized to genomic DNA of the wild-type strain and the six variants. According to the TT01_/I genome sequence, NotI hydrolysis generates 25 fragments with a unique 1,056-kb fragment containing region B. Hybridization of the probe B to NotI-hydrolyzed genomic DNA generated a unique fragment of 1,056 kb in the TT01 lineage and of 1,020 kb in the TT01α lineages (Figure 6b, c). By contrast, in the TT01α' lineage, the B probe hybridized to the 1,020-kb fragment and an additional fragment. This second fragment has a similar size in TT01α'_/II and VAR* variants (610 kb) but is smaller (365 kb) in the REV variant. These findings showed that duplication of region B occurred in all TT01α' lineage variants.

Region B encompasses 275 kb in the TT01_/I genome sequence; thus, we determined whether the resulting amplified genes were dispersed in the genome or co-localized in an unique block. The unique additional fragment detected by the probe B in the TT01α'_/II and VAR* variants indicated that the product of the region B amplification is constituted either of one block or a few blocks co-localized in a genomic region whose size is smaller than 610 kb in TT01α'_/II and VAR* and smaller than 365 kb in REV. The probe B was also hybridized to ApaI-hydrolyzed genomic DNA of the wild-type strain and the six variants. The seven patterns were identical and the probe B hybridized with the two main 74 and 156 kb fragments covering the major part of region B according to the TT01_/I genome reference sequence (data not shown). Because the duplication did not modify the ApaI restriction pattern, we concluded that region B was amplified as a single block.

Our study provides the first extensive investigation into genomic rearrangements in Photorhabdus variants. First, we evaluated phenotypic traits of the three variant lineages (Figure 1). The TT01 lineage is derived from the TT01_/I strain, which was isolated from the H. bacteriophora TH01 nematode collected in Trinidad in 1993 43 and whose genome is sequenced 42. The TT01α lineage is derived from the TT01α_/I genomic variant, which was collected from H. bacteriophora TH01 maintained and multiplied in the laboratory. The TT01α' lineage was derived from the TT01α_/I variant after prolonged culture in synthetic medium. Each lineage is composed of PVs, whereby the primary form is characterized by the presence of typical phenotypic traits that are absent from the secondary form. The TT01α' lineage has an additional level of complexity, because the PVs exhibit features of CVs such as unstable morphotypes.

We then examined the genomic architecture of each variant in macrorestriction experiments and used comparative DNA microarray hybridization experiments to analyze the genomic content of representative variants for each lineage. Our findings revealed that large genomic rearrangements characterize each variant lineage. Consequently, these findings provide insight into probable scenarios underlying each lineage transition. The whole-genome organization of the TT01 lineage is described by the TT01 reference genome 42. Large-scale deletion events in the TT01 flexible gene pool seem to be involved in the TT01 to TT01α lineage transition. Deletion events in the TT01 flexible gene pool and a single block duplication encompassing 4.8% of the TT01 reference genome appear to underlie the TT01 to TT01α' lineage transition. The genomic clusters do not depend on the PV status (see below). Thus, each variant lineage is a genetic lineage.

To explain the molecular mechanisms involved in the rearrangements in our variants, we investigated potential repetitive elements and recombination-mediating elements flanking the rearranged regions. Large genomic architectural changes are often driven by homologous recombination between repeated sequences. The nature of the change then depends on the relative orientation, size, and spacing of the repeated sequences 47484950. Recombination events often occur at the rrn operon in Gram-negative bacteria, such as Salmonella, Rhizobium, Escherichia coli, and Ochrobactrum 1113185152. However, despite the variation detected in PFGE analysis of the rrn skeleton for the three variant lineages, we demonstrated that the rearrangements are not the result of rrn recombination.

Apart from homologous recombination, rearrangements can be induced by site-specific recombination, associated with recombination-mediating elements such as mobile elements, or by illegitimate recombination, linked to shortly spaced repeats 4950. Most of the deleted regions in the TT01α lineage are rich in potential rearrangement-mediating elements, with both repeated sequences - including insertion sequences and Rhs elements - and mobile elements, including phagic and plasmid-related genes.

