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. 2012 Nov 27;14(3):256–264. doi: 10.1111/mpp.12004

Identification of a response regulator involved in surface attachment, cell–cell aggregation, exopolysaccharide production and virulence in the plant pathogen Xylella fastidiosa

Tanja M Voegel 1,4, Harshavardhan Doddapaneni 2, Davis W Cheng 2, Hong Lin 2, Drake C Stenger 2, Bruce C Kirkpatrick 1, M Caroline Roper 3,✉
PMCID: PMC6638743  PMID: 23186359

Summary

Xylella fastidiosa, the causal agent of Pierce's disease of grapevine, possesses several two‐component signal transduction systems that allow the bacterium to sense and respond to changes in its environment. Signals are perceived by sensor kinases that autophosphorylate and transfer the phosphate to response regulators (RRs), which direct an output response, usually by acting as transcriptional regulators. In the X. fastidiosa genome, 19 RRs were found. A site‐directed knockout mutant in one unusual RR, designated XhpT, composed of a receiver domain and a histidine phosphotransferase output domain, was constructed. The resulting mutant strain was analysed for changes in phenotypic traits related to biofilm formation and gene expression using microarray analysis. We found that the xhpT mutant was altered in surface attachment, cell–cell aggregation, exopolysaccharide (EPS) production and virulence in grapevine. In addition, this mutant had an altered transcriptional profile when compared with wild‐type X. fastidiosa in genes for several biofilm‐related traits, such as EPS production and haemagglutinin adhesins.

Introduction

The plant pathogenic, Gram‐negative bacterium Xylella fastidiosa is the causal agent of Pierce's disease (PD) of grapevine and many other economically important diseases, such as almond leaf scorch, citrus variegated chlorosis and oleander leaf scorch (Hopkins, 1989; Hopkins and Purcell, 2002; Purcell and Hopkins, 1996). The bacteria are transmitted in nature by xylem‐feeding insects, mainly sharpshooters (Hewitt et al., 1946). Once introduced into the plant xylem, X. fastidiosa forms biofilms that are thought to contribute, together with tyloses of host origin, to the occlusion of the xylem vessels, which may result in host plant death (Hopkins, 1989; Thorne et al., 2006). During X. fastidiosa biofilm formation, the cells primarily attach to the surface via large afimbrial adhesins and pili. The cells aggregate via haemagglutinins (HAs) and pili, which leads to the formation of microcolonies that subsequently form a mature matrix‐encased biofilm in the xylem vessels (Feil et al., 2007; De La Fuente et al., 2008; Guilhabert and Kirkpatrick, 2005; Li et al., 2007; Meng et al., 2005; Voegel et al., 2010). The X. fastidiosa biofilm matrix has been shown to contain exopolysaccharide (EPS), DNA and proteins (Cheng et al., 2010; Roper et al., 2007b).

Similar to other bacteria, X. fastidiosa has evolved to cope with changes in its physical and biological environment. The ability to rapidly sense and respond to its environment is particularly pertinent when X. fastidiosa makes the abrupt transition from life inside the insect foregut to life inside the plant xylem tissue, and vice versa. Bacteria utilize two‐component signal transduction (TCST) systems to sense and respond to their environment (Heeb and Haas, 2001; West and Stock, 2001). The prototypical TCST system is composed of two proteins, a sensor kinase and a cognate response regulator (RR). Diverse abiotic (pH, temperature, osmolarity) and biotic (host‐produced or quorum‐sensing) signals are typically perceived by the sensor domain of histidine protein kinases (HKs) leading to autophosphorylation of the HK. The phosphoryl group is then transferred to the RR, which modulates the expression of downstream genes, including genes important for biofilm formation and virulence (Heeb and Haas, 2001; West and Stock, 2001). RR proteins typically contain a conserved N‐terminal CheY‐like receiver domain (REC) that can function either as a stand‐alone module (Armitage, 1999; Szurmant and Ordal, 2004) or can be C‐terminally associated with a DNA‐binding (Stock et al., 2000; West and Stock, 2001), RNA‐binding (Shu and Zhulin, 2002), enzymatic output (Galperin, 2006) or protein‐binding (Rosario et al., 1994) domain.

