A shvR mutant is attenuated in virulence in zebrafish embryos

We first analysed the role of B. cenocepacia K56-2 ShvR in virulence in the zebrafish infection model. Zebrafish embryos were micro-injected at 30 hours post-fertilization (hpf) with B. cenocepacia K56-2(mCherry_pCR11), ΔshvR(mCherry_pCR11) or the complemented ΔshvR(pshvR:shvR;mCherry_pCR11). Analysis of embryo survival showed that ΔshvR was unable to cause fatal infection over the time course of the experiment, in contrast to its wildtype parent which caused 100% embryo mortality in 3 days (Fig 1A and [30]). In agreement with the observed reduced host mortality there was a significantly lower bacterial burden of ΔshvR compared to K56-2 at 24 and 48 hours post infection (hpi) (Fig 1B). Although all embryos survived infection with ΔshvR, some contained significantly more bacteria at 24 and 48 hpi compared to T = 0 (Fig 1B), which suggests the mutant bacteria are able to replicate. Expression of shvR from its endogenous promoter introduced on a single copy plasmid restored virulence to the ΔshvR mutant (Fig 1). These results show that ShvR is required for development of acute fatal infection.

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Fig 1. ShvR is important for fatal infection in zebrafish larvae.

Zebrafish larvae were micro-injected with on average 143 ± 76 CFU of wildtype B. cenocepacia K56-2 (black), 118 ± 29 CFU of ΔshvR (red), and 58 ± 16 CFU of complemented ΔshvR (green), and embryo survival (A) and bacterial burden (B) were determined. The complemented strain harboured a low copy pCR11-derivative expressing shvR from its endogenous promoter. Open circles indicate dead embryos. Pooled results from three independent experiments are shown (n = 20 (A) and n = 5 (B), respectively, per group per experiment). Geometric means with each data point representing an individual embryo. A log rank (Mantel-Cox) test was used to determine significance of survival differences (A). Significance between groups was determined using one-way ANOVA with Sidak’s Multiple Comparisons test (B). * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, ns: not significant.

https://doi.org/10.1371/journal.ppat.1007473.g001

ShvR mutant bacteria persist and replicate in macrophages

Our previous work showed that macrophages efficiently phagocytose intravenously injected B. cenocepacia K56-2, and that they are critical for bacterial replication and ensuing fatal pro-inflammatory infection in zebrafish embryos [29]. Phagocytosis of bacteria from the blood results in infected macrophages distributed over the yolk and the tail region [30]. To analyse the interaction of ΔshvR with macrophages in the host, we quantified the percentage of ΔshvR bacteria that were internalised by macrophages using confocal imaging of infected Tg(mpeg:mCherry-F) transgenic embryos, which express a membrane localised mCherry protein (Fig 2A and S1 Movie). One hour after injection, ΔshvR bacteria were as efficiently phagocytosed by macrophages as wildtype bacteria (Fig 2B). Real time experiments showed that the ΔshvR bacteria still localised in macrophages at later time points, where they sometimes reached high numbers (Fig 2C) in agreement with the observed increase in CFU counts (Fig 1B). In contrast, at these later time points wildtype K56-2 bacteria have replicated to very high intracellular numbers, formed local infection sites (Fig 2D), re-entered the blood circulation and spread systemically as shown earlier [29]. Quantification of the number of intracellular ΔshvR bacteria in infected macrophages using Tg(mpeg:mCherry-F) embryos at 1 hpi and 24 hpi (Fig 2E) showed a significant decrease in the number of macrophages containing 1 or 2 bacteria and an increase in the number of macrophages containing more than 4 bacteria, respectively, demonstrating that the ΔshvR mutant is able to survive and replicate inside macrophages. The total number of infected macrophages that were observed per embryo decreased in time from 12.3 ± 2.1 (SEM; n = 15 embryos) at 1 hpi to 7.6 ± 0.9 (SEM; n = 10 embryos) at 24 hpi, suggesting that a proportion of ΔshvR bacteria can establish a replication niche, while others get degraded.

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Fig 2. ΔshvR mutant bacteria are able to persist and replicate in macrophages.

(A) Tg(mpeg1:mCherry-F) embryos were injected intravenously at 30 h post fertilization with ~50 CFU ΔshvR bacteria expressing eGFP (pIN301). Left panel: Confocal stack image over the yolk region of a representative embryo (macrophages marked in red) at 1 hpi. Scale bar 100 μm. Right panel: Detailed confocal (bright field and fluorescent overlay) image stack of two macrophages at 3 hpi containing 2 and 3 ΔshvR bacteria, respectively. Inset shows red and green channels only. Scale bar 10 μm. S1 Movie shows a video of the individual (21) sequential images. (B) Quantification of intracellular and extracellular bacteria in PFA-fixed embryos at 1 hpi over the yolk as indicated in Fig 2A using confocal microscopy. Each data point represents an individual Tg(mpeg1:mCherry-F) transgenic embryo injected intravenously with either wildtype B. cenocepacia K56-2 (black dots, n = 13) or ΔshvR (red dots, n = 15) expressing eGFP. Counts per embryo are presented as percentage of internalized bacteria relative to total numbers of bacteria in the yolk sac region. A total of 220 and 141 bacteria were counted for K56-2 and ΔshvR, respectively. Unpaired t-test, with mean 86.7 ± 5.3 and 90.6 ± 2.8, respectively, p-value 0.51. Data in (A, B) are representative of two independent experiments. (C) Tg(mpeg1:mCherry-F) embryos were injected with ΔshvR expressing eGFP. Images of an infected macrophage at 18 hpi over the yolk (left, bright field and fluorescent overlay image; enlarged inset, red and green overlay) and at 44 hpi (red and green overlay image) with high numbers of intracellular bacteria (representative of at least 5 embryos per treatment in more than 3 experiments). Scale bars, 12.5 μm. (D) Tg(mpeg1:mCherry-F) embryos were injected with K56-2 wildtype expressing eGFP. Image of an infection site at 24 hpi in the tail region (left, bright field and fluorescent overlay image; enlarged inset, red and green overlay) with high numbers of bacteria. Scale bars, 12.5 μm. (E) Quantification of the number of intracellular ΔshvR bacteria per macrophage in PFA-fixed embryos at 1 hpi (mean: 1.3 ± 0.05 bacteria per macrophage) and 24 hpi (4.1 ± 0.47 SEM), presented as frequency distribution histogram. The graph represents a total of 187 (15 embryos) and 80 macrophages (10 embryos), at 1 and 24 hpi respectively. Mann Whitney test p < 0.0001. Representative of 2 experiments.

https://doi.org/10.1371/journal.ppat.1007473.g002

In contrast to wildtype K56-2, which disseminates from initially infected macrophages at around 10–12 hpi culminating in acute fatal infection [29,30], ΔshvR did not spread throughout the embryos. Instead, the infection was characterised by low overall bacterial burden and the absence of tissue inflammation as seen for K56-2 and the complemented mutant (Fig 3A). However, real time observations showed that small infection sites with multiple infected cells sometimes developed from single infected macrophages. Although we have been unable to quantify this event due to the cell dynamics of the in vivo infection with immune cells moving around, and the mechanism behind the local spread is not known, it suggests the ΔshvR mutant can maintain low levels of chronic infection. Together, the results demonstrate that ShvR is not essential for intracellular persistence and replication, but that it regulates factors that are critical for the bacteria to disseminate efficiently and cause robust pro-inflammatory infection.

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Fig 3. A ΔshvR mutant is unable to spread and elicit strong pro-inflammatory responses.

B. cenocepacia K56-2, ΔshvR, and complemented ΔshvR were microinjected intravenously in embryos at 30 hpf. (A) Real time imaging of representative Tg(mpx:eGFP) injected larvae (neutrophils green) with fluorescent bacteria (red filter) at 24 and 48 hpi. For each bacterial strain, the same embryo was imaged at both time points. White asterisks in upper images indicate the region imaged with a higher magnification objective and presented underneath. Black arrows in bright field inset indicate tissue damage. Scale bar, 50 μm. (B) Mean neutrophil numbers in Tg(mpx:eGFP) larvae of non-infected control (grey open circles) and infected with K56-2 (black circles), ΔshvR (red circles) and complemented mutant (green circles). Representative images of each group, and an example of a binary conversion image used for pixel quantification are provided in S1 Fig. Quantification was performed prior to injection (0 hpi) and at 24 and 48 hpi. Only in non-infected and ΔshvR-infected embryos the 72 hpi time points were considered as embryos were still alive. Geometric means with each data point representing an individual embryo. Pooled results from two independent experiments are shown (n = 5 respectively per group per experiment). Significance was determined using one-way ANOVA with Sidak’s Multiple Comparisons test. (C, D) Mean relative il1b and cxcl8 gene expression levels (qRT-PCR) in embryos injected with on average 273 ± 114 CFU of B. cenocepacia K56-2 (black bars) or 184 ± 111 CFU of ΔshvR (red bars). Normalization was performed to the PBS-injected control group at each time point. Asterisks above each bar indicate significance compared to the PBS control at each time point, and significance between groups per time point is indicated with a horizontal line. Error bars represent mean with SEM of three independent experiments. Statistical analysis of qRT-PCR data was performed using one-way ANOVA with Tukey’s Multiple Comparison Test. (B, C, D) * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, ns: not significant.

https://doi.org/10.1371/journal.ppat.1007473.g003

The innate immune response towards the shvR mutant is significantly reduced compared to wildtype

We have shown earlier that acute infection caused by B. cenocepacia K56-2 correlates with systemic phagocyte death, whereas persistent infection, caused by for instance B. stabilis LMG14294, triggers neutrophilic inflammation [29]. Neutrophil numbers were quantified during infection with wildtype K56-2, ΔshvR and the complemented mutant at 0, 24, 43, and 72 hpi using Tg(mpx:eGFP) transgenic embryos, which express GFP specifically in neutrophils (S1 Fig and Fig 3B). Embryos infected with B. cenocepacia K56-2 showed a strong decline in the number of neutrophils, with only few cells remaining at the end stage of infection (Fig 3B and [29]). In contrast, infection with ΔshvR resulted in neutrophilic inflammation with a significant increase in the number of neutrophils compared to non-infected control embryos from 43 hpi (Fig 3B). The complemented mutant caused neutropenia, as seen for the wildtype strain, confirming restoration of virulence by expression of shvR in trans.

B. cenocepacia K56-2 induces a robust increase in host pro-inflammatory cytokine expression ([29]; Fig 3C and 3D). The expression of il1b and cxcl8/il8 upon infection with ΔshvR was, however, only moderately increased compared to PBS-injected embryos (Fig 3C and 3D), as shown earlier for B. stabilis [29]. Moreover, at 24 hpi, no significant difference in gene expression was observed between embryos injected with PBS or ΔshvR. These results confirm that ShvR is required for the induction of robust pro-inflammatory responses.

ShvR tightly regulates factors required for transition from intramacrophage stages to acute fatal infection

Based on the requirement for macrophages for intracellular bacterial replication of K56-2 and subsequent pro-inflammatory fatal infection [29], and the restriction of the ΔshvR mutant to macrophages, we hypothesised that ShvR could be an important switch between intracellular persistence and acute pro-inflammatory infection. To address this hypothesis, we used a rhamnose inducible expression system [31] to temporally control the induction of shvR expression in the ΔshvR mutant during infection in zebrafish embryos. This allowed us to study the effect of experimental induction of shvR gene expression during intramacrophage stages of B. cenocepacia on disease outcome. The shvR coding sequence was cloned under control of the rhamnose-inducible promoter PrhaB, resulting in PrhaB:shvR and introduced in ΔshvR. The PrhaB- control plasmid, lacking the shvR coding sequence, served as a negative control. Zebrafish embryos were injected at 30 hours post fertilisation (hpf) with ΔshvR(PrhaB-) and ΔshvR(PrhaB:shvR). At 24 h post injection of bacteria, at the time ΔshvR resided in macrophages (Fig 2), a 2% rhamnose solution, or PBS as a control, was injected into the circulation of the infected embryos (see Fig 4A for schematic representation). In order to maintain shvR expression, the embryos were further incubated in E3 medium with 2% rhamnose. The presence of PrhaB- in ΔshvR had no effect on embryo survival or bacterial burden, even after injection of rhamnose (Fig 4B and 4C). Strikingly, application of rhamnose, but not PBS, restored virulence to ΔshvR harbouring PrhaB:shvR, measured as host mortality and increase in bacterial burden (Fig 4B and 4C). These results confirm a critical role for ShvR in acute infection.

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Fig 4. ShvR tightly controls bacterial factors needed for fatal infection.

(A) Schematic representation of rhamnose-inducible shvR (PrhaB:shvR) and control (PrhaB-) constructs, cloned on a pBBR replication unit transferred into ΔshvR (right panel). Both strains were injected intravenously at 30 hpf. Zebrafish embryos were incubated for 24 hours in E3 water. At 24 hpi, PBS or 2% rhamnose, respectively, were injected intravenously to induce shvR gene expression (left panel). (B, C) Embryo survival and CFU counts after injection of 183 ± 136 CFU of B. cenocepacia K56-2 ΔshvR(PrhaB-) (red) and 22 ± 10 CFU of ΔshvR(PrhaB:shvR) (green), followed by injection of rhamnose or PBS, respectively at 24 hpi (black arrows). A representative experiment of three is shown (n = 20 (B) and n = 5 (C) respectively per group). (B) Solid lines indicate PBS-injected embryos while dashed lines indicate rhamnose -injected embryos. Differences in survival were determined using a Log rank (Mantel-Cox) test. (C) Bacterial burden was analysed at 0, 24, 48 and 72 hpi. The determination of CFU at 24 hpi was performed prior to rhamnose or PBS injection. Solid circles indicate PBS-injected embryos and open circles rhamnose injected embryos. Geometric means with each data point representing an individual embryo. Significance between groups was determined using one-way ANOVA with Sidak’s Multiple Comparisons test. * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, ns: not significant.

https://doi.org/10.1371/journal.ppat.1007473.g004

To visualise the effect of temporal induction of shvR expression on intracellular B. cenocepacia, fluorescence microscopy was performed using Tg(mpx:eGFP) transgenic embryos infected with ΔshvR(PrhaB:shvR) expressing mCherry. While injection with PBS in embryos infected prior with ΔshvR(PrhaB:shvR) did not alter the infection progression of the ΔshvR mutant, injection of rhamnose resulted in changes that are characteristic of acute infection (Fig 5A). These changes include neutrophil recruitment towards infection sites, indicative of increased pro-inflammatory signalling, and as described for acute infection with B. cenocepacia K56-2 [29]. The infection rapidly worsened and at 24 h post rhamnose injection the embryos were neutropenic (Fig 5A), as seen for infection with wildtype (Fig 3A). The 2-log increase in bacterial burden within 48 hours post rhamnose injection (Fig 4C) was in agreement with real time observations using fluorescence microscopy and time lapse confocal imaging (Fig 5A and S2 Movie). At later time points after rhamnose injection, tissue damage as a sign of strong pro-inflammatory responses could be observed (Fig 5A).

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Fig 5. ShvR-mediated transition from persistent to acute pro-inflammatory infection.

(A) Real time imaging of Tg(mpx:eGFP) embryos (green neutrophils) injected with ΔshvR(PrhaB:shvR) (in red), without (PBS) and with induction of shvR gene expression (rhamnose) at 24 hpi. Fluorescence imaging was performed at 0, 3, 6, 24 and 44 h post PBS or rhamnose injection. A single embryo was followed and visualised in time for each of the two conditions (representative embryos of a total of 20 embryos per experiment per condition are shown, 3 experiments). Green and red overlay images are shown. Inset shows the same image in bright field. White arrow heads indicate cells full of bacteria. Black arrow in bright field inset indicates tissue damage. Scale bars, 50 μm (B, C) Mean relative il1b and cxcl8 expression levels analysed by qRT-PCR at 24 h post injection with rhamnose in non-infected and infected embryos. Embryos were injected with an average of 102 ± 37 CFU of B. cenocepacia K56-2 ΔshvR(PrhaB-) and an average of 74 ± 45 CFU of ΔshvR(PrhaB:shvR). At 24 hpi embryos were injected with either PBS or rhamnose. Black bars indicate PBS-injected embryos and red bars represent rhamnose-injected embryos. Normalization was performed to a PBS-injected non-infected control group. Injection of rhamnose had no significant effect on il1b and cxcl8 expression (non-infected rhamnose-injected control). Significance between groups per time point is indicated with a horizontal line. Error bars represent mean with SEM of two independent experiments. Statistical analysis was performed using one-way ANOVA with Tukey’s Multiple Comparison Test. * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, ns: not significant.

https://doi.org/10.1371/journal.ppat.1007473.g005

Next, we analysed whether global cytokine gene expression levels were induced after restoration of shvR expression in ΔshvR(PrhaB:shvR) bacteria. While no difference in relative expression levels was observed between rhamnose and PBS conditions in embryos infected with ΔshvR(PrhaB-), a significant increase in the expression of cxcl8 and il1b was observed in embryos infected with ΔshvR(PrhaB:shvR) after injection with rhamnose (Fig 5B and 5C). Thus, ShvR strongly controls the expression of genes that are required for the transition from intracellular persistence to acute pro-inflammatory infection.

B. cenocepacia K56-2 afcE determines the difference between intracellular persistence and acute fatal infection

ShvR has been described as a global regulator of virulence factors in B. cenocepacia, moderately (2 to 4 fold) changing the expression of a large set of genes. The antifungal activity cluster afc, comprising 24 genes organised in two divergently expressed operons localised adjacent to shvR, has been described to show strongly decreased expression levels (100 fold) in the absence of shvR, demonstrating afc expression is under tight positive control of ShvR [25]. In addition, mutation of afcE mimics the phenotypes found for ΔshvR, including shiny colony morphology, altered membrane properties, and reduced virulence in a rat infection model [25,27,28], suggesting the afc cluster is a major target of ShvR. In analogy to the experiments described above for ΔshvR, we therefore analysed a role for AfcE in persistent/acute transition.

Infection experiments showed that ΔafcE(DSRed_pBBR) was severely attenuated in virulence in zebrafish embryos (Fig 6A and 6B). In agreement with the inability of the mutant to cause fatal infection, ΔafcE showed significantly lower bacterial burden compared to K56-2 at 24 hpi, similar to that observed for ΔshvR (Fig 1). Expression of afcE in trans (ΔafcE (Plac:afcE;DSRed_pBBR) complemented virulence to the ΔafcE mutant (Fig 6A and 6B). ΔafcE was taken up by macrophages as efficiently as the wildtype and ΔshvR mutant from the blood circulation (Fig 2B and S2A Fig). After phagocytosis by macrophages, ΔafcE bacteria were observed in macrophages also at later time points, sometimes, like ΔshvR, at high numbers (Fig 6C). Quantification of intracellular ΔafcE bacteria at 1 hpi and 24 hpi (S2B Fig) confirmed that the ΔafcE mutant is able to survive and replicate inside macrophages, as shown for ΔshvR (Fig 2E).

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Fig 6. A ΔafcE mutant mimics the persistent infection caused by a ΔshvR mutant.

Bacterial burden (A) and survival assays (B) of zebrafish embryos micro-injected with on average 120 ± 41 CFU of wildtype B. cenocepacia K56-2 (black), 126 ± 55 CFU of ΔafcE (red), and 146 ± 31 CFU of complemented ΔafcE (green) with Plac:afcE;DSRed_pBBR expressing afc from the lac promoter. Pooled results from two independent experiments are shown (n = 20 (A) and n = 5 (B) respectively per group per experiment). Geometric means with each data point representing an individual embryo. Significance between groups was determined using one-way ANOVA with Sidak’s Multiple Comparisons test (A). Statistical analysis of survival percentage was done a Log rank (Mantel-Cox) test (B). (C) Tg(mpeg1:mCherry-F) transgenic embryos, specifically expressing mCherry in macrophages, were injected with ΔafcE harbouring pIN301 (green). Representative image (red and green overlay) of an infected macrophage at 18 hpi. Scale bar, 12.5 μm. (D) Real time imaging of representative Tg(mpx:eGFP) injected larvae (neutrophils in green) with fluorescent bacteria (red) at 24 and 48 hpi. For each bacterial strain, the same embryo was imaged at both time points. White asterisks in upper images indicate the region imaged with a higher magnification objective and presented underneath. Black arrows in bright field inset indicate tissue damage. Scale bars, 50 μm. (E, F) Mean relative il1b and cxcl8 gene expression levels (qRT-PCR) in embryos injected with on average 273 ± 114 CFU of B. cenocepacia K56-2 (black bars) or 101 ± 41 CFU of ΔafcE (red bars). Data for afcE were obtained in the same experiments shown in Fig 3, therefore the values for expression of the cytokines in K56-2 infection are the same in the graphs presented in both figures. Normalization was performed to PBS-injected non-infected control group at each time point. Asterisks above each bar indicate significance compared to the PBS control at each time point, and significance between groups per time point is indicated with a horizontal line. Error bars represent mean with SEM of three independent experiments. Statistical analysis was performed using one-way ANOVA with Tukey’s Multiple Comparison Test. (A, B, E, F) * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, ns: not significant.

https://doi.org/10.1371/journal.ppat.1007473.g006

Using non-invasive real time imaging, we observed that the ΔafcE mutant bacteria did not cause neutropenia nor disseminated infection, while expression of the afcE gene restored these wildtype features (Fig 6D). Further, analysis of the global pro-inflammatory response of the host showed that the expression of the cytokine genes cxcl8 and il1b is induced to lower levels during infection with ΔafcE compared to wildtype K56-2 (Fig 6E and 6F), similar to that shown for ΔshvR (Fig 3C and 3D). These results clearly demonstrate that AfcE is required for acute, disseminated pro-inflammatory infection.

Experiments using rhamnose-mediated induction of afcE during intramacrophage stages, as shown for shvR (Fig 4), were performed. Whereas infection with ΔafcE(PrhaB-) after injection of rhamnose or PBS at 24 hpi remained persistent, embryos infected with ΔafcE(PrhaB:afcE) and injected with rhamnose succumbed to the infection (Fig 7A), which correlated with an increase in bacterial CFU at 72 hpi (Fig 7B). Not all embryos showed increased bacterial numbers, however, and further experiments are needed to determine the reasons for the apparently larger biological variation in this assay for the ΔafcE mutant compared to the ΔshvR mutant (Fig 4C). In addition, global cytokine expression of embryos infected with ΔafcE(PrhaB:afcE) was significantly induced at 24 h post injection of rhamnose, but not PBS (Fig 7C and 7D). Together, these results demonstrate that AfcE is essential for bacterial dissemination and the robust pro-inflammatory fatal response seen for B. cenocepacia K56-2, in agreement with afc being the direct target of ShvR required for acute infection caused by B. cenocepacia K56-2.

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Fig 7. AfcE expression is critical for progression from persistent to disseminated acute fatal infection.

Application of the experimental method depicted in Fig 4A using rhamnose inducible afcE (PrhaB:afcE) and control constructs (PrhaB-) introduced into ΔafcE. (A,B) Embryo survival and CFU counts upon injection of on average 76 ± 29 CFU of B. cenocepacia K56-2 ΔafcE(PrhaB-) (red) and 50 ± 37 CFU of ΔafcE(PrhaB:afcE) (green). A representative experiment of three is shown (n = 20 (A) and n = 5 (B) respectively per group). (A) Solid lines indicate PBS injected embryos and dashed lines rhamnose injected embryos. A Log rank (Mantel-Cox) test was used for statistical analysis. (B) Bacterial burden was analysed at 0, 24, 48 and 72 hpi. The determination of CFU at 24 hpi was performed before rhamnose or PBS injection. Closed circles indicate PBS-injected embryos and open circles rhamnose injected embryos. Semi-filled circle indicates dead embryo. Geometric means with each data point representing an individual embryo. Significance between groups was determined using one-way ANOVA with Sidak’s Multiple Comparisons test. (C, D) Mean relative cxcl8 and il1b expression levels (qRT-PCR) at 24 h post injection of rhamnose. Embryos were injected with an average of 141 ± 42 CFU of ΔafcE(PrhaB-) and average 104 ± 45 CFU of ΔafcE(PrhaB:afcE). At 24 hpi embryos were injected with either PBS or rhamnose. Black bars indicate PBS-injected embryos and red bars represent rhamnose-injected embryos. Normalization was performed to PBS-injected non-infected control group. Significance between groups per time point is indicated with a horizontal line. Error bars represent mean with SEM of two independent experiments. Statistical analysis was done using one-way ANOVA with Tukey’s Multiple Comparison Test. * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001, **** p ≤ 0.0001, ns: not significant.