Genomic annotation of the region H, which underwent successive deletions in the TT01α_/I and VAR* variants, did not describe the presence of typical repetitive or recombination-mediating elements. The region H belongs to a large genomic island containing the genes vgr and hcp, initially described as genes associated with Rhs elements. Rhs elements are repeated sequences in the E. coli genome that mediate major chromosomal rearrangements 515354. Although the TT01_/I genome contains Rhs-like elements 42, no Rhs element is located in the genomic island encompassing the region H. Nevertheless, we identified pairs of approximate long repeated sequences (>1 kb) in direct orientation (RPT) that corresponded to the observed deletion junction points. Therefore, successive deletions in the region H are likely to have been mediated by homologous recombination between RPT during the transition from TT01_/I to TT01α_/I to VAR*, leading to genomic reduction.

There was a strong selective pressure during the TT01α to TT01α' lineage transition (3 months in LB broth without shaking). This environmental constraint could thus be responsible for the rearrangement leading to the region H deletion. However, the region H deletion was already initiated during the former transition (TT01_/I to TT01α_/I in the laboratory-maintained nematode). Therefore, the observed reduction genomic size is more likely to be the result of particular genomic features (the RPT) rather than environmental constraints.

The region H is unique in the TT01_/I genome. Nevertheless, some RPT elements have similarities with sequences elsewhere in the TT01_/I genome, in the Photorhabdus strain W14 genome 55 or in other Enterobacteriaceae genomes such as Yersinia pseudotuberculosis IP32953 (BX936398.1), Yersinia pestis Angola (CP000901.1), Yersinia pestis Pestoides F (CP000668.1), Yersinia pestis CO92 (AL590842.1), Yersinia pestis biovar Microtus str. 91001, (AE017042.1, Yersinia pestis Antiqua (CP000308.1), Yersinia pestis Nepal 516 (CP000305.1), Yersinia pestis KIM (AE009952.1), Yersinia pseudotuberculosis IP 31758 (CP000720.1), and E. coli CFT073 (AE014075.1). Therefore, we propose that the region H represents a new type of bacterial recombination hot spot, which is vgr- and hcp-rich, but lacks Rhs elements.

We described a single block duplication (region B) targeting a 275-kb region of the TT01_/I genome in the TT01α' lineage. This significant duplication encompasses 4.8% of the TT01_/I genome. Region B is not located near the replication origin or termination and does not correspond to genomic islands or enterobacterial variable regions previously identified 4256. GC content or GC skew deviations are not evident.

Gene amplifications can occur through three kinds of known mechanism: homologous recombination between direct repeats, illegitimate recombination, or escape replication. No repeated elements flanking region B were detected, despite the use of the Nosferatu software 46, excluding the possibility of homologous recombination underlying this duplication. Region B duplications may result from illegitimate recombination between short repeats 475758. However, amplification copy number resulting from illegitimate recombination events is often high, even for large amplicons, such as in Acinetobacter sp. ADP1 or Streptomyces kanamyceticus 5960. Escape replication involves amplification of large regions of the host genome (several hundred kilobases), next to phage integration sites after induction of the phage lytic cycle 61626364, or around degraded prophages without the induction of specific phage lysis 65. Although phage remnants represent 4% of the Photorhabdus genome 42, lytic phages have not been identified in Photorhabdus strains, even after extensive investigation of lytic induction conditions 66. We detected the presence of an 11-kb phagic segment (plu0818-plu0826) in region B, potentially representing a degraded prophage. However, whereas the copy number usually resulting from the escape replication mechanism ranges between three and ten, with its intensity decreasing symmetrically from the center, region B in the TT01α' lineage genomes represents a single block homogeneous duplication. We only identified one other previously reported example of a large duplication without repeated flanking sequences - a 250 kb duplication in Mycobacterium smegmatis mc² 155 genome 67. Therefore, the duplication of region B is likely to belong to a new class of duplications.

Large genomic changes such as deletions and duplications are supposed to have important fitness effects. In our study, we firstly demonstrated that the PV status (primary or secondary) is independent from global genomic architecture. This was consistent with previous studies analyzing specific genetic regions 283839 and with partial genome studies 334041, but this is the first time it has been demonstrated using a whole-genome approach.

We showed that the overall genomic pattern corresponds to the variant lineage. Both the phenotype and pathogenic traits of the primary PV (or the secondary PVs) are indistinguishable between the TT01 and TT01α lineages. Therefore, changes in the genomic architecture of these strains did not lead to observable changes in the phenotype. Furthermore, certain regions that were deleted in the TT01α lineage potentially encode biosynthesis pathways for antimicrobial compounds. However, we did not observe any difference in antimicrobial activity between TT01_/I and TT01α_/I. This finding suggests that some TT01_/I genes are redundant. Indeed, genes encoding proteins potentially involved in the biosynthesis of antimicrobial compounds are over-represented in TT01_/I genome 42. Moreover, the encoded proteins in the deleted regions may be adaptive factors required for specific conditions that are not encountered in the laboratory or in our antibiosis assays.

