
==== Front
Mol Plant Pathol
Mol Plant Pathol
10.1111/(ISSN)1364-3703
MPP
Molecular Plant Pathology
1464-6722
1364-3703
John Wiley and Sons Inc. Hoboken

10.1111/mpp.70001
MPP70001
MPP-OA-24-051.R2
Original Article
Original Article
VirB11, a traffic ATPase, mediated flagella assembly and type IV pilus morphogenesis to control the motility and virulence of Xanthomonas albilineans
Li et al.
Li Meilin 1
Xiong Liya 1
Chen Wenhan 1
Li YiSha 1
Khan Abdullah 1
Powell Charles A. 2
Chen Baoshan https://orcid.org/0000-0001-5866-7033
1
Zhang Muqing https://orcid.org/0000-0003-3138-3422
1 zmuqing@163.com

1 Guangxi Key Laboratory for Sugarcane Biology & State Key Laboratory of Conservation and Utilization for Subtropical Agri‐Biological Resources Guangxi University Nanning Guangxi China
2 IRREC‐IFAS, University Fort Pierce Florida USA
* Correspondence
Muqing Zhang, Guangxi Key Laboratory for Sugarcane Biology & State Key Laboratory of Conservation and Utilization for Subtropical Agri‐Biological Resources, Guangxi University, Nanning, Guangxi 530005, China.
Email: zmuqing@163.com

02 9 2024
9 2024
25 9 10.1111/mpp.v25.9 e7000129 7 2024
17 2 2024
13 8 2024
© 2024 The Author(s). Molecular Plant Pathology published by British Society for Plant Pathology and John Wiley & Sons Ltd.
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the terms of the http://creativecommons.org/licenses/by-nc-nd/4.0/ License, which permits use and distribution in any medium, provided the original work is properly cited, the use is non‐commercial and no modifications or adaptations are made.

Abstract

Xanthomonas albilineans (Xal) is a gram‐negative bacterial pathogen responsible for developing sugarcane leaf scald disease, which engenders significant economic losses within the sugarcane industry. In the current study, homologous recombination exchange was carried out to induce mutations within the virB/D4‐like type IV secretion system (T4SS) genes of Xal. The results revealed that the virB11‐deletion mutant (ΔvirB11) exhibited a loss in swimming and twitching motility. Application of transmission electron microscopy analysis further demonstrated that the ΔvirB11 failed to develop flagella formation and type IV pilus morphology and exhibited reduced swarming behaviour and virulence. However, these alterations had no discernible impact on bacterial growth. Comparative transcriptome analysis between the wild‐type Xal JG43 and the deletion‐mutant ΔvirB11 revealed 123 differentially expressed genes (DEGs), of which 28 and 10 DEGs were notably associated with flagellar assembly and chemotaxis, respectively. In light of these findings, we postulate that virB11 plays an indispensable role in regulating the processes related to motility and chemotaxis in Xal.

VirB11, a traffic ATPase from Xanthomonas albilineans, is crucial for motility and virulence. Its loss disrupts swimming, twitching, and swarming behaviour, reducing virulence by impairing flagellar assembly and chemotaxis.

flagella
motility
sugarcane leaf scald disease
VirB11
virulence
Xanthomonas albilineans
the Guangxi Science and Technology Major ProjectAD20207020 China Agricultural Research System 10.13039/501100012453 170109 source-schema-version-number2.0
cover-dateSeptember 2024
details-of-publishers-convertorConverter:WILEY_ML3GV2_TO_JATSPMC version:6.4.8 mode:remove_FC converted:03.09.2024
Li, M. , Xiong, L. , Chen, W. , Li, Y. , Khan, A. , Powell, C.A. et al. (2024) VirB11, a traffic ATPase, mediated flagella assembly and type IV pilus morphogenesis to control the motility and virulence of Xanthomonas albilineans . Molecular Plant Pathology, 25 , e70001. Available from: 10.1111/mpp.70001
==== Body
pmc1 INTRODUCTION

The bacterium Xanthomonas contains 30 species, most of which are plant pathogens. Xanthomonas albilineans (Xal) is a species of Xanthomonas that can infect sugarcane and cause sugarcane leaf scald diseases (Alvarez‐Martinez et al., 2021). Disease symptoms range from single, white, narrow, clear streaks to infected leaves that wither and die entirely, causing plant death and substantial economic losses. Xal is a gram‐negative bacterium with a size of 0.6–1.0 μm × 0.2–0.3 μm, possessing polar flagella and displaying a rod‐shaped pilin structure. It colonizes the xylem and exhibits optimal growth at temperatures ranging from 25°C to 28°C (Li, Li, et al., 2022). Xal lacks the gum gene cluster responsible for producing exopolysaccharides and adhesion (Li, Bao, et al., 2022; Pieretti et al., 2015).

