September 2014

Analysis of a role of the LysR-Type Regulator ShvR in virulence of Burkholderia cenocepacia using zebrafish as a model Margarida Castro Gomes1 1

Master Student in Biological Engineering at Instituto Superior Técnico, Universidade de Lisboa, Av. Rovisco Pais, 1 - 1049-001 Lisboa, Portugal

Burkholderia cenocepacia is an opportunistic pathogen, comprised in the Burkholderia cepacia complex (Bcc), that causes chronic infections in immunocompromised people, mainly in cystic fibrosis patients. These bacteria have been demonstrated to be capable of living and multiplying inside cells by evading host cell degradation mechanism. Zebrafish (Danio rerio) embryos have been developed as a model to study Bcc infections. In this study the zebrafish model was used to further understand a role for the LysR-type transcriptional regulator (LTTR) shvR from B. cenocepacia K56-2 in virulence. ShvR has been shown to be important for virulence in alfalfa and rat models of infection. Here, it is shown that the shvR mutant causes a persistent infection pheno-type in zebrafish embryos, in contrast to the acute infection caused by its parent K56-2. A persistent phenotype, with bacteria residing in macrophages, was further confirmed by analysis of host phagocyte response. Further-more, new tools were developed to start to better understand the role of ShvR in virulence and persistence.

Keywords: Burkholderia cenocepacia, shvR, zebrafish, immune response, intracellular survival, persistence

1 INTRODUCTION This project focuses on virulence mecha-

nisms of Burkholderia cenocepacia, that be-longs to the Burkholderia cepacia complex (Bcc). This complex currently includes 17 phe-notypically similar species that are found ubiqui-tously in the natural environment and in indus-trialized environments, but importantly, they are life-threatening pathogens of immunocompro-mised persons, especially those with cystic fibrosis (CF) (1–7). Bcc bacteria are known to resist under different (environmental) condi-tions, including nutrient limitation, toxic com-pounds (8), antimicrobial peptides (4,9) and almost all clinically used antibiotics, thus com-plicating treatment in infected individuals (10,11). The metabolic diversity and flexibility of these species can be explained by their ge-nome variety and genome plasticity, allowing the bacteria to rapidly adapt to different condi-tions (12). Their genomes are among the larg-est bacterial genomes known, with sizes rang-ing from 7 to 9 Mb, consisting of three repli-cons: chromosome 1, chromosome 2 and chromosome 3 (13). The latter has recently been described as a mega virulence plasmid that confers a competitive advantage to the organism, since it is essential for pathogenicity in various hosts, including rats and zebrafish,

and antifungal activity and is involved in stress tolerance (14,15). Cystic fibrosis is a lethal recessive human ge-netic disorder caused by mutations in the gene encoding the cystic fibrosis transmembrane conductance regulator (CFTR) (reviewed in (16)). The main function of this regulator is as a chloride channel, however many other functions are associated to this protein. The infection of the lungs of CF patients normally occurs during infancy and early childhood with organisms such as Staphylococcus aureus and Haemophi-lus influenza (17). Recent years have seen the emergence of several new pathogens of clinical relevance to CF, which is the case for Burkholderia cenocepacia (4,8). The outcome of infections with these bacteria is unpredicta-ble; it can vary from periods of chronic infection to sudden acute, systemic infection (18), and leaves the physicians with few options for treatment due to the high intrinsic antibiotic resistance. Moreover, infections with B. ceno-cepacia are generally associated with reduced survival and a higher risk to develop “cepacia syndrome” (19,20). Until now the factors that are involved in the sudden fatal changes of the infection are still unknown. In higher organisms, the first line of defense against viruses, bacteria and other disease cau-

Table 1 - Bacterial strains and plasmids used in this study.

sing organisms is provided by macrophages and neutrophils, as part of the innate immune response. In most cases, these cells are capa-ble of eliminating the invading organism through phagolysosomal degradation. B. cenocepacia has been shown to be phagocytized and killed in a ROS-dependent manner in neutrophils (26), however in cell culture macrophages, and macrophages in zebrafish embryos, the bacte-ria have been shown to escape from the classi-cal endocytic pathway and survive and replicate (reviewed in (27)).

Indeed, analysis of lung tissue from patients has demonstrated that B. cenocepacia are mainly present inside macrophages (28,29). Schwab et al. showed that B. cenocepacia and P. aeruginosa create different niches (29). Us-ing immunohistochemistry, they were able to show that Bcc bacteria are found in bronchi and, very frequently, they are inside macro-phages within mucus, without any evidences of biofilm formation (29).

In this study the main focus is on a global regulator of gene expression, the shvR gene (BCAS0225, Genbank: AM747722.1), in B. cenocepacia K56-2 described by Bernier and colleagues that shows that it is attenuated in

various infection models (30) Subramoni, Vergunst et al. unpublished).

The aim of my project is to better under-stand the role of ShvR in virulence in vivo in more detail, using a zebrafish embryo model of infection. The use of this animal as a model for infection studies has great advantages in-cludeng the transparency of the embryos, al-lowing for real-time observation of infection, and the high similarity of its immune system to that of humans. Zebrafish has proven to be a good model to study different microbial infections, allowing for the study of the role of phagocytes, intracellular bacterial stages, and immune re-sponse in large detail. Infection models have been developed for instance for Mycobacterium (31), Staphyloccus (32), Pseudomonas (33) and Bcc species (25).

Moreover new tools were developed to aid in better understanding the role of ShvR in vivo. Deletion of shvR greatly attenuated the acute inflammatory infection caused by B. cenocepa-cia K56-2 in zebrafish embryos; instead, the shvR mutant persisted in macrophages. The results showed further characteristics of persis-tent infection.

2 MATERIALS AND METHODS 1. Bacterial strains and plasmids

The bacterial strains and plasmids used in this study are described in Table 1. Escherichia coli and Bcc strains were cultured at 37°C in Luria-Bertani (LB) broth, with or without 1.6% agar. Both E. coli and Bcc strains carrying plasmids encoding chloramphenicol resistance (CmR) were grown in the presence of 30 and 100 µg/mL, respectively, of the antibiotic. For selection of Bcc with trimethoprim re-sistance (TpR) a concentration of 50 µg/mL trimethoprim was used and for E. coli 100 µg/mL. Selection of E. coli colonies using pUC29 plasmid with ampicillin resistance was used at a concentration of 150 µg/mL of ampicillin.

2. DNA manipulations Polymerase chain reaction (PCR) conditions The

conditions used for a 25 µL reaction volume: 100 ng of B. cenocepacia K56-2 genomic DNA as a template, 0.5 µM of each primer (Table 2), 200 µM of each deoxynucleotide (Invitrogen) and 0.06 U/µL of Pfu DNA Polymerase (Prome ga). The samples were subjected to an initial denaturation at 95°C for 3 minutes, followed by 30 cycles of de-naturation (95°C for 45 seconds), annealing (57°C for 30 seconds) and elongation (72°C for 2 min/kb of expected product). After a final elongation at 72°C for 5 minutes, the samples were stored at 4°C until use. Amplified fragments were cloned as described in the Results, and verified by

Bacterial strains Reference B. cenocepacia K56-2 (LMG18863) (21) B. cenocepacia K56-2 ΔshvR (22) B. vietnamiensis FC441 (LMG18836) (23) B. stabilis (LMG14294) (24) E. coli DH5α

Plasmids Description Reference pIN29 DsRed reporter plasmid (25)

pIN233 mCherry reporter plasmid Subramoni,

Vergunst et al., unpublished

pCR11 Broad host range RK2-based low copy number

cloning vector

Gift from M. Kovach

pAV100 A. Vergunst

pUCP28T-shvR

shvR complementation plasmid

(22)

pAV209 This study pMG1 This study pMG2 This study pMG3 This study pMG4 This study pMG5 This study pMG6 This study

Table 2 - Amplification products and primers used in this study. Amplification

product Primer/Oligo Sequence (5’–3’) Annealing temperature

shvR promoter (596 nt)

pshvR for GAACATATGTCTCACATTAGCCATACCGCCGC 57 ºC pshvR rev TGCAAGCTTCGAATTCCGCCCGACATGCG shvR promoter and gene (1593 nt)

pshvRXhoI for AATTCTCGAGGAATTCCGCCCGACATGCGC 57 ºC

shvRXbaI rev GTTCTAGACTATCCGACGCGATACATCGGC

unst1 GTACAAGCGCCGCAACGACGAATACCGCCTGGTCCGCTAGGAGCT

unst2 CCTAGCGGACCAGGCGGTATTCGTCGTTGCGGCGCTT

sequencing (MWG Operon).