The TT01α' lineage differs from the two other lineages due to its polymorphic genomic pattern. Furthermore, this lineage is composed of three unstable CVs and the virulence of the stabilized VAR* variant is attenuated in insects. This is consistent with previously reports of CVs isolated from the Photorhabdus genus 33. Therefore, changes in genomic architecture might be correlated to phenotypic changes in variants of this lineage. The main rearrangement observed in the TT01α' lineage is the region B duplication. According to TT01_/I genome annotation, region B may be involved in both basal cellular functions and environment and/or host interactions. Gene duplication events can underlie modification of phenotypes 58. However, we did not detect any modification of gene transcription in region B using transcriptomic microarray comparison between the VAR* and TT01α_/I variants (Gaudriault S, unpublished data). Thus, this duplication does not appear to modify gene expression in the VAR* variant. Therefore, the attenuation of virulence of the VAR* variant is not likely to be due to amplified expression in region B. Rather, it is more likely that the 'cost' to the bacteria of the increased genome size is decreased virulence in insects.

We conclude that the observed phenotypes and overall genomic architecture are not systematically correlated in TT01, TT01α, and TT01α' lineages. It is likely that this result is general in the field of bacterial genomic architecture. Similar observations were previously made between strains of the Pseudomonas aeruginosa species 68, but also inside a clonal bacterial population of a wide range of bacterial groups such as Yersinia pestis 19, Pseudomonas aeruginosa 17, and Sinorhizobium meliloti 16.

Do large genomic rearrangements occur randomly or are they shaped by drastic selective evolutionary forces? Several years of comparative genomics between whole bacterial genomes showed that the prokaryotic genome is a heterogeneous entity, with regions of stability and flexibility 44950. Genomic stability is subject to selective pressures such as functional replication 69, gene essentiality 70, or translation 71. The three main routes of evolution of genome repertoire are lateral gene transfer, when several bacterial communities share a same ecological niche, deletions, and duplications 44950. The dynamism of genome repertoire inside a clonal population only arises by the last two phenomena, as illustrated by our study on Photorhabdus variants.

In E. coli, the chromosome is organized in structured macrodomains, limiting genome plasticity. Whereas some genomic rearrangements between these macrodomains have only moderate effects on cell physiology, others have detrimental effects 72. The rearrangements that we observed in our variants may have been selected to preserve chromosomal configurations that are not detrimental for bacterial fitness in the laboratory or in the nematode. We believe that structured macrodomains that restrict chromosome plasticity are likely to exist in other bacterial genus. Identification of structured macrodomains in P. luminescens genome would provide better knowledge on evolutionary forces modeling bacterial genome.

The major genomic variations described in TT01 variants have cryptic consequences in our laboratory conditions. The absence of associated phenotypes makes them difficult to identify, explaining why such genomic variations are rarely observed. However, further studies of such genomic variations may be crucial for a better understanding of bacterial adaptation and evolution.

Indeed, we observed that the extensive genomic rearrangements in Photorhabdus variants were often associated with several genomic subpopulations in the same culture. Similar observations were previously made for a P. luminescens TT01_/I locus encoding a phage tail-like structure 73 and the mrf locus of the P. temperata strain K122 74. In Sinorhizobium meliloti, Yersinia pestis, and Pseudomonas aeruginosa, extensive variations of genome architecture, without obvious changes in phenotype, were also observed during bacterial growth in broth medium 161719. Different pre-existing chromosomal forms in a clonal bacterial population are likely to give this population an adaptive capacity. It is therefore possible that bacterial populations maintain various subpopulations with different genomic structures as a way to cope with different environments during its life cycle.

Additionally, deletion events in TT01α and TT01α' lineages are located within the TT01_/I 'flexible' gene pool. Whereas intragenomic recombination in the 'flexible' gene pool have been widely studied using comparative genomics for different bacterial genera, species, and strains 1234, similar reports for clonal variants are rare. Gene repertoires of the 'flexible' gene pool may evolve through variations in bacterial subpopulations and then become fixed after bacterial speciation. Such pre-existing or currently existing genomic variations have an important role in evolutionary patterns of natural eukaryotic populations 75. They may also have a determinant role in bacterial evolution.