In silico analysis of the X. fastidiosa genome identified a gene, designated xhpT, which encodes an RR containing a prototypical CheY‐like REC domain and a histidine phosphotransferase (HPt) protein‐binding domain. This is an unusual domain organization for bacterial RRs in general, with an occurrence of less than 0.1% (mostly found in proteobacteria), and is the only one of its type found in the X. fastidiosa genome (Galperin, 2006, 2010). Interestingly, gene neighbourhood analysis suggested that xhpT participates in the regulation of EPS synthesis, because of its location directly upstream of the gum operon, which encodes proteins putatively involved in the biosynthesis of the EPS polymer, fastidian gum (da Silva et al., 2001). This polymer is a major component of the X. fastidiosa biofilm matrix in vitro and in planta (Roper et al., 2007b). This observation prompted us to investigate the role of XhpT as a regulator of EPS synthesis, as well as of phenotypes associated with other steps involved in biofilm formation. In addition, microarray analyses on an xhpT null mutant were conducted. The results further elucidated the contribution of this unusual RR to the biology and pathogenicity of X. fastidiosa.

Results

Protein domain analysis of XhpT

The 714‐bp gene coding for PD1386, designated XhpT, for Xylella histidine‐containing phosphotransferase, is located immediately upstream of, but transcribed in the opposite orientation to, the gum operon that encodes genes putatively involved in EPS production (da Silva et al., 2001; Simpson et al., 2000; Van Sluys et al., 2003). XhpT is most likely a nonsecretory cytoplasmic protein as analysed by SMART (simple modular architecture research tool). Located at the C‐terminus (amino acids 10–118) is a REC superfamily domain, and located at the N‐terminus (amino acids 161–235) is an HPt (histidine‐containing phosphotransferase) domain.

The xhpT mutant is altered in cell–cell aggregation and surface attachment

The xhpT null mutant formed fewer bacterial cell aggregates (Fig. 1A) and was compromised in attachment to a solid surface (Fig. 1B) after 9 days of incubation in liquid culture when compared with wild‐type (WT) cells. Both strains reached stationary phase by 9 days of incubation, which was the time point at which all phenotypes and gene expression profiles were measured.

Figure 1.

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(A) Cell–cell aggregation assay of Xylella fastidiosa wild‐type (WT) Fetzer and xhpT mutant. The mutant forms fewer cell aggregates relative to WT cells. (B) Surface attachment of X. fastidiosa WT Fetzer and xhpT. The mutant attaches less well to polystyrene surfaces relative to WT X . fastidiosa. Error bars indicate standard deviation. Letters indicate groups assigned by Tukey's honestly significant difference test. OD, optical density.

The xhpT mutant produces more EPS matrix

Protein A double‐antibody‐sandwich enzyme‐linked immunosorbent assay (ELISA) using X. fastidiosa‐specific EPS antibodies was employed to quantify EPS associated with X. fastidiosa cells grown on PD3 plates (Roper et al., 2007b). The xhpT mutant produced significantly more EPS relative to WT X. fastidiosa when cells were assayed at a concentration of 108 colony‐forming units (cfu)/mL (Fig. 2).

Figure 2.

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Quantification of exopolysaccharide (EPS) production in Xylella fastidiosa wild‐type (WT) Fetzer and xhpT mutant using protein A double‐antibody‐sandwich enzyme‐linked immunosorbent assay (ELISA). The mutant produces more EPS relative to WT cells. Error bars indicate standard deviation. Letters indicate groups assigned by Tukey's honestly significant difference test.

Virulence and host colonization studies

PD symptom onset began at 8 weeks post‐inoculation in grapevines inoculated with either xhpT or WT X. fastidiosa. We scored plants at 18 weeks post‐inoculation on the arbitrary disease scale and observed that more WT‐inoculated plants reached a rating of ‘5’ (dead or dying) relative to plants inoculated with the xhpT mutant strain, meaning that more plants inoculated with the xhpT mutant strain exhibited fewer PD symptoms at this time point and were still alive. Specifically, the survival rate of plants inoculated with WT was only 60% ± 8.94% at 18 weeks post‐inoculation, whereas the survival rate was 90% ± 5.48% for plants inoculated with the xhpT mutant (Fig. 3), indicating that a mutation in xhpT resulted in a reduction in overall virulence.

Figure 3.

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Survival rates of Vitis vinifera var. Thompson seedless grapevines inoculated with Xylella fastidiosa wild‐type (WT) Fetzer and xhpT mutant strain, 18 weeks after inoculation. Plants inoculated with the xhpT mutant showed an increased survival rate relative to plants inoculated with WT.

Twelve weeks after inoculation, bacteria were quantified from infected petioles. There was no significant difference in the bacterial population between the two strains at the point of inoculation (P = 0.5939) or 25 cm above the point of inoculation (P = 0.4725), indicating that the mutant was capable of colonizing the plants to WT levels. Therefore, the decrease in virulence cannot be attributed to a colonization defect, indicating that XhpT may be involved in the regulation of virulence factors that specifically affect PD symptom development. No bacteria were recovered from plants inoculated with buffer alone.

Microarray and quantitative reverse transcription‐polymerase chain reaction (qRT‐PCR) analyses

To test the effects of a mutation in xhpT at the transcriptional level, microarray analysis was performed. Overall, 520 genes were at least two‐fold up‐regulated and 115 genes were at least two‐fold down‐regulated in the xhpT mutant relative to WT, as shown in Tables S1 and S2 (see Supporting Information). Select genes with differences in expression levels were organized into functional groups and are listed in Table 1 (genes that were up‐regulated) and Table 2 (genes that were down‐regulated).

Table 1.

Genes that are at least two‐fold up‐regulated in xhpT, organized into functional groups. A complete list can be found in Table S1 (see Supporting Information)

Gene Gene product Fold change
EPS production
PD1388/gumK GumK protein 3.79
PD1391/gumH GumH protein 2.66
PD1395/gumC GumC protein 3.4
PD1396/gumB GumB protein 3.59
Attachment/aggregation/motility
PD0060/fimD Outer membrane usher protein precursor 2.13
PD0731/xadA Outer membrane protein‐XadA 2.9
PD1928/pilR TCST, regulatory protein 3.1
PD1924/pilA Fimbrial protein 2.45
PD1148/pilU Twitching motility protein 2.43
PD1691/pilQ Fimbrial assembly protein 2.41
PD1147/pilT Twitching motility protein 2.91
Gene regulation
PD0092/lexA LexA repressor 2.16
PD0595/yybA Transcriptional regulator/MarR family 5.07
PD0708 Virulence regulator 3.62
PD0945/exsB Transcriptional regulator 2.73
PD1153/algR TCST, regulatory protein 2.16
PD1276/algH Transcriptional regulator 2.11
PD1367/colR TCST, regulatory protein 9.55
PD1905/xrvA Virulence regulator 2.7
PD1909 Transcriptional regulator 3.1
PD1919/colR TCST, regulatory protein 2.49
PD2050 TCST, hybrid sensor and regulatory protein 2.35
Potential virulence factors
PD0215/cvaC Colicin V precursor 8.72
PD0499/cvaB Colicin V secretion 2.25
PD0529/guxA Cellulose, 1,4‐β‐cellobiosidase 2.34
PD0708 Virulence regulator 3.6
PD0855/virK VirK protein 7.05
PD1341/vapI Virulence‐associated protein 3.73
PD1640/bglX β‐Glucosidase 2.24
PD1702 Hyp protein w/lipase domain 8.41
PD1703 Hyp protein w/lipase domain 33.47
PD1905/xrvA Virulence regulator 2.7
PD1964/tolC Outer membrane export factor 6.01
PD2061/egl Endo‐1,4‐β‐glucanase 3.47
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EPS, exopolysaccharide; TCST, two‐component signal transduction.

Table 2.

Genes that are at least two‐fold down‐regulated in xhpT, organized into functional groups. A complete list can be found in Table S2 (see Supporting Information)

Gene Gene product Fold change
Cell–cell aggregation
PD1792/hxfB Haemagglutinin‐like protein 0.35
PD2118/hxfA Haemagglutinin‐like protein 0.3
Attachment/aggregation/motility
PD0024/pilE PilE protein 0.42
PD00845/pilG Pilus protein 0.43
PD1926 Fimbrial assembly protein 0.25
PD1929/pilS TCST, sensor protein 0.49
Gene regulation
PD0279 GGDEF family protein 0.47
Potential virulence factors
PD0310 Pathogenicity‐related protein 0.44
PD1412/hlyB Toxin secretion 0.42
PD1439/mviN Virulence factor 0.46
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TCST, two‐component signal transduction.