3. Zebrafish infections Zebrafish (Danio rerio) were kept and handled accord-

ing to national regulations for animal welfare (ID 30-189-4 and CEEA-LR-12186), and as described in (25,34). The embryos are raised in E3 egg water (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, 0.00005% meth-ylene blue), at 29.5°C. Experiments were performed using zebrafish embryos and terminated before the larvae reached the independent feeding stage. For infection exper-iments, AB and the Golden variety (35) were used. The transgenic reporter lines Tg(mpeg1::mCherry-F) and Tg(mpx::GFP) expressing mCherry specifically in macro-phages and GFP specifically in neutrophils, were used for analysis of host phagocyte behavior (36,37). Microinjection was performed as described in detail by by Mesureur and Vergunst (34). Briefly, bacterial dilutions of 50 bacteria/nL were prepared in PBS (with 0.05% phenol red to visualize microinjection). The embryos were dechorionated and placed on agar plates with E3 medium containing 0.02% buffered MS222 (tricaine; ethyl-3-aminobenzoate me-thanesulfonate salt; Sigma). About 1 nL of a bacterial suspension was microinjected directly into the blood circula-tion, either in the blood island or in the axial vein, with the use of a Fentojet microinjector and pulled microcappilaries. To determine inoculum size, 5 embryos were individually plated onto LB agar plates immediately after microinjection (T=0). To follow bacterial kinetics, inoculated embryos were incubated individually in E3 medium in 24- or 48-well plates at 29.5°C and sampled at 24 and 48 h post-infection (hpi), five embryos at each time point. To determine colony form-ing units (CFU) at T=0,, embryos were disrupted in 50 µL of tissue-culture-grade trypsin-EDTA (2%) and incubated for 20 min . For other time points, 50 µL of 2% Triton X-100 (in H2O) was added, and tubes were incubated for 30 min at room temperature prior to plating. For survival assays, embryos were maintained individually in 24-well plates in E3 medium at 29°C. At regular time points after infection, the number of dead embryos was determined visually based on the absence of a heartbeat.

4. Statistical analysis

Statistical analyses were performed using Prism 6.0c (GraphPad) and are detailed in each figure legend. Embryo survival data were presented in Kaplan-Meier survival plot, and a log-rank (Mantel-Cox) test was used for statistical analysis. Kinetics data were represented in scatter plots, and t-tests were performed to determine the p-value be-tween different conditions.

5. Microscopic analysis

For microscopy, a Leica DM IRB inverted microscope equipped for bright-field, differential interference contrast

(DIC), and fluorescence imaging was used. GFP, DsRed/mCherry and m2-Turquoise were excited using a 100 W mercury lamp, and fluorescence was detected using filter sets L5 (band pass [BP] 480/40; beam splitter [BS] 505; emission BP 527/30), N2.1 (515 to 560; BS 580; emis-sion long pass [LP] 590), A (340 to 380; BS 400; emission LP 425) and CFP (BP 436/20; BS 455; emission BP 480/40), respectively. For imaging we used a Coolsnap fx (Roper Scientifique). Embryos were transferred to E3 con-taining MS222 in glass-bottom dishes (MatTek Corp., Ash-land, MA) for direct visualization using 40x and 63x oil objectives. A Nikon AZ100 equiped for bright-field and appropriated filter for red and green fluorescence imaging, and coupled with Coolsnap HQ2 (Roper Scientifique), were used for imaging embryos for further fluorescence quantifi-cation. MetaVue software was used for imaging, and imag-es were processed further using Adobe Photoshop, or Image J.

3 RESULTS 1. Survival assays and bacterial kinetics

To analyze virulence of the shvR mutant, survival and kinetics assays were performed. Thirty-five embryos (30 hours post fertilization (hpf)) were microinjected intravenously (iv) with fluorescently labeled B. cenocepacia K56-2 (pIN29) and a K56-2 shvR (pIN29). Figure 1 summarizes the results of survival and kinetics assays from two experiments.

Analysis of bacterial multiplication in the in-fected embryos showed that K56-2 bacteria replicated rapidly in individual embryos, reach-ing high numbers at 48 hpi, in agreement with the acute infection phenotype for this strain, as shown in (25). In contrast, the number of K56-2ΔshvR bacteria remained relatively constant throughout the analyzed time period of 48 h, indicating a persistent infection (Figure 1A).

The embryo survival assay demonstrated that embryos infected with K56-2 wild-type rap-idly died after infection (Figure 1B), whereas all embryos infected with the shvR mutant were still alive at 72 hpi. In fact, the embryos infected with the mutant were still alive at 5 days post infection, at which time point the experiment

Figure 1 - B. cenocepacia K56-2ΔshvR is less virulent

than K56-2 in zebrafish embryos. Thirty-five embryos per experiment were microinjected with on average 31 and 19 CFU. For each strain, five embryos per time point were used to determine CFU, and twenty embryos were kept to determine survival times. The graphs show the results of two independent experiments. (A) A Student’s unpaired t-test showed that the mutant differed significantly from the WT at 24 hpi (P=0.0012) and at 48 hpi (P<0.0001). For K56-2 the two points represented by black triangles were two embryos that were dead at 48 hpi. (B) A log-rank (Man-tel-Cox) test showed that the mutant differed significantly from the WT (P<0.0001).

was terminated. These results show that the shvR mutant is strongly attenuated compared to its wild-type parent K56-2, and induced a per-sistent infection in zebrafish embryos. Thus, ShvR regulates factors needed for the devel-opment of acute fatal infection.

2. Analysis of the behavior of host immune cells in infected embryos

In embryos injected iv with K56-2, the bacteria have been shown to be phagocytized mainly by macrophages, in which they start to replicate about 6 to 7 hours later. The bacteria dissemi-nate from such infected macrophages around 10-12 hpi, first locally, again replicating inside cells, and resulting in local infection sites, sometimes forming visible cell aggregates (25). At such sites neutrophils are visibly recruited to infection sites (Mesureur et al., manuscript in preparation, Figure 2). Then at about 16-24 hpi, the bacteria re-enter the blood circulation result-ing in a systemic infection. At 24 hpi, most em-bryos show signs of severe tissue deterioration and necrosis – some embryos lose parts of the body (cell aggregates) – and the blood flow rate is severely reduced or has even stopped at this time point (Figure 2). Most neutrophils have been recruited to infection sites, and their numbers

have started to decline, with often hardly any macrophages and neutrophils detectable in the embryo by 24 hpi compared to non-infected embryos, which have a steadily increasing neu-trophil number during their development (Mesu-reur et al., manuscript in preparation). The em-bryos succumb to K56-2 infection, depending on the infection dose, from 40-48 hpi. In con-trast to the acute fatal infection seen with K56-2, shvR-infected embryos, at 24 hpi, show no signs of external deterioration but bacteria can be seen in macrophages. After phagocytizing shvR rapidly after microinjection, macrophages full of bacteria have been seen already after 24 hpi (Figure 3).

Often the whole cell was filled with bacte-ria, possibly in one large vacuole, but this has to be further determined. ShvR-infected macro-phages seemed larger than the ones infected with K56-2, although this has to be further quantified. The results suggest that the bacteria replicate inside macrophages and are able to establish a niche inside the macrophages. The shvR mutant bacteria were not able to spread from the infected macrophages as wild- type bacteria, resulting in a (intracellular) per-sistent phenotype. Further microscopic obser-vations indicated that although neutrophils are present in high numbers, they are not recruited massively to the site of infected macrophages, as in K56-2 infected embryos.

At 3 and 5 days post-infection macrophages full of bacteria were still observed. Occasional-ly, macrophage and neutrophil recruitment was seen around infected cells (data not shown).

Macrophage numbers are not affected in shvR infected embryos Tg(mpx::GFP x mpeg1::mCherry) embryos were injected with 10 to 30 CFU of B. cenocepacia K56-2 and shvR mutant (Figure 4B). In control embryos the number of macrophages increased throughout the 72h of the experiment, as expected in the developing embryos (Figure 4). In contrast, em-bryos infected with K56-2 showed significantly lower numbers of macrophages, which reduced in numbers during the infection, consistent with earlier real time observations (Mesureur et al., manuscript in preparation). On the other hand,

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Figure 2 - B. cenocepacia K56-2 infection in zebrafish embryos (Tg( mpx::GFP x mpeg1::mCherry) and Tg(mpx::GFP)). Fluo-rescence images of different time points after infection. The smaller insets show the corresponding bright light image. After injection (2 hpi), bacteria (blue) co-localize mainly with macrophages (red) but not with neutrophils (green). At 24 hpi massive neutrophil (green) recruitment to infection sites (bacteria in red) can be observed. This recruitment causes tissue damage, being visible in the light image (white arrow). At 42 hpi the infection has spread and almost no neutrophils are found, and as seen in the inset, the tissue damage is extensive. Photos at 2 and 24 hpi courtesy of A. Vergunst and 42 hpi of J. Mesureur.