https://doi.org/10.1371/journal.ppat.1007473.g007

The shvR-afc cluster is present in a subset of the Bcc species

To better understand a role for ShvR/Afc in virulence within the Bcc, we performed bio-informatics analysis. The Burkholderia cepacia complex has seen a rapid increase in the number of bacterial species it contains due to improved characterization and reclassification [32]. The current methods for discrimination between species are based on recA gene sequences, multilocus sequencing and whole genome studies (reviewed in [33]). At present, the complex includes 21 formally named species [1,2]. For the construction of a phylogenetic tree, representative genomes of each species were included (in some cases more than one strain) with either a complete genome and annotation status or whole genome sequence (WGS, see S2 Table). A phylogenetic inference tree of these species was constructed using the nucleotide sequences of the Multilocus Sequence Typing (MLST) housekeeping genes (atpD, gyrB, gltB, lepA, recA, phaC and trpD) [34]. Fig 8 shows a maximum-likelihood phylogeny generated from concatenation of these 7 genes. B. pseudomallei K96243, B. thailandensis E264 and Ralstonia pickettii 12J were used as out-groups.

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Fig 8. Phylogenetic inference of the Burkholderia cepacia complex species.

Maximum likelihood phylogeny of 21 Burkholderia cepacia complex species (summarised in S2 Table) based on the alignment of concatenated nucleotide sequences from 7 housekeeping genes (bootstrap 1000 replicates). Two non-Bcc Burkholderia species and a Ralstonia pickettii strain were used as outgroups. The distances for nucleotide data were inferred using the General Time Reversible (GTR) model with gamma distribution (5 rate categories and 49% invariable sites). Scale bar indicates 0.1 substitutions per site. The presence or absence of shvR and the afc gene cluster has been indicated with + or -. B arboris strain has not yet been fully sequenced and the presence of shvR or afcE could not be determined. The strains of each species are clustered except for two B. cepacia strains (GG4 and JBK9). B. stabilis LA20W and B. vietnamiensis strain AU4i. We propose that their classification may need to be reviewed given the following reasons. The strain B. cepacia GG4 clusters closer to B. ambifaria species as has already been reported [61]; the clustering of B. cepacia strain JBK9 together with B. stabilis is consistent with the 73% whole genome similarity shared by the B. cepacia JBK9 and B. stabilis strains [52]. The B. stabilis strain LA20W not only has higher genome similarity with B. pyrrocinia DSM 10685 (91.25%; while only 67.2% with the B. stabilis reference strain ATCC BAA-067 (or LMG14294) [52]), but also encodes the shvR/afc cluster sequence in its genome (S3 Fig), while B. stabilis ATCC BAA-67 does not. B. vietnamiensis AU4i is the only one that appears clustered with B. cepacia ATCC 25416, being consistent with high similarity between their genomes 83–86% (against 52–54% similarity between AU4i strain and other B. vietnamiensis strains) [52] and the presence the shvR/afc cluster in its genome (S3 Fig).

https://doi.org/10.1371/journal.ppat.1007473.g008

To better understand the evolution of the LTTR regulator ShvR within the complex, BLAST-analysis was performed to identify orthologs of the gene BCAS0225 from B. cenocepacia J2315. A gene with high similarity to BCAS0225 (identity >85%) was identified in a subclade containing strains from 10 out of the 20 species for which complete genome sequences are available (B. arboris has not been sequenced). In all cases, the gene was located on the megaplasmid pC3. Similarly, orthologs were identified for afcE (BCAS0208). Fig 8 shows that the presence/absence of shvR within the Bcc complex matches that of afcE. Further investigation of the adjacent afc cluster showed that the 24 afc genes and their organization, including the presence and location of the shvR gene are well-conserved among shvR-containing Bcc species (S3 Fig).

In addition to the 19 analysed Bcc strains (covering 10 species) that carry the shvR-afc cluster, B. pseudomallei and B. mallei have a subset of the afc genes, including afcE. The operon in B. pseudomallei, located on chromosome 1, shares similarity to 15 of the 22 genes in the Bcc afc cluster and is likely regulated by an LTTR regulator unrelated to ShvR, encoded by an ORF adjacent to the cluster (BPSL0494, S3 Fig). This regulator shares amino acid similarities with only the DNA binding domain of the ShvR protein (27% amino acid identity). BPSL0494 shares 77% nucleotide identity with BCAS0283, another LTTR identified in B. cenocepacia J2315 encoded on pC3, with homology mainly in the 5’ 200 bp and the 3’ 150 bp. A maximum-likelihood phylogeny of afcE (S4 Fig) confirmed that the acyl-CoA dehydrogenase from B. pseudomallei is the most distant in the Burkholderia group, although they share 80% of nucleotide similarity.

Analysis of the region upstream of the shvR coding sequence amongst the shvR-positive strains of the Bcc complex showed nucleotide variations (SNPs, indels), not only between species but also between strains belonging to the same species (S5 Fig). As expected, strains belonging to the same species contain fewer variations, compared to, for example, those observed between the B. cenocepacia and B. lata strains. Within the B. cenocepacia species, the recA lineage IIIA and IIIB (Fig 8) each appear to have their own consensus sequence for the shvR upstream region. Interestingly, a 2-nucleotide gap, at position -238 from the translation start site (S5 Fig), followed by a short highly non-homologous region compared to the conserved sequence found in the other cluster, is observed in the strains of B. cepacia, B. pyrrocinia, B. stabilis, B. lata, B. contaminans, B. metallica and B. vietnamiensis. This is in agreement with the phylogenetic relationship shown in Fig 8, and suggests that this deletion and the following non-homologous region occurred before speciation.

Overexpression of shvR in H111 causes an afc-dependent increase in virulence

B. cenocepacia, B. cepacia and B. contaminans, Bcc species which are generally correlated with more severe disease in humans, all carry genes encoding ShvR and Afc proteins. Several B. cenocepacia strains, including the epidemic CF isolates K56-2 and J2315, are highly virulent in different model systems, including zebrafish [29]. However, B. cenocepacia H111, which is very closely related to K56-2, was shown to be less virulent in G. mellonella and rats [35,36]. In zebrafish, we previously described that this strain can show substantial variation between experiments [37], with mortality rates between 20 and 60% at 4 days post fertilisation (dpf), but always less virulent than K56-2 (Fig 9). We also found that H111 does not show afc-dependent antifungal activity as seen for K56-2 [37]. Comparison of the shvR coding sequences showed 3 silent SNPs between H111 and our K56-2 strain. Of note, the sequenced K56-2Valvano strain has a 4^th SNP giving rise to a variant Tyr78 to Cys78. We set out to analyse whether differences in shvR expression could account for the differences in virulence observed between K56-2 and H111, and thus confirm the role of shvR/afc in virulence of B. cenocepacia.

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Fig 9. Differences in shvR expression between B. cenocepacia strains K56-2 and H111 cause afc-dependent differences in virulence.

Embryo survival after intravenous injection of an average of 192 ± 22 CFU of B. cenocepacia K56-2 wildtype (solid black lines), 273 ± 132 CFU of K56-2(Plac-shvR_pBBR) (dashed black lines), 185 ± 47 CFU of B. cenocepacia H111 wildtype (solid blue lines), 174 ± 10 CFU of H111(Plac-shvR_pBBR) (dashed blue lines), 198 ± 53 CFU of H111Δafc (solid red lines) and 228 ± 29 CFU of H111Δafc (Plac-shvR_pBBR) (dashed red lines). Pooled results from three independent experiments are shown (n = 24 per experiment per strain, K56-2 was included only in 2 experiments). Statistical analysis was performed using a Log rank (Mantel-Cox) test. **** p < 0.0001, ns: not significant.

https://doi.org/10.1371/journal.ppat.1007473.g009

To be able to study the role of shvR and afc in virulence of H111 in the absence of upstream regulatory signals, we cloned the shvR gene of H111 under control of the lac-promoter on pBBR1MCS and constructed an H111 mutant lacking the complete afc cluster. K56-2, H111 and H111Δafc were transformed with Plac-shvR_pBBR or the pBBR1MCS control plasmid. Expression of shvR from the lac promoter led to a change in colony morphotype of H111 from shiny to rough, in a manner that depended on afc (S6 Fig). This suggests that the endogenous shvR gene is not expressed in H111 on agar plates, but that expression of shvR in trans induces the expression of a functional AFC lipoprotein. H111 caused fatal disease in on average 40% of injected embryos over a 4-day time period, compared to 100% mortality in 2 to 3 days for K56-2 (Fig 9). Zebrafish embryos injected with the H111Δafc mutant survived and remained without any clinical signs of infection during the experimental time, in agreement with the experiments performed with K56-2ΔafcE (Fig 6B). The finding that afc is essential for acute fatal infection also in H111 suggests that shvR and afc expression is induced in wildtype H111 upon injection in zebrafish larvae, but probably does not reach the level of that in K56-2. Strikingly, overexpression of shvR from the lac promoter increased virulence in H111, but not the H111Δafc mutant, almost to that seen with B. cenocepacia K56-2 (Fig 9). These data validate the results obtained with K56-2 that shvR/afc are critically important for acute virulence in zebrafish larvae. They also unambiguously demonstrate that shvR/afc are functional in H111, and suggest that differences in inducing signals or upstream elements render H111 less virulent in zebrafish compared to K56-2.

Ethics statement

Zebrafish (Danio rerio) were kept and handled in compliance with the guidelines of the European Union for handling laboratory animals (http://ec.europa.eu/environment/chemicals/lab_animals/home_en.htm). Zebrafish studies performed at VBMI are approved by the Direction Départementale de la Protection des Populations (DDPP) du Gard (ID 30-189-4) and the Comité d'Ethique pour l'Expérimentation Animale Languedoc-Roussillon (CEEA-LR-12186). Infection experiments in this study were terminated before the larvae reached the free feeding stage and did not classify as animal experiments according to the 2010/63/EU Directive.

Zebrafish

The zebrafish line AB was used as wildtype (WT), and the transgenic reporter lines Tg(mpx:eGFP)ⁱ¹¹⁴ [47] and Tg(mpeg1:mCherry-F)^ump2Tg [48] were used to analyse host phagocyte behaviour. Care and maintenance of zebrafish was as described previously [29]. Eggs were obtained by natural spawning and incubated at 29°C in Petri dishes containing E3 medium (5mM NaCl, 0.17mM KCl, 0.33mM CaCl₂, 0.33 mM MgSO₄) as described [49]. Methylene blue was omitted.

Bacterial strains, plasmids and growth conditions

The bacterial strains, plasmids and primers used in this study are listed in S1 Table. Strains were grown at 37°C in Lysogeny Broth (LB), supplemented with ampicillin at 100 μg/mL for Escherichia coli, chloramphenicol at 30 μg/mL and 100 μg/mL for E. coli and Burkholderia cenocepacia respectively, tetracycline at 250 μg/mL (B. cenocepacia) and trimethoprim at 50 μg/mL (B. cenocepacia). No differences in growth were observed between B. cenocepacia K56-2 and the ΔshvR and ΔafcE mutants when grown in LB medium.

Generation of plasmid constructs

For the construction of pIN233, the mCherry coding region from plasmid pSAT1:mCherry-MCS-nVenus (pE3370) was amplified using primers mCherry-3 and mCherry-4. The PCR fragment was digested with XbaI and NdeI and cloned into pIN29 [30], replacing the DSRed coding sequence. Plasmid pIN298 resulted from cloning the mCherry coding sequence from pIN233 into pCR11 (a gift from M. Kovach), a Cm^R derivative of the single copy plasmid pMR10 (GenBank: AJ606312.1), using HindIII and XbaI restriction sites.

A ~1.6 kb fragment containing full length shvR and upstream region PshvR (primers pshvRXhoI for and shvRXbaI rev), a 995 bp fragment containing the shvR coding region (primers shvRNdeI for and shvRXbaI rev) and a ~1.8 kb fragment containing the afcE coding region (primers 0208_NdeI for and 0208_XbaI rev) were separately amplified from K56-2 genomic DNA by PCR using Pfu polymerase (Life technologies, USA). The fragments were first subcloned in pUC29 and verified by sequencing (MWG Operon Eurofins, Germany).

To construct pIN308, the PCR product of PshvR:shvR was cloned into pIN29 as a XhoI/HindIII fragment, and named pIN307. From this plasmid, the PshvR:shvR fragment was cloned into pIN298 using the SpeI and BamHI restriction enzymes (XhoI and XbaI).

The rhamnose inducible system from pSCPrhaB2 [31], containing the rhamnose-regulated PrhaB promoter and the genes rhaR and rhaS of the rhamnose operon, was cloned into pIN177 using XbaI and PstI restriction sites, and named pIN299 (or PrhaB-, serving as negative control in rhamnose induction experiments, for simplicity). pIN177 is a derivative of pBBR1MCS plasmid lacking the oriT/mob region and containing a strong trp termination signal (pIN32; [30]), a tac promoter sequence (PstI/NdeI fragment from pIN17; [30]) and an additional NdeI/XbaI linker (catatgaagctttcgcgagctcgagatctaga).

To create PrhaB:shvR (pIN310), allowing rhamnose-inducible expression of shvR, the shvR coding sequence was cloned downstream the PrhaB promoter sequence as follows: the 995 bp shvR PCR fragment (primers shvRNdeI for and shvRXbaI rev) was cloned into XbaI/NdeI-digested pIN299, and named pIN309. A Ptac:mCherry fragment from pIN233, cloned in pUC29 as a SpeI/NotI fragment, was then inserted in the KpnI site of pIN309, resulting in pIN310. Plasmid pIN311 (PrhaB:afcE) was constructed by digesting pIN299 with NdeI/XbaI restriction sites and inserting the afcE fragment amplified using the primers 0208_NdeI for and 0208_XbaI rev.

The afcE complementation plasmid pINR139 was generated by cloning a Ptac:DSRed fragment from pIN29 as a SpeI/SacI fragment into pBBRS0208.

Plasmids were transferred into B. cenocepacia K56-2, ΔshvR and ΔafcE mutants by electroporation as described earlier [30].

The plasmid Plac-shvR_pBBR carries the coding sequence of shvR amplified from B. cenocepacia H111, using primers shvRHindFor and shvRBamRev. The restriction sites BamHI and HindIII were used to clone the fragment in pBBR1MCS.

Deletion of the afc cluster from B. cenocepacia H111

Deletion of the afc cluster was carried out using vectors pSHAFT2-FRT and pEX18Tp-FRT, as described in [37]. A fragment adjacent to the region to be deleted (UP) was amplified from B. cenocepacia H111 with primers upXhoF and upBglIIR (~1.3 kb), and digested with XhoI and BglII. The fragment was then inserted in pSHAFT2-FRT using the same sites, resulting in pSHAFT2-FRT-afcUP. This plasmid was inserted into the genome of B. cenocepacia H111 by single crossover recombination. Correct integration was confirmed with primer pairs AFCdelUPcheckF (GATCATCTTCTTCTCGCTCG) and pSHAFTseqR4 (GAACACTTAACGGCTGACAT, ~1.5 kb) and pSHAFT2For3 (GATTATTTTGCCCCGGTTTT) and AFCdelUPcheckR (GGAGATTTCGCATGATGTTT, 2 kb).

A fragment bordering the afc cluster on the other side (DOWN) was amplified from B. cenocepacia H111 using primers downpstF and downBamR. The resultant ~1.3 kb fragment was digested with PstI and BamHI and inserted between these sites in pEX18Tp-FRT. This vector (pEX18Tp-FRT-afcDOWN) was inserted into the genome of B. cenocepacia H111 bearing the previously integrated pSHAFT2-FRT derivative by single crossover recombination. Correct integration was confirmed with primer pairs AFCdelDOWNcheckF (GAATTGAACCGCTATCGCC) and M13R (~1.4 kb) and M13F and AFCdelDOWNcheckR (GCACAGGTTGCAGGTATT, 1.7 kb).

Plasmid pBBR5::FLP [50] was then introduced by conjugation, to stimulate recombination between the FRT sites integrated into the genome.

Flippase-mediated recombination was carried out at 30°C, and colonies were selected using the marker present on the pBBR5::FLP derivative. Colonies were then checked for loss of the markers present on the pSHAFT-2FRT and pEX18Tp-FRT derivatives (chloramphenicol and trimethoprim, respectively). Finally, deletion was confirmed by amplifying across the ends of the deleted region with primers AFCDELcheckFor and AFCDELcheckRev. Successful deletants gave rise to a band of ~700 bp.

Microinjection conditions and rhamnose-mediated induction of gene expression

Microinjection of zebrafish embryos was performed as previously described [49]. Briefly, B. cenocepacia strains were grown overnight in LB broth with appropriate antibiotics at 37°C. Bacterial dilutions to obtain the desired inoculum concentration were prepared in PBS (with 0.05% phenol red to visualize microinjection). Embryos were dechorionated 2 hours prior to microinjection and at 30 hours post fertilization (hpf) they were injected in the blood island. For microinjection, embryos were placed on agarose plates containing E3 medium with 0.02% buffered MS222 (tricaine; ethyl-3-aminobenzoate methanesulfonate salt). Injection was performed with a Femtojet microinjector (Eppendorf) and a micromanipulator with pulled microcapillary pipettes, under a stereo light microscope (Leica MS5). The pool of infected embryos was then randomized over CFU plating and survival assays. Embryos were maintained individually in 48-well plates in E3 medium at 29°C. In order to specifically induce the expression of shvR and afcE in ΔshvR and ΔafcE mutants, respectively, at 24 h after injection of the bacterial inoculum, rhamnose (2% in PBS) was injected directly in the blood circulation. As a negative control, half of the embryos of each group were injected with PBS. The embryos were then kept in E3+2% rhamnose or in E3 medium, respectively. Immediately after bacterial injection (T = 0 hpi), 5 embryos per strain were disrupted and plated on LB agar with the appropriate antibiotics to determine the precise inoculum size. At 24, 48 and 72 hpi (in the rhamnose assays), 10 μl of each serial dilution was deposited on a squared LB-agar plate for determination of CFU. Embryo survival was determined at regular time points, starting at 42 or 44 hpi and every two hours during periods with high mortality rates. Time of death was based on the absence of a heartbeat.

