The acrystalliferous B. thuringiensis 407 Cry^- strain (Bt 407^-) (Lereclus et al., 1989) was used as the parental strain to create all the strains used in this study. E. coli strain DH5α (Taylor et al., 1993) was used as the host strain for plasmid construction. E. coli strain ET12567 (MacNeil et al., 1992) was used to prepare DNA prior to electroporation in B. thuringiensis. Cells were grown in LB medium (1% tryptone, 0.5% yeast extract, 1% NaCl) or HCT medium (0.7% casein hydrolysate, 0.5% tryptone, 0.68% KH2PO4, 0.012% MgSO4 7H2O, 0.00022% MnSO4 4H2O, 0.0014% ZnSO4 7H2O, 0.008% ferric ammonium citrate, 0.018% CaCl2 4H2O, 0.3% glucose, pH 7.2) (Lereclus et al., 1982) at 37°C and stored at -80°C in LB containing 15% glycerol.

For B. thuringiensis cultures, t0 corresponds to the beginning of the transition between the exponential and stationary phases of growth.

The antibiotic concentrations used for selection of B. thuringiensis and E. coli were as follows: erythromycin, 10 μg/mL; ampicillin, 100 μg/mL. Chloramphenicol was used at a concentration of 100 μg/mL to block the synthesis of proteins (Periago et al., 2002) in B. thuringiensis.

When required, xylose was used at a concentration of 20 mM.

DNA manipulations are detailed in the Supplementary Experimental Procedures. All the plasmids and strains used in this study are indicated in Tables 1, 2. Oligonucleotides are listed in Supplementary Table S1.

To assay the GFP-based fluorescence of cells harboring the _sfgfp gene and its derivatives designed to improve fluorescence, overnight cultures incubated at 30°C in HCT medium supplemented with erythromycin were diluted 1000-fold in HCT and incubated at 37°C under agitation until an OD₆₀₀ of 0.5. Xylose was then added and growth was resumed. Cells were harvested at the time of xylose addition (T0) as well as at other time points after xylose addition (Tn). Cultures were carried out in the same way to assay the GFP-based fluorescence of cells harboring the gfp_Bte gene and its derivatives designed to destabilize the GFP, except that chloramphenicol was added to the cells 1 h after xylose addition.

In all cases, the cells were harvested and fixed as described in Verplaetse et al. (2015). Essentially, the cells were centrifuged, fixed for 7 min in PBS-formaldehyde 4% then washed in PBS. The pellet was then resuspended in GTE buffer (Vlamakis et al., 2008) and kept at 4°C until flow cytometry analysis or microscopy.

Intrahemocoelic injection experiments with G. mellonella were carried out essentially as described previously (Salamitou et al., 2000; Verplaetse et al., 2015). For each strain, 20 larvae were injected each with 2.10⁴ bacteria and kept at 30°C for 72 h. 18 h after injection, surviving insects were eliminated. At each time point, B. thuringiensis cells were harvested from 2 or 3 dead insects as follows: the larva was cut open, transferred to a 1.5 mL Eppendorf tube containing 1 ml of PBS and vortexed. The suspension was pipetted into a new 1.5 mL Eppendorf tube (leaving behind most of the large insect debris). The sample was centrifuged and the pellet was resuspended in PBS-formaldehyde 4%, fixed for 7 min then washed in PBS. This suspension was filtered onto a cotton pad in a 1 mL syringe in order to retain the cadaver debris and recover the bacterial cells in the filtrate. These were then concentrated by centrifugation, resuspended in GTE buffer (Vlamakis et al., 2008) and kept at 4°C until flow cytometry analysis or microscopy.

For the assessment of the percentage of live cells in the bacterial population in insect cadavers, the experiments were carried out in the same manner except that the cells were not fixed with formaldehyde. After filtration on the cotton pad, the suspension was centrifuged, resuspended in saline, incubated with the SYTOX Green Dead Cell Stain (Molecular Probes) according to the manufacturer’s instructions and analyzed immediately with the flow cytometer. Dead bacteria will present a bright green fluorescence compared to live cells.

Fluorescence was measured on a CyFlow Space cytometer (Partec, France). Details about the parameters used to collect fluorescence and the softwares used to analyze the data are given in the Supplementary Experimental Procedures. The different populations were identified using histograms or bi-parametric cytograms. GFP- or mCherry-expressing cells were identified as cells giving a higher signal intensity than the reporterless cells used as controls.

Cells were observed with an AxioObserver.Z1 Zeiss inverted fluorescence microscope equipped with a Zeiss AxioCam MRm digital camera and with Zeiss fluorescence filters. GFP was imaged using the 38 HE filter (excitation: BP 470/40, beam splitter: FT 495, emission: 525/50). mCherry was imaged using the 45 HE filter (excitation: BP 590/20, beam splitter: FT 605, emission: 620/14). Images were processed using the ZEN software package.

The DNA sequence of the plasmid containing the gfp_BteAAV sequence was submitted to GenBank and is available under the accession number MF673728.

To facilitate the analysis of gene expression in B. thuringiensis, we engineered a highly expressed version of the sfGFP. We used the reporterless _sfgfp (Pedelacq et al., 2006) under the control of the xylose-inducible PxylA promoter as a template for our improvement procedure. _sfgfp encodes a version of the Aequorea victoria GFP that has been shown to be four times brighter and that folds 4 times faster in vivo than GFPmut3 (Milde, 2008). We modified the _sfgfp gene and the features required for its translation to adapt it to B. thuringiensis. Our first step was to change the codon usage to one that would correspond better to that of our strain, using the GENEius software and the associated B. thuringiensis serovar thuringiensis codon usage table (Eurofins Genomics). We designated this allele gfp_Bt. We then modified the RBS present in the promoter region of PxylA to the improved RBS+, that resulted in a twofold increase in protein production in B. megaterium, as described in Stammen et al. (2010). Finally, we added the first 24 bp of comGA to the 5’ end of the _sfgfp coding sequence. The corresponding eight amino acids have been shown to enhance the stability of fluorescent proteins (Veening et al., 2004). The resulting cassette comprising the modified RBS and the comGA’gfp_Bt sequence was designated gfp_Bte. Using flow cytometry, we showed that the gfp_Bte-expressing cells grown in HCT medium were about 20 times brighter than the _sfgfp-expressing cells, 2 h after the addition of xylose (Figure 1A; median fluorescence of 34 and 1.6 AU and pink and orange lines, respectively). Codon adaptation resulted in the most efficient improvement compared to the RBS modification and the comGA sequence addition. Indeed, the gfp_Bt-expressing cells are fivefold brighter than the _sfgfp-expressing cells, whereas each subsequent modification step increased the fluorescence of the cells by approximately twofold (Figure 1A; median fluorescence of 8, 1.6, 22, and 34 AU and blue, orange, green and pink lines, respectively). This increase in fluorescence intensity is visible on the microscopy pictures taken 2 h after xylose addition (Figures 1B–F).

To monitor variations in gene expression, we decided to generate an unstable version of the GFP_Bte. We used a previously described method that takes advantage of the ssrA-mediated peptide tagging system (Andersen et al., 1998). The principle of the method is to tag a protein for specific degradation by a protease. We added 10 amino acids at the N-terminal end of GFP_Bte with a variation in the last 3 amino acids of the sequence (GKQNNLSLAA/-LVA/-AAV/-ASV), generating GFP_BteLAA, GFP_BteLVA, GFP_BteAAV, and GFP_BteASV. Amino acids GKQNNLSLAA correspond to the putative proteolytic tag added by the ssrA tmRNA we identified in the genome of strain B. thuringiensis 407. These amino acids have been shown to address the tagged protein to the Clp degradation machinery in E. coli and in B. subtilis (Keiler et al., 1996; Gottesman et al., 1998; Wiegert and Schumann, 2001). Each tag variant should alter the stability of the protein (Keiler and Sauer, 1996; Andersen et al., 1998). We monitored the fluorescence of the cells harboring the GFP_Bte tagged variants during growth in HCT medium and the results are presented on Figure 2. At the time of inducer addition, all the cells present a fluorescence similar to that of the reporterless cells (Figure 2A, left panel). 30 min after xylose induction, there is a striking difference between the cells harboring the wild-type -LAA or the -LVA tag and the cells harboring the -AAV or the -ASV tag (Figure 2A, second panel from the left). The latter present a fluorescence closer to that of the cells producing GFP_Bte whereas the fluorescence of the former is weaker. 1 h after xylose induction (Figure 2A, third panel from the left), the cells harboring the GFP_BteLAA or the GFP_BteLVA tag had a similar fluorescence (median fluorescence of 2 and 2.3 AU which is 33- and 28-fold lower than that of GfpBte-producing cells, respectively). The cells expressing gfp_BteASV were the most fluorescent (median fluorescence of 43AU, 1.5-fold lower than that of gfp_Bte-expressing cells), however, the histogram profile was not satisfying since it showed two peaks which indicate that gfp expression and/or GFP degradation in the population was highly heterogeneous. The cells harboring the GFP_BteAAV presented a median fluorescence of 19 AU that was 3.5-fold lower than that of the GFP_Bte-producing cells (median fluorescence of 65 AU) and a histogram profile similar to that of the latter. Two hours after addition of the inducer (Figure 2A, right panel), all the histograms were similar to those observed 1 h after induction. gfp_Bte-expressing cells were more fluorescent than 1 h before (median fluorescence of 125 AU vs. 65 AU, respectively) whereas the fluorescence of gfp_BteAAV-expressing cells was similar at both times (median fluorescence of 19 AU and 15 AU 1 and 2 h after addition of xylose, respectively). gfp_BteAAV was thus chosen as a reporter of gene expression for the subsequent experiments.

The decrease in fluorescence of GFP_BteAAV-producing cells was then monitored after the addition of chloramphenicol, an inhibitor of translation (Periago et al., 2002), to determine GFP stability (Figure 2B). The median fluorescence of gfp_Bte-expressing cells was similar at the time of and 1 h after chloramphenicol addition (median fluorescence of 73 AU and 77 AU, respectively) (Figure 2B, second and third panels from the left, respectively). In contrast, the median fluorescence of gfp_BteAAV-expressing cells decreased by 17-fold between these two time-points (median fluorescence of 26 AU and 1.5 AU, respectively). 22 h after chloramphenicol addition, the majority of the gfp_Bte-expressing cells remained fluorescent whereas the gfp_BteAAV-expressing cells presented a fluorescence close to that of the control cells (Figure 2B, right panel). These results indicate that, in these conditions, the GFP_BteAAV-based fluorescence shows a half-life of approximately 15 min.

In order to monitor the vegetative state of B. thuringiensis cells during the infectious cycle we chose to use the promoter of the abrB gene. This gene encodes a transition state regulator transcribed and active during the exponential phase (Perego et al., 1988; O’Reilly and Devine, 1997; Banse et al., 2008; Lucking et al., 2009). We monitored the fluorescence of the cells harboring a PabrB’gfp_BteAAV transcriptional fusion during growth in LB medium and the results are presented on Figure 3. The data are from two independent experiments. We represented 2 negative control samples on the graph to account for the difference between the auto-fluorescence of the reporter-less cells at t-2 and the cells harvested at the other time-points (Figure 3A, left panel). All the negative control histograms are shown in Supplementary Figure S1. The flow cytometry histograms show that the PabrB’gfp_BteAAV-expressing cells at t-2 are already fluorescent compared to the negative control (median fluorescence of 2.1 vs. 0.4 AU, respectively) and their fluorescence increases until it reaches a maximum at t-1 and t0 before decreasing at t1 and t2 (median fluorescence of 5.2, 5.8, 3.1, and 2.7 AU, respectively, to be compared to the light gray negative control with a median fluorescence of 0.8 AU) (Figure 3A, left panel). At t24, the cells harboring PabrB’gfp_BteAAV do not present any fluorescence (Supplementary Figure S2). The shape of the histograms suggests that the PabrB’gfp_BteAAV fusion is expressed in a homogeneous fashion in the population (only 1 peak is visible). The right panel of Figure 3A recapitulates the data for both experiments as kinetics of median fluorescence intensity and shows that the peak of PabrB’gfp_BteAAV expression was reached between t-1 and t0. Microscopy pictures of PabrB’gfp_BteAAV-expressing cells harvested at t0 and at t2 support the flow cytometry data (Figure 3B). These results show that the PabrB’gfp_BteAAV transcriptional fusion is expressed during exponential phase and that we can visualize a decrease in its expression when the cells enter the transition phase.

We investigated the physiological state of B. thuringiensis cells during infection in order to characterize in more detail the composition of the population during this cycle. To achieve this objective we examined the state of the cells that did not activate the promoters previously used, in particular the necrotrophic reporter (Verplaetse et al., 2015). We infected G. mellonella larvae with B. thuringiensis cells harboring the PnprA’mcherry_LGC transcriptional fusion as well as the PabrB’gfp_BteAAV transcriptional fusion on the same vector. nprA is under the control of NprR, the regulator of the necrotrophic state, and reflects its activity (Perchat et al., 2011). We monitored the fluorescence of the cells harvested from the insect cadavers at various times following infection. The results are presented in Figure 4. The kinetics profile of the Nec+ population (i.e., cells expressing nprA) is similar to what has been published before (Verplaetse et al., 2015) with a low percentage of the population expressing nprA 18 h post-infection (pi) (mean value of 11%) that increases between 24 and 48 h pi (mean values of 27 and 68%, respectively) to reach a maximum 72 h pi (mean value of 74%). We observed that the vegetative PabrB’gfp_BteAAV reporter is expressed at 18 h pi in 13% of the population. Interestingly, less cells expressed it at 24 h pi (mean value of 4%). The percentage of cells in which PabrB is expressed increased at 48 and 72 h pi (mean value of 15 and 22%, respectively). The expression of PabrB’gfp_BteAAV is almost exclusively restricted to cells that have activated the nprA gene promoter from 48 to 72 h pi. Figure 4 also shows that the proportion of cells that did not express the necrotrophic or the vegetative reporters changed from 71% at 18 h pi to 25% 72 h pi. In order to verify that the expression patterns observed were not due to loss or copy number heterogeneity of the plasmid during the infectious process, we infected G. mellonella larvae with B. thuringiensis cells harboring the pPaphA3’_sfgfp vector carrying a transcriptional fusion between the promoter of a constitutive gene in B. thuringiensis, and the reporter gene encoding _sfGFP. The results show that the fusion is expressed in a homogeneous fashion in the population 24 and 72 h pi (only 1 peak is visible) (Supplementary Figure S3). Furthermore, a previous study has reported that 100 and 99.3% of the bacteria harvested from insect cadavers still carried a vector with the same backbone 48 and 96 h post-infection, respectively (Verplaetse et al., 2015). This indicates that the phenotypic heterogeneity observed for the reporters mentioned above is not due to plasmid instability or copy number heterogeneity in B. thuringiensis during infection.

In order to verify that the bacteria that did not express any of the reporters used were alive, we infected G. mellonella larvae with B. thuringiensis cells harboring the PnprA’mcherry_LGC transcriptional fusion and incubated the samples harvested from the insect cadavers at various times following infection with the SYTOX Green Dead Cell Stain. We then immediately monitored the mCherry- and SYTOX Green-based fluorescence of the bacteria. The cells recovered at each time-point were mostly alive. Indeed 89, 79, and 84% of the bacteria were not stained by the SYTOX Green Dead Cell dye at 24, 48, and 72 h pi, respectively (Supplementary Figure S4). Figure 5A shows that in these conditions we recovered 78, 26, and 29% of Nec^- cells at 24, 48, and 72 h pi, respectively. In these samples, the SYTOX Green stain shows that 81, 69, and 62% of the Nec^- bacteria were live cells, respectively (Figure 5B). This indicates that the majority of the cells that did not express any of the reporters used were not dead.