Figure 3 - B. cenocepacia K56-2ΔshvR infection in zebrafish embryos (Tg(mpx::GFP)). Fluorescence images of different time points after infection. The smaller insets show the corresponding bright light image. Throughout the infection time, macrophag-es become fuller with bacteria (red, indicated by white arrows) and generally no neutrophil recruitment is observed, although their numbers increase. (B) Macrophage infected with B. cenocepacia K56-2ΔshvR in zebrafish embryos (Tg(mpx::GFP)). At 48 hpi the macrophages are full with bacteria and occasionally neutrophils are seen around the infected cells. Scale bar 25 µm.

counts in embryos infected with the shvR mu-tant did not show a reduction in macrophage numbers as K56-2, but showed similar numbers of macrophages as the control embryos (Figure

4A). In parallel with the phagocyte cell counting,

embryos were plated at the different time inter-vals to determine CFU (Figure 4B), and showing that each strain was reproducibly injected and the infection progressed as shown above. Alt-hough the experiment was only performed once and should be repeated, macrophage numbers do not seem affected as for K56-2.

Neutrophil numbers increase in shvR in-fected embryos Tg(mpx::GFP) embryos were injected with 20 to 80 CFU of K56-2ΔshvR, K56-2ΔshvR+pUCP28T-shvR (complemented strain) and K56-2. Figure 5A, shows the number of neutrophils at different time points in infected and non-infected embryos. During infection with wild-type bacteria, neutrophil numbers de-creased drastically, already at 24 hpi, causing neutropenia, confirming results obtained by Mesureur et al. (manuscript in preparation, Fig-

ure 5A). However, infection with the shvR mu-tant resulted in an increase in the production of new neutrophils compared to non-infected con-trol embryos. This neutrophilia was observed from 48 hpi onwards.

Additionally, the results obtained with com-plemented strain K56-2ΔshvR+pUCP28T-shvR demonstrate complementation in most embryos to neutrophil numbers found after infection with wild-type K56-2, showing the shvR mutation was almost fully complemented by the overex pression of the shvR gene in trans. Bacterial enumeration was performed as a control to verify infection progression (Figure 5B). Although this experiment should be repeated to analyze a more significant number of embryos, the data show that infection with shvR does not result in neutropenia, as with its wild type parent, but in neutrophillic inflammation.

3. Development of tools to better study the role of ShvR in virulence

Complementation plasmids Studies using genetic mutants require complementation of the mutant phenotype by introducing a plasmid expressing the gene, to verify that the pheno-type is due to the specific gene deletion. In the laboratory the plasmid pUCP28T-shvR, provid-ing trimethoprim resistance (kindly provided by P. Sokol), has thus far been used for comple-mentation studies of the shvR mutation in viru-lence in zebrafish embryos. The complemented strain almost fully restores virulence, measured as embryo survival (Figure 6) and bacterial mul-tiplication (Figure 5B, and Subramoni, Vergunst

2"hpi"" 24"hpi" 42"hpi"Macrophage*K56.2*Neutrophil*

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K56$2ΔshvR'Neutrophil'

K56$2ΔshvR'Neutrophil'

K56$2ΔshvR'Neutrophil'

24#hpi# 48#hpi# 72#hpi#

K56$2ΔshvR'Neutrophil'

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Figure 4 - Macrophage quantification analysis (A) Number of macrophages per Tg(mpx::GFP x mpeg1::mCherry) embryos uninfected (black dots) or infected with ~29 CFU of B. cenocepaci K56 2 (black squares) and ~12 CFU of the shvR mutant (black diamond) at 0, 24, 43 and 72 hpi. The number of macrophages was assessed by pixel counting, each dot represents one embryo. A Student’s unpaired t-test showed that the mutant differed significantly from the WT at 24 (P=0.0064) and 43 hpi (P< 0.0001). We also observed a small but significant difference between the mutant and the non-infected embryos at 24 hpi (P=0.0465). (B) The CFU counts are represented in a scatter plot, and a Student’s unpaired t -test showed that the mutant differed significantly from the WT at 24 hpi (P=0.0005). The 43 hpi time point for K56-2 was not determined due to a problem with the plating.

Figure 5 - Neutrophil quantification analysis. (A) Number of neutrophils per Tg(mpx::GFP) embryos uninfected (black dots) or infected with ~35 CFU of B. cenocepacia K56-2 (black squares), ~76 CFU of the shvR mutant (black diamond) or with ~17 CFU of the complementing strain (black triangle) at 0, 24, 43 and 72 hpi. The number of neutrophils was assessed by pixel counting, each dot represents one embryo. A Student’s unpaired t-test showed that the mutant-infected embryos differed significantly from the non-infected control embryos at 43 (P=0.0010) and 72 hpi (P=0.0004), it also differed significantly from the WT at 24 (P=0.0002) and 43 hpi (P< 0.0001). No significant variation was verified between the WT and the complementing strain. (B) The CFU counts are represented in a scatter plot, and a Student’s unpaired t-test showed that the mutant differed significantly from the WT at 24 (P=0.0004) and at 43 hpi (P=0.0047). At T=43 hpi significant difference between the WT and the comple-menting strain was verified (P=0.0326). et al., unpublished) but the complementation does not reach 100% wild-type levels. The plasmid pUCP28T-shvR has the shvR gene under control of its own shvR promoter.

Although the exact copy number of pUCT28 in B. cenocepacia is not known, overexpression of shvR from plasmid pUCP28T-shvR (in B. gladioli there are over 100 copies per cell (38)), may prevent full complementation for virulence. We decided to clone the pshvR-shvR cassette on two different plasmids, a derivative of pBBR, a medium copy plasmid, and a derivative of the low copy plasmid pMR10 (1-2 copies per cell), called pCR11. In addition, we generally use CmR resistant plasmids in our assays, and therefore we used Cm resistant versions of these plasmids.

To construct pMG3, the shvR gene, includ-ing the promoter sequence (~1.6 kb), was am-

plified by PCR (primers pshvRXhoI and shvRXbaI rev, sequence in Table 2). The blunt end amplification product was ligated into SmaI-digested pUC29 (pMG2). An XhoI/HindIII frag-ment of pMG2 was then inserted in pIN29, cre-ating pMG3. pMG3 also contains a ptac-DsRed reporter gene for simultaneous real time visual-ization of bacteria, thus avoiding the need to transfer two compatible plasmids, each with a different selectable marker, to make the com-plementing mutant strain. The plasmid pMG3 was introduced into B. cenocepacia K56-2ΔshvR by electroporation, and the strain was compared to the original complementing strain, K56-2ΔshvR (pUCP28-shvR) described for the neutrophil counting experiment, in zebrafish infections. Preliminary experiments demon-strated that the new plasmid, pMG3, was not able to fully restore the wild-type virulence

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Figure 6 - pUCP28T-shvR almost fully restores virulence to a shvR mutant. A log-rank (Mantel-Cox) test showed that the mutant differed significantly from the WT (P<0.0001) and from the complementing strain (P<0.0001).

levels of the shvR mutant, as seen with pUCP28-shvR (not shown). These experiments will be repeated.

To construct the second plasmid, a low copy plasmid, the pshvR-shvR casette (~1.7 kb) was isolated from pMG3 using NcoI and NarI re-striction sites and the DsRed reporter gene was obtained from pAV100 plasmid (~800bp, Table

1), using XbaI and ClaI (compatible with NarI) restriction sites. These two fragments were ligated (~2.5 kb) and the gel-extracted fragment was cloned into pUC29, digested with NcoI and XbaI (pMG6). A fragment corresponding to shvR gene and DsRed was then cloned as as XbaI/SpeI fragment into similarly digested pCR11 creating pMG4.

Visualizing shvR expression in vivo in zebrafish embryos In order to analyze the expression of the shvR gene in vivo, during infection in zebrafish embryos, and other exper-imental growth conditions in vitro, a plasmid was constructed that places the reporter gene mCherry under control of the shvR promoter sequence. To construct this plasmid the up-stream region of the shvR gene, encompassing the promoter sequence (596kb), was amplified by PCR (primers pshvR for and pshvR rev, sequence in Table 2) The blunt end amplification product was first ligated into EcoRV-digested pUC29 (pMG1), and the correct sequence veri-fied by sequencing. pMG1 was digested with NdeI and HindIII enzymes (both sites previously added by PCR), and inserted in equally digest-ed pIN233. This resulted in replacement of the tac promoter region in pIN233 with the shvR promoter sequence, placing the mCherry re-porter gene under control of shvR regulatory signals. Read through expression from plasmid encoded sequences was prevented by the presence of a strong termination signal up-stream the shvR-mCherry cassette.

Wild-type K56-2 and the shvR mutant strains were electroporated with the pAV209 plasmid. The colonies on LB-agar plates were bright red fluorescent under a fluorescence microscope, indicating that shvR is expressed under these growth conditions. Also in liquid cultures the bacteria were red fluorescent. A preliminary zebrafish infection experiment showed that K56-2 and the shvR mutant, carry-ing plasmid pAV209, strongly expressed mCherry throughout the infection. The mCherry protein is very stable, and will not allow analysis of changes in expression levels.

Therefore, to be better able to verify if shvR expression is repressed, or activated during infection, including the intracellular stages, an unstable version of the mCherry protein was made. Therefore, we created a small peptide tag based on the studies of unstable GFP vari-ants in Pseudomonas putida (39). We transla-tionally fused the C-terminal peptide tag AANDENYALVA, that has been shown to lead to a faster destabilization of a GFP protein (39), to mCherry in pAV209. Two complementary oligos (Table 2, codon optimized for B. cenoce-pacia and including a TAG terminator codon) were annealed, resulting in cohesive BsrGI and SacI ends. Plasmid pAV209 was digested with BsrGI and SacI enzymes and the fragment was inserted by ligation, resulting in pMG5. The plasmid is now being verified by sequencing, and will be introduced into wild-type K56-2, and the shvR mutant for further analysis.

4 DISCUSSION Recent studies on B. cenocepacia K56-2, an

ET12 strain recovered during an epidemic out-break, described the appearance of shiny vari-ant colonies that encoded a mutation in a global regulator, named ShvR. This regulator belongs to the LysR-type transcriptional regulator family (LTTR), which is a highly conserved family of regulators in bacteria (40). In a BLAST analysis, using the PATRIC bioinformatics tool (41), the gene appeared highly conserved in B. cenoce-pacia and B. cepacia strains, and orthologs of ShvR were not detected in most of the other species in the complex (data not shown). ShvR is encoded on pC3, a non-essential megaplas-mid in B. cenocepacia demonstrated to be

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needed for full virulence in different models (15). In previous experiments with a K56-2 shvR mutant, the bacteria were shown to be less virulent in alfalfa seedlings and showed reduced inflammation in rat lung (30), suggest-ing that ShvR regulates virulence factors in-volved in establishing an acute infection.

Previous studies using a chronic rat agar bead model, showed that the shvR mutant had reduced virulence and reduced lung inflamma-tion; however, the bacteria were able to persist sometimes better than the wild-type in the lungs (30). In addition, the shvR mutant showed low biofilm formation in vitro. Thus ShvR might be important in regulating factors involved in the excessive inflammation and dissemination of the bacteria, and negatively regulating factors needed for a persistent infection.

The aim of this study was to start to better understand the behavior of the shvR mutant and the immune response during infection, using the zebrafish embryo model. The zebrafish has recently been developed as a model for Bcc infections allowing us to detect differences in virulence caused by different strains, for example B. cenocepacia K56-2 de-velops an acute and fatal infection whereas B. stabilis LMG14294 causes a persistent infection (25). My results show that in the absence of ShvR the lethal K56-2 can only cause persis-tent infection in zebrafish. This is seen by re-duced bacterial growth, reduced inflammation and virulence, and absence of host phagocyte cell death, compared to K56-2. Instead the mu-tant somehow activates the host immune re-sponse, observed by the increased number of neutrophils, similar to the response seen to infection with B. stabilis (Mesureur et al., manu-script in preparation). Embryos infected with K56-2 die after 2 dpi with a high number of bac-teria, in line with published data that this strain creates an acute fatal infection (25). In contrast, embryos infected with K56ΔshvR are all still alive at 5 dpi with a relatively constant number of bacteria, consistent with a persistent pheno-type as shown for B. stabilis LMG14294 by Vergunst et al. (25). In agreement with the find-ings in experimental rat infections, these results suggest that ShvR regulates factors involved in acute inflammatory infection caused by K56-2.

Microscopic observations showed that shvR mutant bacteria survived inside host macro-phages, and were still observed 5 dpi. The bac-teria are able to reach high intracellular num-bers. Such infected macrophages sometimes had extreme sizes, but further analysis will be performed to estimate bacterial numbers inside the macrophages, in order to affirm a possible difference between shvR-infected and K56-infected macrophages. Moreover, the im-portance of the macrophages for the bacterial replication and survival should be assessed by performing a macrophage ablation assay.

Additionally, the number of host immune cells was analyzed in order to further demon-strate a persistent infection phenotype. Con-cerning macrophages, Mesureur et al. (manu-script in preparation) found that in K56-2 infect-ed embryos, the number of non-infected mac-rophages decrease rapidly during infection, reaching very low numbers, whereas infected macrophages were kept alive by the bacteria. Similar results were obtained in this study for K56-infected embryos, however, in embryos infected with the shvR mutant the macrophage numbers increased in a similar manner as in control embryos. Although we do not know the reasons for the disappearance of macrophages during K56-2 infection, this shows that the shvR mutant lacks the regulation of the factors in-volved in the macrophage killing, or induces altered innate immune responses leading to the difference in host phagocyte response. When neutrophil numbers during infection were ana-lyzed, it was found that in K56-2 infected em-bryos neutrophils were reduced to very low numbers, as found also by Mesureur et al. (manuscript in preparation). Thus, these em-bryos are not able to resolve the inflammation caused by the bacteria. As for the mutant, it showed that the bacteria induced neutrophilia in the embryos, again similar to an infection with B. stabilis, where Mesureur et al. (manuscript in preparation) verified that the strain induced neutrophil production at higher levels even at 5 dpi. The analyses of host immune cell numbers suggest once more that the infection with the shvR mutant follows a persistent phenotype, similar to B. stabilis infection and strikingly dif-ferent from K56-2 infection.