qRT‐PCR was used to validate differences in gene regulation found by microarray analysis for PD0146, PD0215 (cvaC), PD0956, PD1299, PD1367 (colR), PD1391 (gumH), PD1395 (gumC), PD1396 (gumB), PD1702, PD1703, PD1711 (tonB) and PD3039, and the down‐regulation of PD1792 (hxfB) and PD2118 (hxfA). The qRT‐PCR data for all genes listed above followed the same trend as the microarray data, thus confirming the robustness of the microarray data.

Discussion

RRs are part of TCST systems and act as phosphorylation‐regulated switches coupling microbial responses to environmental cues. With an increasing number of annotated bacterial genomes, new RRs that possess atypical domain architectures have emerged (for an overview of domain architectures, see Galperin, 2006, 2010). Among the different possibilities for output domains, less than 0.1% of RRs from a nonredundant set of completely sequenced bacterial genomes possess an HPt domain (Galperin, 2010) and, in X. fastidiosa, XhpT is the only protein that has been identified with a REC–Hpt domain architecture. In this study, we sought to characterize the role of the XhpT RR based on genetic linkage to the gum operon and its unique domain structure within the X. fastidiosa genome.

The simplest and most common example of a signal transduction pathway is a TCST system composed of a sensor HK and its cognate RR. Because XhpT does not possess an obvious DNA‐binding domain, direct action as a transcriptional regulator is unlikely. Instead, XhpT may serve as a ‘signal integration locus’ within a multistep signal transduction network (e.g. a phosphorelay). This more complex type of two‐component phosphotransfer scheme involves several proteins, rather than just a sensor kinase and its cognate RR. The phosphoryl group is commonly transferred from a hybrid kinase via a histidine‐containing phosphotransfer protein (HPt) to the DNA‐binding RR (West and Stock, 2001). In X. fastidiosa, we speculate that XhpT transfers phosphate to the receiver domain of an unidentified downstream RR that acts as a transcription factor, initiating a change in gene expression (Goulian, 2010). Twelve genes listed in Tables 1 and 2 that putatively encode proteins involved in various regulatory pathways were differentially expressed in the xhpT mutant, suggesting that there may be branched regulatory pathways in X. fastidiosa. Two general scenarios of branched pathways have been described in other bacteria (Laub and Goulian, 2007). The first is a ‘many‐to‐one’ pathway in which more than one HK is responsible for the phoshorylation of the same RR. The second is a ‘one‐to‐many’ pathway in which one HK phosphorylates several RRs, as exemplified in bacterial chemotaxis systems, where the HK CheA phosphorylates the two RRs CheY and CheB (Goulian, 2010; Kirby, 2009). The existence of 19 RRs and only nine HKs in the X. fastidiosa genome further suggests considerable branching among the regulatory networks in X. fastidiosa. This strategy could be advantageous for X. fastidiosa by allowing greater signalling precision within the pathways that control complex phenotypes, such as biofilm formation. Biofilm formation is a process that generally follows a developmental sequence beginning with initial attachment to a surface and microcolony formation, followed by EPS matrix production and biofilm maturation (O'Toole et al., 2000), and appropriate timing of the steps in biofilm formation is often critical for successful host colonization. The integration of XhpT in a phosphorelay controlling biofilm formation may also allow X. fastidiosa to integrate multiple environmental signals into this pathway, thereby providing robust regulation over the phenotypes involved in this complex microbial behaviour.