The bacterial type IV secretion systems (T4SS) form a functionally diverse superfamily prevalent across various bacterial species (Costa et al., 2021). The T4SS facilitates the translocation of DNA and mono‐ and multimeric protein substrates across bacterial and eukaryotic cells. In gram‐negative bacteria, T4SS assembly progresses from VirB1 through VirB11 and VirD4 (Wu et al., 2014). Structural insights have been gained into several critical components of the T4SS, including the conserved ATPases (VirD4, VirB4, VirB11) and various channel/pilus subunits (VirB5, VirB8, VirB9, VirB10). These components are organized into integral inner membrane (IM) subunits (VirB3, VirB6, and VirB8), which probably form an IM channel, whereas outer membrane (OM)‐associated subunits (VirB7, VirB9, and VirB10) form a structural scaffold that facilitates the channel's passage through the periplasm and OM of Xanthomonas (Li et al., 2019; Sgro et al., 2019). VirB1, VirB2, and VirB5 are implicated as putative components of the T‐pilus, which probably mediates binding to plant cells and the subsequent transfer of genetic material (Alvarez‐Martinez et al., 2021; Li et al., 2019). The T4SS system in Xanthomonas citri is involved in producing effectors with lethal properties against other bacterial species (Souza et al., 2015). The T4SS‐mediated transport of plasmids and proteins plays essential roles in bacterial survival, evolution, virulence, and drug resistance (Alvarez‐Martinez & Christie, 2009; Atmakuri et al., 2004; Dunger et al., 2016; Mary et al., 2018; Sgro et al., 2019; Yuan et al., 2005).

VirB11 proteins are traffic ATPases that belong to the large AAA+ secretion protein superfamily, which includes proteins of type II secretion systems, type IV pilus biogenesis, and archaeal flagellar assembly machinery (Planet et al., 2001; Ripoll‐Rozada et al., 2013). These soluble, hexameric proteins are located on the cytoplasmic side of the inner membrane and characterized by distinct N‐terminal (NTD) and C‐terminal (CTD) domains connected by a flexible linker crucial for the catalytic cycle (Ripoll‐Rozada et al., 2012). VirB11, a conserved P‐ring ATPase, is an essential component located within the intermembrane region of the bacterial cell, containing Walker A and Walker B motifs and playing a crucial role in biogenesis processes (Christie, 2016; Christie et al., 2014; Sgro et al., 2019). VirB11 assists in the polymerization of pili by relocating pilus protein components from the inner membrane to the outer periplasmic region (Kerr & Christie, 2010). The flaHIJ gene, essential for flagellar assembly in all flagellated bacteria, shares homology with VirB11 and many other secretion‐associated ATPases (Thomas et al., 2002). Flagella, composed of a flagellar filament, a motor, and connecting components between the rod and the hook, significantly affect bacterial colonization, biofilm formation, motility, and virulence (Malamud et al., 2011; Suaste‐Olmos et al., 2010; Wadhwa & Berg, 2022; Zhu et al., 2021).

Bacterial motility includes various forms, such as flagellum‐mediated swimming and swarming and flagella‐independent surface twitching and gliding (Raina et al., 2019; Wadhwa & Berg, 2022). Bacterial swimming primarily occurs in liquid environments, allowing bacteria to explore their surroundings, seek nutrients or favourable conditions, and avoid harmful factors (Wadhwa & Berg, 2022). During bacterial swimming, individual bacteria move independently, relying on the action of flagella and modulating flagellar rotation to change direction or speed, as observed in 0.3% soft agar (Ha et al., 2014; Kuchma et al., 2015). On the other hand, bacterial swarming occurs on solid surfaces and involves the coordinated movement of bacteria as multicellular structures facilitated by flagella‐mediated motility and chemotaxis (Jeckel et al., 2019; Wadhwa & Berg, 2022). This rapid surface migration is preceded by a cell density‐dependent lag period, which can be eliminated if actively swarming cells are used as the inoculum (Kearns & Losick, 2003). The collective movement in swarming is notably aided by flagella and type IV pili on semisolid surfaces containing 0.6%–1% agar (Kearns, 2010). Twitching, a type of surface‐associated motility, is mediated not by flagella but by slender filaments called type IV pili (Craig & Forest, 2019). These pili play diverse roles in host adhesion (Craig et al., 2004), virulence (Craig et al., 2004; Persat et al., 2015), chemotaxis (Oliveira & Foster, 2016), and surface quorum‐sensing (QS) (Dufrêne & Persat, 2020). Twitching motility involves the type IV pili (T4P) cycle, which is characterized by the extension and attachment of pili to a solid substrate, followed by retraction that acts as an anchor, drawing the cell forward (Merz et al., 2000).

Bacterial chemotaxis, a behavioural trait of bacteria, plays a crucial role in adhesion, cell auto‐aggregation, and motility processes. It is essential for establishing symbiotic associations with plants and colonizing host organisms by pathogens (Huang et al., 2017; Johnson & Ottemann, 2018; Scharf et al., 2016; Wadhwa & Berg, 2022). Chemotaxis involves the active movement of bacteria towards or away from specific chemical gradients, enabling them to seek advantageous conditions or avoid harmful environments strategically.

This study aimed to elucidate the regulatory function of traffic ATPase of VirB11 in Xal through mutational and transcriptomic analysis using RNA sequencing. Our results revealed that the deletion of virB11 impaired the flagella formation, pilus morphogenesis, cell motility, and virulence in Xal. Transcriptome and reverse transcription‐quantitative PCR (RT‐qPCR) analyses further revealed differentially expressed genes (DEGs) involved in various functions, including flagellar assembly and chemotaxis in the ΔvirB11 mutant strain. Our results prove that VirB11 regulates flagellar assembly and pilus morphogenesis.

2 RESULTS

2.1 Identification of VirB11, a traffic ATPase of Xal and generation of virB11 ‐deletion mutant (ΔvirB11 )

Our comparative genomics suggests that the T4SS is required for the virulence of Xal JG43 (Li, Li, et al., 2022). This T4SS system comprises 12 protein complexes encoded by the VirB/VirD operon (VirB1–VirB11/VirD4; