Extracellular DNases of Ralstonia solanacearum ...

Environmental Microbiology (2016) 00(00), 00–00

doi:10.1111/1462-2920.13446

Extracellular DNases of Ralstonia solanacearum modulate biofilms and facilitate bacterial wilt virulence

Tuan Minh Tran,1 April MacIntyre,1,2 Devanshi Khokhani,1 Martha Hawes3 and Caitilyn Allen1* 1 Department of Plant Pathology, University of Wisconsin-Madison, Madison, WI 53706, USA. 2 Microbiology Doctoral Training Program, University of Wisconsin-Madison, Madison, WI 53706, USA. 3 Department of Soil, Water and Environmental Science, University of Arizona, Tucson, AZ 85721, USA. Summary Ralstonia solanacearum is a soil-borne vascular pathogen that colonizes plant xylem vessels, a flowing, low-nutrient habitat where biofilms could be adaptive. Ralstonia solanacearum forms biofilm in vitro, but it was not known if the pathogen benefits from biofilms during infection. Scanning electron microscopy revealed that during tomato infection, R. solanacearum forms biofilm-like masses in xylem vessels. These aggregates contain bacteria embedded in a matrix including chromatin-like fibres commonly observed in other bacterial biofilms. Chemical and enzymatic assays demonstrated that the bacterium releases extracellular DNA in culture and that DNA is an integral component of the biofilm matrix. An R. solanacearum mutant lacking the pathogen’s two extracellular nucleases (exDNases) formed non-spreading colonies and abnormally thick biofilms in vitro. The biofilms formed by the exDNase mutant in planta contained more and thicker fibres. This mutant was also reduced in virulence on tomato plants and did not spread in tomato stems as well as the wild-type strain, suggesting that these exDNases facilitate biofilm maturation and bacterial dispersal. To our knowledge, this is the first demonstration that R. solanacearum forms biofilms in plant xylem vessels, and the first documentation that plant pathogens use DNases to modulate their biofilm structure for systemic spread and virulence. Received 11 May, 2016; accepted 1 July, 2016. *For correspondence. E-mail [email protected]; Tel. 608-262-9578; Fax: 608-2632626. C 2016 Society for Applied Microbiology and John Wiley & Sons Ltd V

Introduction Bacterial biofilms are critical for interactions of many animal and plant pathogens with their eukaryotic hosts (Costerton et al., 1999; Morris and Monier, 2003; Parsek and Singh, 2003). Biofilms protect bacteria from stresses such as desiccation, antibiotics and host antimicrobial defenses and help them attach, feed and dispose of waste in fluid environments (Morris and Monier, 2003; Danhorn and Fuqua, 2007; Flemming and Wingender, 2010). Microbial biofilms develop according to a predictable program that begins when cells aggregate on a surface. These sessile cells multiply and produce an extracellular polymeric matrix, which is often regulated by a quorum sensing system (Vu et al., 2009; Limoli et al., 2015). Eventually, a subpopulation of microbes in mature biofilms become motile, escape from the matrix and resume planktonic life; this allows them to disseminate to new sites. Although biofilms vary in composition and structure, they generally consist of live and dead cells embedded in a matrix of polysaccharides, proteins and DNA (Montanaro et al., 2011). Extracellular DNA (eDNA) plays an important structural role in biofilms of environmental and animal pathogenic bacteria, including Pseudomonas aeruginosa, Staphylococcus aureus and Streptococcus spp. (Whitchurch et al., 2002; Allesen-Holm et al., 2006; Moscoso et al., 2006; Brown et al., 2015). These microbes all use extracellular nucleases to partially degrade the biofilm matrix and disperse (Mann et al., 2009; Nijland et al., 2010; Kiedrowski et al., 2011; Steichen et al., 2011). Ralstonia solanacearum is a major vascular pathogen of plants. It causes bacterial wilt disease on over 200 species, including diverse economically important crops such as potato, tomato and banana (Denny, 2006). The pathogen enters host roots through wounds or lateral root emergence sites and colonizes the water-transporting xylem vessels. There the bacteria multiply rapidly, reaching population sizes greater than 108 CFU/ml in this habitat, which is characterized by flowing liquid and relatively low levels of oxygen and nutrients (Else et al., 1994; Vasse et al., 1995; Digonnet et al., 2012; Netting et al., 2012; Dalsing et al., 2015). In vitro, this bacterium forms biofilms that depend on diverse traits, including aerotaxis, the mannose-binding lectin LecM, twitching and swimming motility and the

2 T. M. Tran et al. extracellular polysaccharide that is essential for R. solanacearum virulence (Saile et al., 1997; Kang et al., 2002; Yao and Allen, 2007; Meng et al., 2011; Meng et al., 2015). EPS is thought to contribute to wilt symptoms by blocking host xylem. This high-molecular weight polymer is probably also an essential component of the pathogen’s biofilm matrix (Saile et al., 1997; Morris and Monier, 2003). Ralstonia solanacearum forms microaggregates on tomato root surfaces (Yao and Allen, 2007) and the bacterium formed biofilms on leaf mesophyll cell surfaces when it was artificially infused into tomato leaf apoplast (Mori et al., 2016). However, it is not known if R. solanacearum forms biofilms in its natural habitat in xylem vessels, nor is it known if biofilms contribute to host colonization or bacterial wilt virulence. In a separate study we found that R. solanacearum produces two structurally distinct non-specific endonucleases: NucA, a membrane-localized 30 kDa Mg11-dependent enzyme, and NucB, a secreted 15 kDa cation-independent nuclease (Tran et al., 2016). We showed that R. solanacearum uses these enzymes to degrade microbe-trapping extracellular DNA nets that are released by plant root border cells, thereby enabling the bacterium to invade plant roots more efficiently. Mutants lacking NucA and NucB could not escape from root border cell DNA traps and were less virulent than the wild-type strain in a soil-soak inoculation assay that required the bacterium to infect intact tomato roots (Tran et al., 2016). The DnucA and DnucB mutants were also defective in initial attachment to pea roots. Since bacterial eDNA is known to be important for biofilm formation in many bacteria, we hypothesized that R. solanacearum’s secreted DNases also function after plant infection by modulating biofilm eDNA in xylem vessels. We used a combination of microscopy, in vitro biofilm assays and in planta analyses to describe biofilm formation and dispersal in R. solanacearum and to determine the role of eDNA and DNases in these processes. We found that this pathogen releases eDNA and that DNase can disrupt its biofilms. Virulence assays with mutants missing one or both nucleases suggested that these enzymes are essential for bacterial wilt virulence. Specifically, these nucleases were important for bacterial translocation inside tomato hosts, likely because they degrade biofilm eDNA, thus facilitating the pathogen’s systemic spread during disease. Results Ralstonia solanacearum forms biofilms in host plant xylem vessels Scanning electron microscopy of stem sections from R. solanacearum-infected tomato plants revealed that colonized host xylem vessels contained biofilms, defined here

as multicellular aggregates of bacterial cells embedded in a matrix of amorphous material and fibres (Fig. 1A–C). These masses sometimes occluded the entire vessel (Fig. 1A). The matrix fibres were straight and smooth and varied in thickness from 20 to 30 nm (Fig. 1C). Extracellular DNA is an important component of R. solanacearum biofilm DNA is a constituent of many bacterial biofilms (Flemming and Wingender, 2010). We used confocal microscopy to visualize the structure of biofilm formed by R. solanacearum strain GMI1000 on chambered glass slides. Baclight live/dead staining revealed that R. solanacearum biofilm contained strings of both live and dead cells, with microcolonies connected by threads that were stained by SYTO9, which binds nucleic acids (Fig. 2A). DNase disperses the biofilms of several microbes (Okshevsky et al., 2015). To determine if digestion of DNA could degrade R. solanacearum biofilm, we treated biofilms formed on glass coverslips with increasing concentrations of commercial DNase I. The structure of the biofilm was progressively disrupted as enzyme concentrations increased, and it was almost completely degraded after exposure to 5 lg ml21 DNase I for 1 h (Fig. 2B). This indicated that extracellular DNA is an essential structural element of R. solanacearum biofilm. R. solanacearum secretes two non-specific endonucleases, NucA and NucB (Tran et al., 2016). Deletion mutants lacking one or both extracellular nucleases grew as well as the wild-type strain in both rich and minimal media (Tran et al., 2016). Colonies of these mutants were indistinguishable from those of wild type on regular CPG plates. However, on soft agar plates the DnucA, DnucB and DnucA/B mutant strains formed colonies noticeably different from those of wild type. Colonies of the nuclease mutants were round rather than irregular and did not spread like those of the wild-type strain, suggesting that DNA also plays a structural role in the bacterium’s extracellular matrix on plates (Fig. 3A). In addition, all three nuclease-deficient mutants formed significantly more biofilm than the wild-type strain on PVC plates, as measured by a spectrophotometric crystal violet assay (Fig. 3B). An ELISA assay with anti-EPS1 antibody found that the nuclease mutants produced wild-type EPS levels, indicating that their increased biofilm formation did not result from altered EPS production (Fig. S1). To determine how extracellular DNase changes the structure of R. solanacearum biofilm, we used confocal microscopy to visualize the biofilms formed on chambered glass slides by wild-type strain GMI1000 and the DnucA/B double nuclease mutant. The wild-type strain formed microcolonies, as previously observed (Mori et al., 2016);

C 2016 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 00, 00–00 V

Extracellular DNases modulate R. solanacearum biofilm and virulence 3

Fig. 1. Biofilms formed by wild-type and nuclease-deficient R. solanacearum strains in tomato plant xylem. Tomato plants were inoculated with 2000 bacterial cells through a freshly cut leaf petiole and sampled after 4 days. For each plant, two stem cross-sections were fixed, sputter-coated and visualized under a high-resolution scanning electron microscope (Zeiss LEO-1530 FE-SEM). A–C. Wild-type strain GMI1000. D–F. Extracellular nuclease-deficient DnucA/B mutant. Threads approximately the diameter of typical 30 nm chromatin-like fibres were present in the aggregates formed by both wild-type and DnucA/B strains in plant stems (C and F). Wild-type strain aggregates contained strikingly fewer threads, and these threads were thinner and more variable (C) whereas they were more uniform in thickness in the biofilm formed by the DnucA/B mutant (F). Images are representative of three independent biological replicates, each containing samples from three plants. Images in the same column had the same magnification.

this biofilm was thin and consisted of only a few layers of cells. In contrast, biofilm formed by the nuclease-deficient mutant was thicker and more complex (Figs 3C and S2). Since deletion of both extracellular nuclease genes resulted in biofilm overproduction, it is likely that these enzymes normally modulate biofilm structures by degrading the DNA in the extracellular matrix. To determine if R. solanacearum cells release DNA that could be integrated into biofilms and if that DNA was affected by the pathogen’s secreted nucleases, we compared the eDNA levels in spent culture medium following growth of wild-type and the nuclease-deficient DnucA/B mutant. DNA was present in spent medium of both strains after 24 h (Fig. 4A). However, while eDNA levels decreased over time in cultures of wild-type GMI1000, it continued to accumulate in cultures of DnucA/B strain for 96 h. This suggests that the double nuclease mutant did not degrade eDNA as effectively as the wild-type strain. We characterized eDNA in cell-free supernatant from GMI1000 cultures using PCR with primers specific for several sites spaced around the GMI1000 chromosome (759/ 760, gspM-F/R, Endo-F/R) and one site on the megaplasmid (Rsol_fliCF/R). eDNA from culture supernatants yielded the same PCR products as purified DNA from

lysed bacteria (Fig. 4B), suggesting that the eDNA is composed of total genomic DNA rather than only one or a few specific sequences.

Purified NucA and NucB disrupted R. solanacearum biofilm Extracellular DNA and secreted nucleases could be important for both initial bacterial attachment and for modification of mature biofilms. To determine if exogenous application of R. solanacearum’s extracellular nucleases affect its biofilms, we allowed the wild-type strain to form biofilms on PVC plates for 24 h and then treated them with purified NucA or NucB protein. Only 0.01 lg ml21 of NucA was enough to significantly degrade these mature in vitro biofilms. In contrast, 1 lg ml21 of NucB was required for similar effects (Fig. 4C and D). This suggests that NucA may make a larger contribution to biofilm modification, at least under these assay conditions. It should be noted that the overexpressed and purified NucA lacks 55 N-terminal amino acids predicted to embed the protein in the bacterial outer membrane; this deletion may have altered the enzyme’s activity (Tran et al., 2016).