Both surface attachment and cell–cell aggregation are early steps in the process of biofilm formation. A visible reduction was observed in the ability of the xhpT mutant to attach to the side‐wall of a polystyrene tube and to form cell–cell aggregates when the mutant was grown in liquid medium. These observations, together with the microarray analysis results, prompted further study of the phenotypes involved in the different stages of biofilm formation, including surface attachment, cell–cell aggregation and EPS production. Indeed, there was a quantitative difference in the ability of the xhpT mutant to initially attach to a polystyrene surface and to form cell aggregates. A gene encoding a cell surface adhesin, XadA, was up‐regulated in the xhpT mutant (Table 1). In WT X. fastidiosa biofilms, very little XadA protein is evident in the early phase of biofilm formation, but, as the biofilm matures, XadA accumulates and displays a distinct temporally regulated spatial ‘island‐like’ distribution, suggesting that it plays a role in the maintenance of biofilm structure (Caserta et al., 2010). We speculate that the deregulated/increased expression of xadA probably leads to an improperly skewed distribution of the XadA protein within the biofilm, causing an overall defect in biofilm stability/formation. Furthermore, an increased EPS layer surrounding the cells might prevent secretion of the extracellular XadA protein, thereby rendering the cells unable to form a stable biofilm. HxfA and HxfB function as cell–cell adhesins in X. fastidiosa (Guilhabert and Kirkpatrick, 2005) and the decrease in expression of HxfA and HxfB in the xhpT mutant probably contributes to the marked decrease in cell–cell aggregation observed for the xhpT mutant.

EPS production is a primary component of the matrix during late biofilm formation and maturation in X. fastidiosa (Roper et al., 2007b). Interestingly, the xhpT gene is directly upstream from, but transcribed in the opposite direction to, the gum operon encoding genes that probably synthesize EPS (da Silva et al., 2001; Simpson et al., 2000; Van Sluys et al., 2003), suggesting that XhpT plays an important role in the regulation of EPS production. The up‐regulation of genes involved in EPS biosynthesis and export probably accounts for the increased EPS production phenotype observed for the xhpT mutant. Indeed, a marked increase in total EPS matrix production was observed in the xhpT mutant, indicating that XhpT acts as part of a regulatory pathway that represses EPS synthesis. The overproduction of EPS could further explain why the xhpT mutant was compromised in attachment to a solid surface, as well as cell–cell aggregation. In other bacterial systems, the constitutive or early production of EPS can prevent initial surface adhesion and disrupt overall biofilm formation (Koutsoudis et al., 2006). Deregulated expression of the EPS layer that coats the bacterial cell surface could prevent physical access of other X. fastidiosa adhesins and surface proteins, thereby disrupting surface attachment and aggregation.

Genes PD1702 and PD1703 were highly up‐regulated in the xhpT mutant with fold changes of 8.41 and 33.47, respectively. Both genes encode for conserved hypothetical proteins that possess a LIP domain potentially functioning as a lipase. In Xanthomonas oryzae pv. oryzae, the causal agent of bacterial leaf blight on rice, lipase proteins are potent inducers of host defence responses. This may also be the case for X. fastidiosa, which could account for the decreased virulence observed for the xhpT mutant (Jha et al., 2007). In addition, the increased layer of EPS could trap secreted lipases, preventing access to the plant cell wall.

Although iron is an essential nutrient and necessary for cell growth, an excess of free iron can cause the production of damaging oxygen radicals via the Fenton reaction, in which iron catalyses the breakdown of hydrogen peroxide to hydroxyl radicals which are damaging to cell components (Imlay, 2008; Zheng et al., 1999). A fine balance is established by various genetic regulatory systems to ensure that a sufficient amount of essential iron is available to the cell without having an excess (Zheng et al., 1999). Fur, a well‐studied repressor of iron uptake (Hantke, 2001), was down‐regulated in the xhpT mutant, but, at the same time, several genes encoding proteins involved in the detoxification of reactive oxygen species (ROS) were up‐regulated. We speculate that the xhpT mutant responds to this deregulated increase in iron uptake by overproducing ROS scavenging enzymes to compensate for the uncontrolled uptake of iron and subsequent formation of damaging ROS.

Interestingly, 12 genes that encode putative virulence factors were up‐regulated in the xhpT mutant, yet a decrease in virulence was observed. This strongly suggests that appropriate temporal expression of certain virulence factors is critical for normal progression of PD.

Although we did not elucidate the exact mechanism by which XhpT regulates gene expression in X. fastidiosa, we demonstrated that this protein plays a central role in controlling a variety of phenotypes affecting biofilm formation and virulence. These findings suggest that XhpT is a critical component of the regulatory hierarchy governing host invasion for this plant pathogen.