All bacterial strains and plasmids used in this study are listed in Additional data file 6. Primers are listed in Additional data file 7. P. luminescens was grown at 28°C in LB broth or on nutrient agar 1.5% (BD Difco™, Franklin Lakes, New Jersey, USA) for 48 hours. Escherichia coli was grown at 37°C in LB broth or on LB supplemented with 1.5% agar (BD Difco™, Franklin Lakes, New Jersey, USA). Strains were stored at -80°C in LB broth containing 16% glycerol (vol/vol). Secondary variants were obtained by prolonged culture of primary variants at 28°C for 10 days in Schneider's insect medium (Cambrex Bio Science, Walkersville, Maryland, USA) with shaking (TT01_/II 30), for 10 days in LB broth with shaking (TT01α_/II), or for 3 months in LB broth without shaking (TT01α'_/II). Secondary variant phenotypes were evaluated from culture on NBTA (nutrient agar 1,5%, 25 mg/l bromothymol blue and 40 mg/l triphenyl-2,3,5-tetrazolium chloride) plates and on TreGNO plates (see below) at 28°C. Secondary variants were identified by performing phenotypic tests as previously described 76 and controlled by PCR-restriction fragment length polymorphism of the 16S rRNA gene 77.

Xenorhabdus and Photorhabdus secondary variants are typically selected on NBTA plates to distinguish red secondary variants colonies from blue primary colonies 76. Because of the high level of pigmentation of Photorhabdus colonies, the use of color assays does not allow clear distinction between primary and secondary variants for Photorhabdus genus. We found that TT01 secondary variants were able to undergo trehalose fermentation, whereas primary variants can not. On nutrient agar plates supplemented with trehalose (10 g/l) and bromothymol blue (25 mg/l), secondary colonies acidified the bromothymol blue and became yellow at 28°C after 48 hours. Primary colonies remained green. Furthermore, secondary colonies were flat and large with irregular borders. This new medium was named TreGNO medium and was routinely used for the discrimination of Photorhabdus luminescens strain TT01 phenotypic variants.

Intact genomic DNA was extracted in agarose plugs as follows. Bacterial cells grown on nutrient agar plates were suspended in phosphate-buffered saline (GIBCO^® Invitrogen, Carlsbad, California, USA) to a turbidity of 1.25 at 650 nm, included in 1% (vol/vol) low melting agarose (SeaPlaque^® GTG, FMC BioProducts, Rockland, Massachusetts, USA) solution and then subjected to lysis as described previously 78.

NotI and ApaI hydrolysis were performed by incubation of the agarose plugs overnight with 40 units of the endonuclease in buffer recommended by the supplier (New England Biolabs, Hertfordshire, UK), at 37°C for NotI and 25°C for ApaI. PFGE was carried out in a contour-clamped homogeneous field electrophoresis apparatus CHEF-DRII (Bio-Rad, Hercule, California, USA) in a 0.8% agarose gel in 0.5× Tris-borate-EDTA (TBE) at 10°C. PFGE conditions were as follows: for NotI fragments, a 35 to 5 second pulse ramp for 47 hours followed by a constant pulse time of 50 seconds for 6 hours at 4.5 V/cm; and for ApaI fragments, 35 to 5 seconds for 35 hours, followed by 5 seconds to 1 second for 4 hours at 4.5 V/cm.

I-CeuI hydrolysis was performed as described previously 79. For the separation of I-CeuI fragments, different electrophoresis conditions were selected according to fragment size: a pulse ramp from 5 to 50 seconds for 24 hours at 6 V/cm for fragments with size below 700 kb; and a pulse ramp from 150 to 400 seconds for 45 hours at 4.5 V/cm for I-CeuI fragments for fragments between 700 kb and 1 megabase. For I-CeuI fragments larger than 1 megabase, PFGE was performed on Rotaphor apparatus (Biometra, Goettingen, Germany) using 0.7% agarose gels in 0.5× TBE buffer. The electrophoresis conditions used were as follows: 50 to 47 V (linear ramp), 6,000 to 1,000 seconds decreasing pulses (logarithmic ramp), with a increasing angle from 96 to 105°, buffer temperature 11°C, for 240 hours. I-CeuI PFGE patterns were compared by calculating the Dice coefficient for each pair 80. Patterns were clustered by UPGMA using the Phylip program package 81.