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Fig. 2. DNA is an important component of R. solanacearum biofilm. A suspension of wild-type strain GMI1000 in rich medium was seeded into 24-well plates with glass coverslips at the bottom of each well and incubated for 72 h without shaking at 288C to allow biofilm formation. A. Visualization of extracellular DNA in R. solanacearum biofilm stained with SYTO9 (live cells) and propidium iodine (PI, dead cells). Live and dead bacterial cells were connected by DNA-containing strings (white arrowheads in merged image) in the biofilm (bar 5 10 lm). B. Exogenous application of DNase I disrupted R. solanacearum biofilm. Coverslips were washed twice with PBS to remove unbound cells and then incubated at 378C for 1 h with the indicated concentrations of DNase I and stained with crystal violet.

Extracellular nucleases are required for normal biofilm structure in plant xylem To determine if the ability to digest extracellular DNA affects the structure of R. solanacearum biofilms inside plant hosts, we used high-resolution scanning electron microscopy to compare stems of tomato plants infected with either wild-type or the DnucA/B mutant. Following petiole inoculation of tomato plants, DnucA/B multiplied extensively in host xylem, like its wild-type parent. However, the DnucA/B aggregates contained a much denser web-like matrix of fibres that were thicker, rougher and kinked, while fibres in the wild-type biofilm were thinner, smoother and straighter (Fig. 1D and E). In shape and size, the DnucA/B biofilm fibres resembled the eDNAcontaining ‘30 nm chromatin-like fibres’ that were observed in other bacterial biofilms (Griffith, 1976; Wu et al., 2007). They were more homogenous in diameter (30 nm) than the thinner, more variable fibres observed in wild-type biofilm (20–30 nm) (Fig. 1C and F). Biofilms formed by the DnucA/B mutant in planta contained fibre strands associated with both cells deep in the biofilm matrix interior and cells in the outermost layers of the biofilm. In contrast, few fibres were associated with bacteria on

the surfaces of wild-type biofilm. These observations suggest that R. solanacearum requires extracellular DNases to develop its normal open biofilm architecture in xylem tissue.

Extracellular DNases are required for full virulence of R. solanacearum We previously observed that NucA and NucB are required for full R. solanacearum virulence when the pathogen infects from the soil and must overcome DNA-containing traps released by plant root border cells (Tran et al., 2016). Our scanning electron microscopy (SEM) analyses showed that R. solanacearum needs extracellular DNases to develop normal biofilm in xylem, so these enzymes may also contribute to virulence after the bacterium has colonized the plant. To test this hypothesis, we bypassed the root border cell trap defenses by introducing the bacteria directly into the xylem of wilt-susceptible tomato plants through a cut leaf petiole. All three exDNase mutants (DnucA, DnucB and DnucA/B) were slightly but significantly reduced in virulence compared to their wild-type parent strain GMI1000 (Fig. 5) (P < 0.05, repeated measures


Extracellular DNases modulate R. solanacearum biofilm and virulence 5 Fig. 3. Ralstonia solanacearum exDNase mutants form abnormal colonies and biofilms. A. Single and double nuclease R. solanacearum mutants formed denser round colonies on 0.8% agar CPG plates. B. Biofilms formed by wild-type GMI1000 and exDNase mutants in CPG broth on crystal violet-stained PVC plates, measured as A590. Bars represent standard error. Columns with different letters are different from each other (P < 0.05, Tukey’s HSD test). C. Orthogonal views of 4-dayold biofilms formed by wild-type GMI1000 and the extracellular nuclease-deficient DnucA/B mutant strain on glass chambered slides, as observed by confocal microscopy. COMSTAT analysis of these data is presented in Fig. S2.

ANOVA). Thus, DNases are necessary for full virulence of R. solanacearum even after the bacterium has entered host roots. Extracellular nucleases facilitate bacterial translocation in tomato stems To determine if extracellular DNases of R. solanacearum contribute to the bacterium’s systemic spread inside the plants, we used fluorescence microscopy to track the movement of GFP-tagged wild-type GMI1000 and DnucA/ B strains following cut-petiole inoculation into tomato stems. After 7 days, wild-type R. solanacearum had spread laterally from the inoculated xylem vessels to adjacent vessels around the vascular ring, and bacteria were abundantly present two and four cm above and below the inoculation point, indicating that they had moved both with and against the direction of xylem flow, consistent with previous observations (Fig. 6A) (Addy et al., 2012). In contrast, the nuclease-deficient DnucA/B double mutant was visible in only a few xylem bundles at the site of inoculation. The mutant did not translocate around the tomato stem vascular bundles as well as the wild-type parent, as indicated by the percentage of GFP signal/stem area in wild-type and DnucA/B inoculated plants at 2 and 4 cm above and below the inoculation site (P < 0.05, Student’s t-

test) (Fig. 6B). Importantly, this difference in the spatial distribution of the wild-type and mutant strains was not explained by a difference in bacterial population size. Ground dilution plated tomato stem sections contained statistically similar numbers of wild-type GMI1000 and the DnucA/B double mutant at all sampled locations, although the DnucA/B mutant population sizes did trend smaller than those of wild-type at 4 cm above and below the inoculation site (Fig. 6C). This indicated that DnucA/B multiplied in susceptible tomato plants as well as its parent strain, at least for the first few days after stem inoculation. We suspect that the DnucA/B mutant occupied less stem area because it was unable to escape from its own biofilms in planta, resulting in reduced systemic colonization along and across xylem vessels. The two-component regulatory system VsrBC negatively regulates extracellular DNase activity Measuring DNase activity produced by R. solanacearum mutants lacking various virulence regulators revealed that most had no significant effect on DNase activity (Fig. 7). However, a vsrBC mutant produced fivefold more DNase activity than the wild-type strain (P < 0.0001, one-way ANOVA). This indicates that VsrBC, a two-component regulatory system that controls motility as well as EPS


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Fig. 4. Extracellular DNases and eDNA modulate biofilm formation in R. solanacearum. A. Kinetics of extracellular DNA release in broth cultures of wild-type strain GMI1000 and the DnucA/B double mutant, quantified spectophotometrically. B. Extracellular DNA of R. solanacearum contains regions from diverse genomic locations. Extracellular DNA extracted from cell-free spent media of GMI1000 was amplified with primers that target different regions on the R. solancearum chromosome (759/760, gspM-F/R, Endo-F/ R) and megaplasmid (Rsol_fliCF/R) (M: molecular weight markers, G: total genomic DNA control, E: extracellular DNA from culture supernatants). Purified NucA (C) and NucB (D) degraded preformed biofilm on PVC. Wild-type strain was allowed to form biofilm for 24 h in rich CPG broth and then treated with either NucA or NucB for an additional 6 h at 288C. Crystal violet-stained biofilm was quantified spectrophotometrically as A590. Asterisks indicate treatments different from water-treated control biofilm (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, one-way ANOVA).

production (Yao, 2007; Meng et al., 2011), is also a negative regulator of extracellular DNase activity. Discussion Microbes living in a fluid environment like a river, a catheter or a plant xylem element form biofilms to remain in one

place, absorb nutrients, shed waste and communicate with each other. Biofilms also shelter microbes from a range of stresses including desiccation, radiation and, in the case of pathogens, host defense responses (Morris and Monier, 2003; Ramey et al., 2004). More than 80% of human infections are associated with organisms in biofilms (Bryers, €mling and Balsalobre, 2012). The success of 2008; Ro

Fig. 5. Extracellular DNases contribute to bacterial virulence on tomato. Three-week-old Bonny Best tomato plants were inoculated by placing 500 R. solanacearum cells on a freshly cut petiole. WT, wild-type strain GMI1000, DnucA single mutant (A), DnucB single mutant (B), or DnucA/B double mutant (C). Disease severity was rated daily for 14 days using a disease index scale, in which 0: no wilted leaves; 1: 1–25% leaves wilted; 2: 26%–50% leaves wilted; 3: 51%–75% leaves wilted and 4: >75% leaves wilted. Data represent the means of three independent experiments; each experiment included 14 plants per treatment and error bars represent standard errors. Disease progress curves of plants inoculated with nuclease mutants were different from that of wild-type (P < 0.05, repeated measures ANOVA). C 2016 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 00, 00–00 V

Extracellular DNases modulate R. solanacearum biofilm and virulence 7

Fig. 6. Ralstonia solanacearum needs extracellular DNases to spread in tomato stems, but not to multiply. Four-week-old tomato plants (cv. Bonny Best) were petiole-inoculated with GFP-expressing variants of either wild-type GMI1000 or the DnucA/ B double nuclease mutant. A. Stem sections from several locations from the point of inoculation were sampled three days after inoculation and observed under a fluorescence dissecting microscope equipped with a GFP filter set. Red signal is plant tissue autofluorescence. B. The percent area of each cross section colonized by wild-type and DnucA/B strains was quantified using ImageJ 1.48. The experiment was repeated twice with 10 plants for each biological replicate (P < 0.05, Student’s t-test). C. GMI1000 and DnucA/B mutant reached comparable population sizes in plant stems (P > 0.05, Student’s t-test). The experiment was repeated twice, with 10 plants per treatment in each experiment.

Fig. 7. Extracellular DNase activity is affected by the twocomponent system VsrBC. The DNase Alert Kit was used to measure nuclease activity of culture supernatants of R. solanacearum wild-type strain GMI1000 and regulatory mutants grown in minimal medium. Activity was adjusted to bacterial population size as determined by dilution plating and normalized to wild-type DNase activity (WT 5 1). Bars represent mean of three experiments, each with three technical replicates. Asterisks indicate significant difference from WT (P < 0.0001, one-way ANOVA).

several bacterial plant pathogens also depends on their ability to form biofilms on biotic surfaces (Ramey et al., 2004). These include the soft-rot pathogen Dickeya dadantii (Rojas et al., 2002), the epiphytic and apoplastic pathogen Pseudomonas syringae (Dulla and Lindow, 2008; Yu et al., 2014), and the xylem-dwelling wilt pathogens Xylella fastidiosa (Feil et al., 2003), Pantoea stewartii (Koutsoudis et al., 2006) and Erwinia amylovora (Koczan et al., 2009; Koczan et al., 2011). We found that R. solanacearum formed dense biofilm structures inside tomato xylem vessels. This environment contains few nutrients, little oxygen and toxic host defense compounds (Dalsing et al., 2015; Lowe et al., 2015). Additionally, tomato xylem sap flows fairly rapidly during active transpiration (estimated at 35–80 ll min21); this likely exerts drag and shear forces that hinder bacterial attachment to vessel walls (Else et al., 1994; Netting et al., 2012). Ralstonia solanacearum cells in biofilms can probably survive these stresses better than single planktonic cells. In addition, the biofilm matrix immobilizes the pathogen against xylem cell walls, facilitating injection of type IIIsecreted effectors into adjacent xylem parenchyma (Genin and Denny, 2012). Although it was recently reported that R. solanacearum strain OE1-1 formed mushroom-like


8 T. M. Tran et al. biofilm structures on leaf mesophyll cells following injection of the bacteria into tomato leaves, we did not observe mushroom shapes in the bacterial aggregates formed in xylem vessels of tomato plants infected by strain GMI1000 (Mori et al., 2016). This difference in behaviour could be strain-specific, but it is more likely that R. solanacearum does not form normal biofilms in the static environment of the leaf apoplast since it is not a leaf pathogen. The extracellular matrix of R. solanacearum biofilms in tomato xylem vessels included many smooth fibres that appeared to connect and contain bacterial cells. Our experiments indicated that these fibres contain eDNA and that eDNA production and degradation play critical roles in R. solanacearum biofilms. First, biofilms formed in vitro were dissolved by purified nuclease. Second, extracellular DNase-deficient mutants formed abnormally thick biofilms in vitro, and the in planta biofilms of the nuclease mutant were qualitatively different from those of the wild-type strain, with cells embedded in a much denser web of 30 nm diameter rough fibres that were similar in size and structure to the DNA cables documented in biofilms of Enterococcus feacalis (Barnes et al., 2012). Extracellular DNA has been reported in biofilms of many bacterial species, including clinically important human pathogens such as P. aeruginosa, S. aureus, Listeria monocytogenes and E. feacalis (Allesen-Holm et al., 2006; Rice et al., 2007; Harmsen et al., 2010; Montanaro et al., 2011; Barnes et al., 2012; Svensson et al., 2014). Among plant pathogenic bacteria, eDNA has been found in biofilm of X. fastidiosa (Cheng et al., 2010) and Xanthomonas citri lez et al., 2016). eDNA promotes bacterial adhe(Sena-Ve sion, especially to hydrophobic surfaces, through acidbase interactions (Das et al., 2010), and it also stabilizes biofilms by interacting with other matrix components (Harmsen et al., 2010; Jennings et al., 2015; Schwartz et al., 2015). Interestingly, biofilm eDNA can induce a drug-resistant state (Mulcahy et al., 2008; Rose et al., 2015; Wilton et al., 2015). What is the source of the eDNA in R. solanacearum biofilms? Although we cannot rule out the possibility that the biofilm fibres observed in xylem contain plant DNA, the thicker biofilms formed by DNase mutants in vitro and the digestion of in vitro biofilms by purified DNase both suggest that the eDNA comes from the bacterium. The mechanism of eDNA release in R. solanacearum is unknown. One possibility is that it arises passively following lysis of dead bacteria; live/dead staining revealed numerous dead cells in R. solanacearum biofilms (Fig. 2A). Alternatively, the eDNA could be the result of secretion, possibly through outer membrane vesicles (Dorward and Garon, 1990; Kulkarni and Jagannadham, 2014; Grande et al., 2015), or from active lysis of a subpopulation (Allesen-Holm et al., 2006; Mann et al., 2009; Paganelli et al., 2013; Beltrame et al., 2015). On-going studies are

exploring the role of bacterial autolysins in R. solanacearum eDNA release and biofilm maturation. It was previously reported that exogenous DNA increases X. fastidiosa biofilm formation in vitro (Cheng et al., 2010). However, adding purified genomic DNA did not enhance R. solanacearum biofilm formation in PVC plates even at 1 lg ml21 (data not shown). Because the extracellular polysaccharides of X. fastidiosa and R. solanacearum are chemically unrelated, they may interact differently with eDNA. Interactions between exogenous eDNA and other components of the biofilm, like proteins, could also explain this difference between the two xylemdwelling pathogens. Like many other microbes, R. solanacearum secretes DNases. Our data indicate that one function of these enzymes is to modify the bacterium’s extracellular matrix, both in planta and in vitro. Ralstonia solanacearum can be identified in culture by its characteristic fluid, irregular, highly mucoid colony morphology. Our results suggest that this morphology results from degradation of eDNA in the extracellular polymeric matrix by secreted DNases. Ralstonia solanacearum nuclease mutants formed atypically round and condensed colonies on plates, indicating that the bacterium releases eDNA when it grows on agar surfaces, as well as in biofilms in vitro. Extracellular DNases of the human pathogen S. aureus also mediate colony spreading (Kaito et al., 2011) and facilitate bacterial escape from biofilm (Kiedrowski et al., 2011; Seper et al., 2011; Kiedrowski et al., 2014). Extracellular DNases are required for normal R. solanacearum biofilm structure in planta, as indicated by the abnormally dense web-like mats formed in xylem vessels by the exDNase-deficient mutant. We speculate that the more open structure and the straighter, thinner and less abundant fibres in wild-type biofilms result from extracellular DNase degradation of these fibres, which may consist of many eDNA strands twisted together into yarn-like strings. Extracellular DNases play similar roles in biofilm maturation in other microbes (Nijland et al., 2010; Okshevsky et al., 2015). It has been proposed that eDNA plays dual sequential roles in biofilms. Initially, it contributes to structural stability of developing biofilms, but as biofilms mature the DNA in the matrix hinders bacterial escape and may also restrict liquid flow through the biofilm; at that point, partially degrading the matrix eDNA increases bacterial fitness (McDougald et al., 2012). Ralstonia solanacearum mutants lacking one or both of the pathogen’s extracellular endonucleases, NucA and NucB, were less virulent than wild type in a direct petiole inoculation assay that bypassed the DNA-containing extracellular traps released by plant root border cells. Interestingly, the single and double mutants had similarly reduced virulence, suggesting that loss of either enzyme is as bad as a loss of both. This may indicate that surface-


Extracellular DNases modulate R. solanacearum biofilm and virulence 9 bound, cation-dependent NucA and freely secreted, cofactor-independent NucB are not functionally redundant. This reduction in virulence did not result from poor multiplication by the nuclease mutants, because the DnucA/B and wild-type strains reached comparably high population sizes (up to 1010 CFU/g tissue) at each location sampled along infected tomato stems. However, the lateral spreading pattern of GFP-tagged bacteria in stem cross-sections revealed that the DnucA/B mutant was present in fewer xylem vessels and vascular bundles than the wild-type strain. Why would a hyper-biofilm forming strain spread poorly in planta? Dispersal of bacterial cells from biofilm is an essential and highly regulated process; without it, bacteria become trapped in their own biofilms and cannot move on to colonize new sites (McDougald et al., 2012). In the insecttransmitted pathogen X. fastidiosa, bacterial attachment and biofilm formation are regulated by quorum sensing (Ionescu et al., 2014). Xylella fastidiosa mutants that adhere excessively and form aberrantly thick biofilms have reduced fitness because they spread poorly within plants and are not efficiently transmitted by their leafhopper vector (Newman et al., 2004; Ionescu et al., 2014). Similarly, the more abundant eDNA in the biofilm of the R. solanacearum DNase mutant apparently limits bacterial spread inside plant hosts. This is likely because this mutant does not go through the normal biofilm maturation process, which includes degradation of some biofilm eDNA. Thus, one function of R. solanacearum exDNases is to digest the eDNA in the biofilm matrix to allow some bacteria to escape from established biofilms, spread systemically inside the plant and eventually return to saprophytic life in soil or water. This suggests that targeted disruption of R. solanacearum biofilms in planta could improve control of bacterial wilt disease. The reduction in spreading of the nuclease mutant is similar to the phenotype caused by the natural R. solanacearum phage URSM3; phage-infected R. solanacearum cells displayed abnormal colony morphology, spread less efficiently across tomato stems and were less virulent (Addy et al., 2012). It would be interesting to determine if phage URSM3 affects the bacterium’s extracellular nuclease activity. Expression of R. solanacearum virulence factors is orchestrated by a complex multicomponent regulatory network (Genin and Denny, 2012). Among several regulatory mutants tested for extracellular DNase activity, only a strain lacking vsrBC differed from wild type. VsrBC is a twocomponent regulator that positively regulates swimming motility, EPS production and virulence (Huang et al., 1993). Like many other virulence regulators, vsrBC expression is affected by the PhcA quorum sensing system, but also by other unknown factors (Huang et al., 1993; Meng et al., 2011). A mutant lacking VsrBC may mimic the behaviour of sessile R. solanacearum cells living in a biofilm. As described above, it appears that degradation of

eDNA facilitates bacterial dispersal from mature biofilms. It would be logical for newly planktonic bacterial cells to upregulate motility to improve spread and coordinately downregulate extracellular DNase production to conserve energy and facilitate eventual DNA-dependent biofilm formation on uncolonized surfaces. We propose a model for R. solanacearum biofilms in planta in which planktonic cells attach to the xylem surfaces and release eDNA either by active autolysis or passively as a result of cell death (Fig. 8). This eDNA facilitates aggregation of cells to form thicker biofilms. At high cell densities or inside developing biofilms, the PhcA quorum sensing system downregulates motility and increases EPS production, promoting biofilm formation (Brumbley and Denny, 1990; Clough et al., 1997). High levels of active PhcA lead to subsequent upregulation of the VsrBC two-component system, which induces exDNase activity and loosens the scaffold structures within the maturing biofilm. Gene expression profiles of cells at biofilm boundaries are likely different than those of cells deep in biofilm, because they experience different environmental conditions (Xu et al., 1998; Lenz et al., 2008). Unknown environmental cues may override the expression of DNase early in biofilm formation to keep eDNA degradation in check and avoid premature dispersal of bacterial cells. The timing of biofilm dispersal may be delayed until the eDNA matrix is loosened substantially, since flagellin can be blocked by EPS at high cell densities (Blair et al., 2008). Alternatively, extracellular DNase activity could also promote expansion of established colonies on the xylem wall. These possibilities are not mutually exclusive and both warrant further investigation. Taken together, our data indicate that eDNA plays a key structural role in R. solanacearum biofilm formation and that the pathogen secretes DNases that modulate biofilm structure to facilitate bacterial dispersal within plants, thereby contributing to bacterial virulence. The more open structure of mature biofilms may also optimize flow through the structure, improving feeding and waste removal. Several lines of evidence indicate that extracellular DNase activity is coordinated with motility to form biofilms in the dynamic flowing environment of xylem vessels. Further studies are needed to elucidate how and when R. solanacearum biofilm eDNA is released, how the DNA-containing fibres interact with other biofilm components, if plant DNA is present in the biofilm matrix in xylem vessels and what factors trigger biofilm maturation and degradation during bacterial wilt pathogenesis. Experimental procedures Media and growth conditions Strains and plasmids used in this study are listed in Table S1. Ralstonia solanacearum cultures were maintained on either Casamino Acids-Peptone-Glucose (CPG) broth or solid


10 T. M. Tran et al. Fig. 8. Proposed model for in planta biofilm formation and DNase-dependent dispersal of biofilm in R. solanacearum. The pathogen’s biofilm is made of eDNA, EPS and other unknown components. When the biofilm is mature and/or nutrients are depleted, localized expression of extracellular DNases degrades eDNA in the matrix to loosen the biofilm structure. Dispersal cells, which are primed by DNases, become motile and disperse into the xylem fluid. These founder cells become attached to xylem walls and eDNA release (via an unknown mechanism) facilitates bacterial aggregation as well as biofilm formation. DNases may also facilitate expansion of established colonies on xylem walls.

CPG 1 TZC (1.8% agar 1 0.05% tetrazolium chloride) at 288C (Kelman, 1954). Escherichia coli was grown in Luria–Bertani medium at 378C. For nuclease assays, strains were grown in Boucher’s minimal medium (Boucher et al., 1985). Culture media were supplemented with antibiotics as needed: kanamycin (25 mg ml21), gentamicin (25 mg ml21) and tetracycline (15 mg ml21).

Strain constructions The DnucA/B GFP-expressing strain was created by transforming DnucA/B with genomic DNA of K60-GFP (Yao and Allen, 2007) by natural transformation as previously described (Bertolla et al., 1997). Transformants were selected on CPG amended with gentamicin, kanamycin and tetracycline and screened for fluorescence using an Olympus BX60F5 compound fluorescence microscope (Olympus, Japan) equipped with a GFP filter set. To create R. solanacearum strain GMI1000 flhDC and vsrBC deletion mutants, 500 bp upstream and downstream of the target genes were amplified by PCR using Kapa Hifi polymerase (Kapa Biosystems, Wilmington, MA). The fragments were then joined to both sides of a gentamicin resistance cassette (amplified from pUCGM) by splicing overlap extension PCR (Horton et al., 1989). Desired PCR products were gel-purified using Zymo Gel Extraction kit (Zymo Research, Irvine, CA), ligated to pCR-Blunt and transformed into E. coli TOP10. Transformants were selected on LB 1 kanamycin and confirmed by PCR and sequencing with primers M13F/R before subcloning into pST-

Blue1 using EcoRI. The resulting deletion constructs in suicide plasmids were electroporated into GMI1000. Gm-resistant transformants were confirmed for correct deletions by PCR using the diagnostic primers listed in Table S2.

PVC biofilm assay To measure biofilm formation of R. solanacearum, we used a standard PVC assay (Yao and Allen, 2007). Briefly, cell pellets from overnight bacterial cultures in CPG were re-suspended in fresh CPG and adjusted to A600 5 0.1 (108 CFU ml21). Five microliters of this suspension were seeded into each well of a PVC 96-well microtiter plate containing 95 ll of fresh CPG. Plates were sealed with Breathe-Easy membrane (Sigma– Aldrich, St Louis, MO) and incubated statically at 288C. After 24 h, each well was stained with 25 ll of 1% w/v crystal violet for 25 min at room temperature, PVC plates were washed twice with 200 ll of sterilized water, and the remaining liquid was aspirated from the bottom of the plate. Adhering crystal violet was dissolved in 200 ll of 95% ethanol, transferred to a new polystyrene plate and measured as absorbance at 590 nm using a Synergy HT microtiter plate reader (Biotek Instruments, Winooski, VT).

Treatment of R. solanacearum biofilm on glass with DNases We used a glass biofilm assay to determine if DNA is a component of R. solanacearum biofilm (Tetz et al., 2009). One


Extracellular DNases modulate R. solanacearum biofilm and virulence 11 8

21

millilitre of a 10 CFU ml bacterial suspension was seeded into each well of a 24-well plate with a round glass coverslip at the bottom of each well and incubated at 288C without shaking for 72 h. Spent medium was aspirated and wells were washed once with PBS. Different concentrations of DNase I (Ambion, Life Technologies, Carlsbad, CA) were added and plates were incubated at 378C for 1 h. The bacterial biofilm on the glass well bottoms was stained with 1% crystal violet, washed twice with PBS buffer and imaged with a Leica DMLB compound light microscope (Leica Biosystems, Buffalo Grove, IL). To confirm that R. solanacearum biofilm is dispersed by the bacterium’s two extracellular nucleases, GMI1000 biofilm were grown on glass-bottomed wells as described above except that the initial bacterial suspension was at A600 5 0.1 to maximize the amount of biofilm available for nuclease digestion. After 24 h, purified NucA and NucB enzymes from overexpressing E. coli strains (Tran et al., 2016) were added and incubated for another 6 h at 288C. Biofilm was quantified using the crystal violet assay as described above. The experiment was repeated twice, with 12 technical replicates for each biological replicate.

Visualization of extracellular DNA in biofilm by confocal microscopy To determine if eDNA is present in R. solanacearum biofilms in vitro, we visualized structures formed by GMI1000. Briefly, 500 ll of bacterial suspension in CPG was seeded into each well of an 8-well glass chambered slide and incubated at 288C, changing the medium daily to maintain bacterial growth. After 4 days, spent medium was removed by pipetting and wells were washed with PBS. Baclight live/dead stain diluted in PBS (Life Technologies, Carlsbad, CA) was added to each well following the manufacturer’s protocol, incubated at room temperature for 15 min and washed twice with PBS gently to avoid disruption of the biofilm. The glass slides were separated from the plastic chambers, mounted with Prolong Gold Antifade reagent (Life Technologies, Carlsbad, CA) and sealed with a cover glass. Bacterial biofilm was visualized using a Zeiss Elyra 780 confocal laser scanning microscope (Carl Zeiss, Germany) with appropriate filter sets for SYTO9 and propidium iodine. Confocal imaging was performed at the Newcomb Imaging Center (Department of Botany, University of Wisconsin-Madison).

Visualization of R. solanacearum biofilm in planta by SEM Scanning electron microscopy was conducted using a modified protocol (Koutsoudis et al., 2006). Three-week-old tomato plants (cv. Bonny Best) were inoculated by placing 2000 cells of either GFP-expressing variants of GMI1000 wild type or the DnucA/B double nuclease mutant on a freshly cut petiole. Plants were grown at 288C with 12 h light/day and sampled at 4 days post-inoculation stems were surfacesterilized by wiping briefly with 70% ethanol, two thin slices from each stem were excised using a sterilized razor blade and immediately put into primary fixative (2.5% glutaraldehyde, 2% formaldehyde, 0.003M MgCl2, 0.003M CaCl2 in 0.05M PIPES buffer, pH 7) for 8 h. Another set of stems from

each plant was sectioned and observed under an epifluorescence microscope; only plants colonized by GFP-tagged R. solanacearum were processed further. Stem sections in primary fixative were then washed twice with 0.05 M PIPES buffer and fixed overnight in secondary fixative (1% OsO4 in PIPES). Samples were washed twice with PIPES the next day and dehydrated with an ascending series of ethanol ending at 100%, followed by critical point drying. Stem sections were sputter coated with gold-palladium and examined with a Zeiss LEO 1530 high-resolution scanning electron microscope at the Material Sciences Center, University of Wisconsin-Madison. The experiment included three biological replicates with three plants per treatment.

Quantification of bacterial eDNA in culture To measure the kinetics of eDNA release in bacterial cultures, we grew R. solanacearum in test tubes containing 10 ml of CPG at 288C with agitation (225 r.p.m.) for 4 days. Test tubes were sampled every 24 h and the bacterial suspensions were transferred to a 15 ml conical tube, vortexed vigorously and centrifuged at 6000 r.p.m. for 5 min. The supernatant was carefully moved to a new 15 ml conical tube by pipetting after passage through a 0.2 mm filter to remove remaining bacterial cells. Five millilitre of filtrate from each sample was then transferred to a DNA concentrating column (Zymo Research) affixed to a Qiagen Manifold with vacuum (Qiagen, Germany). The column was washed with washing buffer and DNA was eluted in 35 ml of RNase-free water by spinning the column at 13000 r.p.m. for 2 min. DNA concentrations were determined spectrophotometrically as described (Allesen-Holm et al., 2006). The experiments were repeated twice, with three technical replicates for each time point. The eDNA in R. solanacearum culture supernatants was characterized by polymerase chain reaction (PCR) with primers specific for the R. solanacearum chromosome (egl, 759/760 and gspM) and the megaplasmid (Rsol_fliC-F/R). Fifty micrograms of GMI100 eDNA or genomic DNA extracted from lysed cells by Epicentre genomic DNA extraction kit according to the manufacturer’s instruction (Epicentre, Madison, WI) was used as template for PCR, using the following programs: (i) 759/760 primers: 968C for 3 min, 35 cycles of 948C for 15 s, 588C for 30 s and 728C for 30 s; followed by 728C for 5 min with expected product size of 230 bp (Opina et al., 1997); (ii) Endo-F/R primers: 968C for 5 min, followed by 30 cycles of 958C for 1 min, 708C for 1 min 728C for 2 min; with a final extension step of 728C for 10 min with expected product size of 750 bp (Poussier et al., 2000); (iii) Rsol_fliCF/R primers: 948C for 5 min, followed by 35 cycles at 948C for 45 s, 688C for 1 min, with no final exten€nfeld et al., sion and the expected product size of 400 bp (Scho 2003); (iv) gspM-F/R primers: 948C for 5 min, followed by 35 cycles at 948C for 30 s, 618C for 30 s and 728C for 30 s, with a final extension step at 728C for 10 min and an expected product size of 427 bp). Polymerase chain reactions (PCR) were analysed by electrophoresis in 1% agarose gel.