Experimental Procedures

Bacterial strains and growth conditions

Xylella fastidiosa was grown on solid PD3 medium (Davis et al., 1981) with appropriate antibiotics when needed (kanamycin at 5 μg/mL) at 28 °C for 9 days. Liquid cultures were grown by the inoculation of a bacterial suspension of 108 cfu/mL in a 1:100 dilution into liquid PD3 medium. Cultures were grown for 9 days at 28 °C with agitation at 100 rpm.

For the microarray gene expression study, X. fastidiosa WT Fetzer strain and the xhpT null mutant were first grown from −80 °C glycerol stock on PD3 agar medium at 28 °C for 14 days. Cells were harvested from plates and transferred into PD3 liquid medium (Davis et al., 1981). Both strains were adjusted to a cell concentration equal to 105 cfu/mL and plated onto solid PD3 agar medium and grown for another 8 days. Cells were then harvested from plates and immediately stored in liquid nitrogen. Each strain was cultured on nine 15‐cm plates from which three replicates were obtained by pooling three plates per replicate. All strains used in this study are described in Table 3.

Table 3.

Primers, plasmids and strains used in this study

Primers, plasmids and strains Sequences (5′–3′) and characteristics Source
Primer
1386for TCGCAGATTGCGTTCCAT This study
1386Srev TGGTGCTGATACTCTGTGTGTACT This study
kan‐2FP‐1 ACCTACAACAAAGCTCTCATCAACC Life Technologies
kan‐2RP‐1 GCAATGTAACATCAGAGATTTTGAG Life Technologies
 xhpTfor GACCTGCGGAACCACTGTAT This study
 xhpTrev TGTTGGTGCGTTGTTTGATT This study
 xhpTcDNAfor ACACCGGCGCTGGCACATAC This study
 xhpTcDNArev GCAGCTGGCTTGTAGGCGGT This study
Plasmids
pCR2.1‐TOPO TA‐cloning vector, ApR KmR lacZ, T7 Life Technologies
pCR2.1‐xhpT pCR2.1‐TOPO with xhpT PCR product, KmR This study
pUC18 pMB1 derivate, rep (pMB1), ApR, lacZ Roche
pUC18‐xhpT pUC18 with xhpT PCR product, ApR This study
pUC18‐xhpT:kan pUC18‐xhpT with kanamycin cassette, ApR, KmR This study
Strains
 X. fastidiosa Fetzer Wild‐type (WT) Hendson et al., 2001
 xhpT X. fastidiosa Fetzer PD1386::(EZ::TN<Kan‐2>Tnp) This study
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Generation of the xhpT mutant strain

PD1386 (xhpT) was PCR amplified from WT X. fastidiosa (Fetzer) genomic DNA with the primer pair 1386for/1386rev (Table 3). The resulting PCR product was cloned into pCR2.1‐TOPO (Life Technologies, Carlsbad, CA, USA) to generate pCR2.1‐xhpT. The xhpT insert was subcloned into pUC18, a suicide plasmid in X. fastidiosa, to generate pUC18‐xhpT. A kanamycin resistance gene was randomly inserted into pUC18‐xhpT using the EZ‐Tn5™ <KAN‐2> Insertion Kit (Epicentre, Madison, WI, USA) according to the manufacturer's protocol, generating pUC18‐xhpT::kan. The location of the kanamycin cassette insertion was confirmed by sequencing using the primers kan‐2RP‐1/kan‐2FP‐1 (Life Technologies). The pUC18‐xhpT::kan mutagenesis construct was electroporated into X. fastidiosa WT Fetzer as described previously (Guilhabert et al., 2001), creating strain xhpT. Primer pair xhpTfor/xhpTrev was used to confirm double cross‐over events in the transformants (Fig. 4A,B). In addition, Southern blot analysis was used to confirm single insertion of the kanamycin resistance gene cassette into the WT Fetzer genome as described previously (Guilhabert and Kirkpatrick, 2005) (data not shown).

Figure 4.

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(A) Primer binding and EZ‐Tn5™ <KAN‐2> cassette insertion sites in xhpT. (B) Confirmation of the integration of the EZ‐Tn5™ <KAN‐2> cassette into the wild‐type (WT) X ylella fastidiosa Fetzer chromosome by polymerase chain reaction (PCR). Lane 1, 1‐kb plus ladder; lanes 2 and 3, PCR products obtained using the primer pair xhpTfor and xhpTrev and DNA from WT Fetzer (lane 2) and xhpT (lane 3). (C) Reverse transcriptase‐PCR. Lane 1, 1‐kb plus ladder; lanes 2 and 3, PCR products obtained using the primer pair xhpTcDNAfor and xhpTcDNArev and cDNA from WT Fetzer (lane 2) and xhpT (lane 3).