HindIII-hydrolyzed DNA was subjected to electrophoresis for 3 hours at 2.6 V/cm in a 0.8% agarose gel in 0.5× TBE using SubCell apparatus (Bio-Rad) 13.

Electrophoresis gels were transferred onto a Nytran N SuperCharge nylon membrane (Schleicher and Schuell, Dassel, Germany) by vacuum blotting in 20 × SSC (Euromedex, Souffelweyersheim, France).

A digoxigenin-labeled probe targeting 16S rRNA gene was obtained by PCR from genomic DNA of P. luminescens strain TT01_/I, using primers 27f and 1492r with a dNTP mixture containing 0.1 mmol/l digoxigenin-dUTP 13.

Probes B and H were obtained using respectively small fragment insert from plg2711 and large fragment inserts from plbac4g08, plbac6h12, plbac3a10, plbac3c04, and plbac2f12. Fragment inserts were purified, sonicated into fragments of between 1 and 10 kb if insert size was higher than 10 kb, and labeled with digoxygenin by random priming (Dig DNA labeling Kit; Roche, Meylan, France). Hybridization of the probes was detected using a CSPD chemiluminescent system (Roche).

Genomic DNA was extracted as previously described 56 and stored at 4°C. We PCR-amplified the lopT1 deletion region with Taq polymerase (Invitrogen, Carlsbad, California, USA), in accordance with the manufacturer's recommendations, using the PlopT1.fw an PlopT1.rev primers. The region H was amplified by PCR with the Herculase Enhanced DNA polymerase (Stratagene, Amsterdam Zuidoost, Pays Bas), in accordance with the manufacturer's recommendations, using the R-3236, F-3249, R-3238bis, and F-3254 primers. For sequencing region H deletions, we purified the 4.8 kb and 5.2 kb fragments using the Montage PCR kit (Millipore, Guyancourt, France) and sequenced using PCR primers and chromosome walking (Millegen, Toulouse, France). Sequencing of the 5.2 kb fragment central region of the fragment failed probably because of the presence of repetitions. A 3.2 kb central region was therefore amplified by PCR with PstIdMutF and XbaIdMutR primers. The amplicon was hydrolyzed by PstI and XbaI, ligated into PstI- and XbaI-hydrolyzed pUC19, and inserted into E. coli XL1blue by transformation. The resulting plasmid was purified by Nucleobond AX-100 kit (Macherey-Nagel, Hoerd, France), and the insert was sequenced with PstIdMutF and XbaIdMutR primers and then by chromosome walking.

DNA microarray hybridization and analysis were performed as previously described 56.

Quantitative PCR was performed in triplicate using the LightCycler FastStart DNA Master^PLUS SYBR Green I kit from Roche Diagnostics with 1 ng genomic DNA and 1 μmol/l specific primers targeting fliC (L-1954 and R-1954), mrfJ (L-0778 and R-0778), dnaQ (L-0943 and R-0943), and pilN (L-1051 and R-1051). The enzyme was activated for 10 minutes at 95°C. Reactions were performed in triplicate at 95°C for 5 seconds, 60°C for 5 seconds and 72°C for 10 seconds (45 cycles), and monitored in the Light Cycler (Roche). Melting curves were analyzed for each reaction; all reactions exhibited a single peak. The amount of PCR product was calculated with standard curves obtained from PCR with serially diluted TT01_/I genomic DNA. All data are presented as ratios, with gyrB (primers L-0004 and R-0004) as a control (95% confidence limits).

Sequence annotation of the TT01_/I genome was obtained from the MaGe database 82. We evaluated amino-acid and nucleotide similarity using BLASTP and BLASTN software 83. We used Repseek software, previously Nosferatu 46, to detect approximate repeats in large DNA sequences.

In vivo infection assays were performed as previously described 45. We performed three independent experiments for each variant. Statistical analysis were performed as previously described 84.

Antibiosis assays were performed as previously described 76 with the following bacterial species: Micrococcus luteus, Staphylococcus epidermidis CIP 6821, Staphylococcus aureus CIP 7625, Escherichia coli CIP 7624, Proteus vulgaris CIP 5860, Pseudomonas aeruginosa CIP 76.110, Corynebacterium xerosis, Ochrobactrum intermedium LMG 3301^T, Ochrobactrum anthropi ATCC 49188^T, Ochrobactrum sp. FR49, Erwinia amylovora CFBP1430, Pseudomonas sp. BW11M, Salmonella enterica 14028s, and Yersinia enterocolitica serotype 08.