Earlier, Bernier et al. found that shvR mutant bacteria reached numbers that could be equal or even higher than the wild-type in rats lungs, yet, the mutant showed reduced capacity to form biofilms (30). Together with the results of survival of the shvR mutant in macrophages in zebrafish embryos, this raises the question whether the shvR bacteria could better persist inside cells than the wild-type in the rat lungs. Experimental evidence for this could change the current idea of persistence, and show a more important role for intracellular bacteria in vivo. This would be in line with recent observa-tions that in lung tissue of patients Bcc have been found primarily inside cells (28,29).

During this study, tools were developed to better analyze the role of ShvR in virulence in vivo: new complementing plasmids were creat-ed to optimize complementation studies, and two other plasmids were created to study the expression of shvR in vivo during infection. In this study, genetic complementation of the shvR mutant for virulence in zebrafish embryos was performed using the plasmid pUCP28T-shvR. Although this plasmid that expresses the shvR gene from its own promoter sequence restored virulence to the mutant in zebrafish infections, the complementation was not 100%. It is possi-ble that full complementation is not achieved because, as ShvR regulates over 1000 genes, reaching the correct expression levels of ShvR is important; although copy number of the pUCP28T-shvR plasmid in B. cenocepacia has not been determined exactly, in B. gladioli there are over 100 copies per cell (38). It is thus likely that ShvR is produced at higher levels than from its genomic position.

Therefore two plasmids were constructed: pMG3 with a pBBR backbone vector (CmR), which has a medium number of copies per cell, and pMG4 with a pMR10 backbone vector (CmR), that only has 1-2 copies. Preliminary experiments with plasmid pMG3 indicated that also this plasmid was not able to fully restore virulence to the shvR mutant. Although the con-struction of pMG4 is still underway, it will be interesting to find out if the further reduction in copy number will allow full complementation. It has been shown previously for instance for a virB5 mutant of Brucella, that VirB5 overex-

pression affected expression and/or stability of other VirB proteins, resulting in attenuation, even of the wild-type strain (42). Future experi-ments include transforming wild-type K56-2 with pMG3, the pBBR-based plasmid, to verify if overexpression reduces its virulence (42).

To better understand a role for ShvR in regulating target genes involved in acute, or persistent infection, it is important to know when during infection the shvR gene is expressed. Therefore, a new plasmid that allows for obser-vation of the regulator’s expression in vivo was constructed. The plasmid pAV209 encoding a mCherry fluorescent protein under control of the shvR promoter, was constructed. Although not brightly expressed in E. coli (data not shown), experiments, using K56-2 and the mutant bac-teria transformed with this plasmid, indicated that the expression of ShvR started at some time point during bacterial growth in LB medi-um, since the bacteria were bright red fluores-cent in overnight grown cultures. During infec-tion experiments in zebrafish, the bacteria were visible with fluorescence microscopy throughout the experimental time. Due to the stability of the mCherry protein (although the half-life of mCherry is not known in Bcc, in E. coli the half-life of GFPmut3, which is also very stable, has been shown to be more than 24 h (39)), we cannot study any changes in the expression levels of shvR. To be able to better analyze a possible regulation of ShvR expression in vivo, also in other Bcc strains, an unstable m-Cherry reporter gene was created. In studies with Shi-gella flexneri, a GFP reporter was created that matured faster in order to assess T3SS activity (43). In our study we included a small peptide tag (AANDENYALVA) to the C-terminal end of mCherry to allow its rapid degradation by intra-cellular tail-specific proteases (39). Further experiments, including infection and fluores-cence extinction experiments, will be performed to assess the efficiency of this plasmid.

5 CONCLUSIONS AND FUTURE PERSCPECTIVES

In conclusion, in agreement with other infection models, ShvR has a role in virulence of B. cenoce-pacia in the zebrafish embryo model. The techniques used in this study contributed to confirm and better understand the persistent infection phenotype of B.

cenocepacia K56-2ΔshvR. More detailed studies have to be performed to determine a more precise role for ShvR in regulating factors that determine the difference between acute and persistent infection, using not only the created tools but also other tech-niques, including proteomic and transcriptional stud-ies (dual RNAseq to analyze both bacterial and host transcriptome) that could give more insights in ShvR regulation.

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