RT‐PCR was carried out to confirm the absence of xhpT RNA transcript in the mutant strain (Fig. 4C). RNA was isolated and cDNA generated as described below (microarray analysis) from WT Fetzer and the xhpT mutant. The cDNA was employed as a template for a subsequent PCR using the primer pair xhpTcDNAfor/xhpTcDNArev. All plasmids and primer sequences used in this study are shown in Table 3.

Growth curve, cell–cell aggregation and surface attachment assays

For growth curve analysis, the strains were grown in 40 mL of PD3 medium. Cell growth was monitored daily for 9 days by dispersing the cells and measuring the turbidity as an optical density at 600 nm (OD600nm).

For cell–cell aggregation assays, X. fastidiosa cultures were incubated for 9 days in liquid PD3 medium in 15‐mL polypropylene tubes without shaking. The turbidity of the upper culture medium (ODs), composed mostly of dispersed cells, was measured using a spectrophotometer at 600 nm. The culture medium was returned to the original tube, the settled aggregate masses were dispersed by pipetting and the total cell culture was measured (ODt). The relative percentage of aggregated cells was estimated as follows: percentage of aggregated cells = (ODt − ODs)/ODt × 100 (Burdman et al., 2000).

For surface attachment assays, X. fastidiosa cultures were incubated for 9 days in liquid PD3 medium in 15‐mL polystyrene tubes in a vertical position without shaking. Attachment to the surface walls of the tubes was assessed by a crystal violet staining method (Espinosa‐Urgel et al., 2000; Leite et al., 2004). After the incubation period, the PD3 medium was discarded and a 0.1% (w/v) aqueous solution of crystal violet was added to each tube, allowed to incubate for 15 min and rinsed with deionized H2O. The remaining stain was eluted from the bacterial ring using ethanol. The absorbance of the ethanol–crystal violet solution was measured at 540 nm. All assays were performed in triplicate and each experiment was repeated twice.

Quantification of X. fastidiosa EPS in vitro

Protein A double‐antibody‐sandwich ELISA was used to detect EPS produced by X. fastidiosa cells grown on PD3 plates for 9 days as described previously (Roper et al., 2007b). Anti‐EPS F(ab)2 fragments were prepared from anti‐Xf EPS immunoglobulin G (IgG) (Roper et al., 2007b) and EPS was quantified using protein A–alkaline phosphatase conjugate. Each sample was replicated four times and the experiment was repeated twice.

Virulence assay and quantification of in planta bacterial populations

Pathogenicity assays using Vitis vinifera var. Thompson seedless grapevines grown in a glasshouse were performed as described previously (Guilhabert and Kirkpatrick, 2005; Hill and Purcell, 1997; Purcell et al., 1999; Roper et al., 2007a). Fifteen plants were used for inoculation of each strain. In brief, 20 μL of bacterial inoculum was mechanically inoculated into the stem at the base of the grapevine using a 20‐gauge syringe needle. Eight weeks post‐inoculation, symptoms were rated biweekly using a disease rating scale of 0–5, with ‘0’ being healthy and ‘5’ being dead or dying (Guilhabert and Kirkpatrick, 2005). In planta bacterial populations were quantified from petiole tissue. Petioles were harvested, surface sterilized, ground in 1 × phosphate‐buffered saline (PBS) and 20 μL of the ground tissue was plated onto PD3 medium. The plates were incubated at 28 °C for 9 days and the colonies were enumerated.

Microarray analysis

RNA isolation and cDNA synthesis

The RiboPure‐Bacteria Kit (Life Technologies) was used to isolate total RNA. RNA was purified and concentrated using an RNeasy Mini Kit (Qiagen, Germantown, MD, USA) with on‐column DNase treatment. Samples were processed to yield ∼20 μg of bacterial RNA. RNA samples were then dissolved in water and quantified on a fluorometer using RiboGreen reagent according to the manufacturer's instructions (Life Technologies). Double‐stranded cDNA was synthesized using a cDNA Synthesis Kit (Life Technologies), and cDNA samples were shipped to the Nimblegen hybridization service facility (Roche Nimblegen, Madison, WI, USA). Both RNA and cDNA integrity were evaluated on a Bioanalyzer 2100 (Agilent, Santa Clara, CA, USA).