Quantification of extracellular polysaccharide We used double antibody sandwich ELISA (DAS-ELISA) to quantify EPS produced by GMI1000 wild-type and nuclease


12 T. M. Tran et al. mutants (Dalsing and Allen, 2014). Briefly, cells from overnight cultures of R. solanacearum strains in CPG were collected by centrifugation, rinsed, re-suspended in sterile water and adjusted spectrophotometrically to A600 5 1.0. One-hundred microliters of each suspension was spread on fresh CPG plates and incubated at 288C for 3 days. Cells were collected by scraping plates in 5 ml of sterile water and 10 ll of this bacterial suspension was used for DAS-ELISA according to the manufacturer’s instruction (Agdia, Elkhart, IN). EPS was quantified by measuring absorbance at 450 nm using a microtiter plate reader after adding 50 ll of 2M H2SO4 to each well to stop the reaction. The experiment was repeated three times, with three technical replicates in each experiment.

Plant growth conditions and disease assay Twenty-one-day-old tomato plants (cv. Bonny Best) grown in 4 inch pots at 288C with 12 h day/night cycle were inoculated by placing a droplet containing approximately 2000 cells of R. solanacearum on the cut petiole of the first true leaf of each tomato plant. Inoculum was prepared by growing bacteria overnight in CPG with appropriate antibiotics at 288C with agitation, collected by centrifugation (8000 r.p.m. for 5 min) and re-suspended in sterile water. Plants were rated daily for disease severity using a modified Horsfall-Barratt index scale: 0: no leaves wilted; 1: 1%–25% leaves wilted; 2: 26%–50% leaves wilted; 3: 51%–75% leaves wilted; 4: >75% leaves wilted (Tans-Kersten et al., 2001). The experiment was repeated three times, with 14 plants per treatment per biological replicate.

In planta translocation assay Bacterial translocation was measured as previously described (Addy et al., 2012). We inoculated 4-week-old Bonny Best tomato plants by placing 104 cells of GMI1000 WT-GFP or DnucA/B-GFP on a cut petiole. One week after inoculation, the stems were harvested and trimmed to remove all petioles. A sterilized razor blade was used to collect thin stem sections at 0, 2, 4 cm above and below the point of inoculation. Sections were observed under a Leica MZFLIII fluorescence dissecting microscope equipped with a GFP2 filter set (Leica Biosystems, Buffalo Grove, IL). To determine bacterial population sizes along tomato stem, 100 mg of stem tissue was harvested from each point sampled above, homogenized and serially dilution plated on CPG-TZC plates with antibiotics. Colonies were counted after 48 h incubation at 288C. The experiment was repeated twice with 10 plants per treatment.

Regulation of extracellular DNase production in R. solanacearum We used the DNase Alert assay (IDT, Coralville, IA) to measure extracellular DNase activity produced by mutants lacking genes for various virulence regulators (vsrAD, vsrBC, flhDC, phcA and hrpB (Genin et al., 1992; Schneider et al., 2009)). Strains were grown in BMM 1 2% glucose overnight and centrifuged at 6000 r.p.m. for 5 min, and culture supernatants were filter-sterilized to remove remaining cells. Eighty microliters of the cell-free supernatant was mixed with 10 ll of

DNase Alert substrate and 10 ll of DNase Alert buffer and seeded into a 96-well black microtiter plate (Corning, NY). Fluorescence signal from the cleaved DNA substrate was monitored with a plate reader for 6 h at 378C. Relative fluorescence units were standardized to cell numbers in the cultures, which were determined by serial dilution plating on solid CPG plates. The experiment was repeated three times, each with three technical replicates.

Acknowledgements This research was supported by grants from the Vietnam Education Foundation, the National Science Foundation and the University of Wisconsin-Madison College of Agricultural and Life Sciences. The authors gratefully acknowledge Sarah Swanson, Department of Botany, for technical advice on confocal microscopy. We also thank Beth Dalsing, Tiffany LowePower, Heidi Goodrich-Blair, Patricia McManus and Jeri Barak for critical review of the manuscript.

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Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site: Fig. S1. Loss of exDNases did not alter EPS1 production in R. solanacearum. Immunological quantification of EPS1 in R. solanacearum cultures was performed with ELISA using EPS1 antibody (A450). Ralstonia solanacearum strains were grown on CPG for 4 days at 288C and re-suspended in water to A600 5 1.0 (109 CFU ml21) and 10 ll of this bacterial suspension was taken for EPS1 quantification. Columns represent mean of three biological replicates, each with three technical replicates. Bars indicate standard errors. Only DepsB mutant (negative control) showed a significant different in EPS1 compared to wild-type (****P < 0.0001, oneway ANOVA). Fig. S2. COMSTAT analysis of 4-day-old biofilms formed by R. solanacearum wild-type GMI1000 and the extracellular nuclease-deficient DnucA/B mutant strain on glass chambered slides, as observed by confocal microscopy. Table S1. Bacterial strains, plasmids and strains used in this study. Table S2. Primers used in this study.