RNA extraction, cDNA synthesis and quantitative RT-PCR

RNA extraction, cDNA synthesis and qPCR analysis of zebrafish genes were performed as described [49]. The peptidylprolyl isomerase A-like (ppial) gene was used as a reference gene. The ΔΔCt method was used for analysis of the data, represented as column bar graphs normalized to a PBS-injected control group at each time point. Three biological replicates were performed (unless mentioned otherwise), each with two technical replicates. So far we have been unable to optimise qPCR of bacterial genes from infected zebrafish larvae, due to the high ratio of host to bacterial RNA.

Microscopy, image processing and quantification of fluorescent host immune cells

Embryos were imaged using a Leica DM IRB inverted microscope coupled with a Coolsnap fx black and white camera (Roper Scientific) as described [49], or a Nikon AZ100, coupled with Coolsnap HQ2 (Roper Scientifique). MetaVue software was used for imaging. Adobe Photoshop was used to colour black and white images, prepare overlay images taken with different channels, include scale bars, and crop images for the purpose of showing enlargements/insets. For analysis of phagocytosis and quantification of intracellular ΔshvR bacteria (sample image Fig 2A), the embryos were fixed in 4% PFA for 2 hours at RT, or overnight in 3% PFA at 4°C and analysed using an Olympus confocal laser scanning microscope Fv10i and Fluoview software.

Quantification of neutrophil cell numbers was performed according to [51]. In brief, images taken with identical camera settings (S1A Fig) were converted to binary in ImageJ 1.47v, resulting in images in which fluorescence was converted into black pixels onto a white background (S1 Fig). Five randomly selected individual cells per image (embryo) were taken to determine pixel size per phagocyte, as described [51]. Total pixel counts were then divided by the average of the 5 individual cells to determine the total number of fluorescent cells.

Phylogenetic analyses of Burkholderia species

A maximum likelihood phylogeny was created from 43 Burkholderia cepacia complex strains and 3 outgroup species (B. pseudomallei K96243, B. thalandensis E264 and Ralstonia pickettii 12J, in S2 Table). The tree is based on the concatenated alignment of fragments of nucleotide sequences of the Multilocus Sequence Typing (MLST) gene set, atpD, gyrB, gltB, lepA, phaC, recA and trpB [34]. For strains with MLST ID indicated in S2 Table, the sequences were retrieved from PubMLST database (pubmlst.org/bcc), for the others the coding sequence of the genes were obtained from BLAST searches in NCBI [52], PATRIC (patricbrc.org, [53]) or the Burkholderia Genome Database (burkholderia.com, [54]). For those cases, the sequences were shortened upon the alignment. The sequences of each gene were first aligned in MEGA6.06 [55] using ClustalW [56,57] and then all gene sequences were concatenated using the program SequenceMatrix [58]. The concatenated genes sequences were then introduced in MEGA6.06 to construct a maximum likelihood tree. The General Time Reversible (GTR) model [59] was used to determine nucleotide distances, with gamma distribution (5 rate categories and 49% invariable sites), assuming partial deletions in the missing data and a cut-off of 95%. The maximum likelihood heuristic method used was the Nearest-Neighbour-Interchange (NNI). A bootstrap of 1000 replicates was used.

Phylogenetic analysis of shvR and afcE

A BLAST analysis of the promoter region of shvR (500bp) from B. cenocepacia J2315 (BCAS0225) was performed using PATRIC against a genome database with the Burkholderia strains summarised in S2 Table (except for B. arboris LMG 14939). The sequences of each of the identified genes were obtained (either from PATRIC, NCBI or Burkholderia Genome Database) and aligned in MEGA6.06 [55] using ClustalW [56,57]. A maximum likelihood phylogenetic tree for the afcE gene was created in MEGA6.06 using the same parameters as described above for the Burkholderia species tree, except that the model used was Tamura 3-parameters model (T92), with gamma distribution (5 rate categories), and were considered 40% invariable sites.

Analysis of the Afc cluster

To analyse the similarities of the afc clusters of Burkholderia species, the pC3 genomic sequences (or contigs) of the 43 sequenced Bcc strains (except for B. arboris LMG 14939) and B. pseudomallei K96243, summarized in S2 Table, were retrieved from NCBI. Sequence alignments were performed with MAUVE using default parameters (default seed weight set to 15; set for determination of Local Collinear Blocks (LCB); full alignment (default minimum LCB weight); set for iterative refinement; set for sum-of-pairs LCB scoring) [60].

Statistical analyses

For statistical analysis GraphPad Prism 6.0 software was used. The average inoculum is calculated as the average CFU of 5 embryos T = 0 values ± SD, and indicated in the legend to the respective graphs. CFU counts from individual larvae at later time points were log₁₀ transformed and the significance between the multiple selected groups was determined using one-way ANOVA with Sidak’s Multiple Comparisons test. To include larvae in which no CFU were detected at 24 or 48 hpi, the 0 count was converted to 1, prior to log-transformation. Since the geometric mean cannot be calculated for groups that contain log₁₀ values of 0, we have set this value at 0.0001 to be able to calculate the geometric mean. No difference in significance was found when using the 0 or 0.0001 log value.

In survival assays statistical analysis was done using a Log rank (Mantel-Cox) test. For statistical analysis of macrophage and neutrophil cell counts and intracellular bacterial numbers, one-way ANOVA with Sidak’s Multiple Comparisons test, unpaired t-test, and Mann-Whitney were used as indicated in the legends to each graph.

The data from qRT-PCR of cytokine genes were log₂-transformed, and significance of the data was analysed using one-way ANOVA with Tukey’s Multiple Comparison Test. Columns indicate mean fold with SEM. For each treatment, normalized to the corresponding PBS control, significance in relative fold-change is indicated with an asterisk above the column, significance between treatments is indicated with a connective line between the bars. In rhamnose assays, rhamnose-induction and PBS treatments in both conditions (control plasmid and gene expressing plasmid) were normalized to the non-infected PBS control. Significance is indicated as: ns, non-significant, *, p ≤ 0.05; **, p ≤ 0.01; ***, p ≤ 0.001; ****, p ≤ 0.0001.