Probe design

Probe design was carried out in collaboration with Nimblegen, Inc. based on the available genomes for four Xylella strains (9a5c, Ann1, Dixon, Temecula). To design 60‐mer oligos for all the chromosomal annotated genes, three levels of stringency were used.

Group I oligos

In this category, oligos were selected with a GC value of 45%–59%, with no more than five homopolymer stretches of As, Ts, Gs or Cs. The GC cut‐off was determined based on the analysis of the GC distribution of the 60‐mer probes on a scale of 0–100. For this, two strains of X. fastidiosa (Ann1 and 9a5c) were used and the GC range for the 60‐mers was plotted at 5% intervals to determine the number of probes per interval. GC averaged around 50%–60%. To ensure optimal hybridization specificity and to eliminate cross‐hybridization, these oligos were blast screened so that none matched more than 15 bp consecutively to any other location in the genome. This set had the largest number of oligos representative of the four strains (82.4%–89.7%).

Group II oligos

For this category, the blast screening was omitted (8.5%–12.5%).

Group III oligos

For this category, both blast and GC screening were omitted (1.6%–5.7%).

Next, for genes which had more than 10 oligos that passed the set criteria, the 10 oligos from the 3‐end of the gene were selected. For Xylella plasmids and NPT2 gene oligos, no blast or GC restrictions were placed.

Microarray analysis

To reduce experimentally and hybridization dye introduced biases, three biological replicates per experimental stage using single colour hybridization were carried out. Slides were scanned using a GenePix 4.0 scanner and associated software (Molecular Devices Corp., Sunnyvale, CA, USA), and data representing raw spot intensities were subjected to robust multi‐array average (RMA) (Irizarry et al., 2003). We used the RMA model of probe‐specific background correction to the perfect match probes. Next, these corrected probe values were subjected to quantile normalization, and a median polish method was applied to compute a single expression value for all the probes of a particular gene. Differentially expressed genes were identified using sam software to generate fold differences and q values with a delta value of 1.2. Only genes with at least a two‐fold change in expression value were selected for further analyses (Tusher et al., 2001).

Validation of microarray data

RNA was isolated as described above and cDNA was generated using the High Capacity RNA‐to‐cDNA Kit (Life Technologies). Primers were generated using Primerexpress software (Applied Biosystems). DnaQ (PD1217), coding for the epsilon chain of DNA‐polymerase III, was used as an endogenous control. PCRs were performed using SYBR Green PCR Master Mix (Applied Biosystems) employing an ABI PRISM 7000 Sequence Detector System (Applied Biosystems). The PCR thermal cycling conditions were as follows: an initial step at 50 °C for 2 min; 10 min at 95 °C; and 40 cycles, with one cycle consisting of 15 s at 95 °C and 1 min at 60 °C. Serial dilutions of WT Fetzer cDNA were used to calculate the standard curve for all primer pairs. The primer efficiency is E = 10(−1/slope), with the slope derived from the standard curve. Each sample was processed in duplicate on each plate, with two plates in total. The Pfaffl equation was used for relative quantification (Pfaffl, 2001): ratio = (E target)ΔCt,target(control‐treated)/(E ref)ΔCt,ref(control‐treated).

Supporting information

Table S1 Genes that are at least two‐fold up‐regulated in xhpT.

Click here for additional data file. (162.3KB, docx)

Table S2 Genes that are at least two‐fold down‐regulated in xhpT.

Click here for additional data file. (106.9KB, docx)

Acknowledgements

This research was supported by the California Department of Food and Agriculture Pierce's Disease/Glassy Winged Sharpshooter Research Program. The authors would like to thank Jerome Braun (Department of Statistics, University of California, Davis, CA, USA) for assistance with the statistical analysis.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Table S1 Genes that are at least two‐fold up‐regulated in xhpT.

Click here for additional data file. (162.3KB, docx)

Table S2 Genes that are at least two‐fold down‐regulated in xhpT.

Click here for additional data file. (106.9KB, docx)

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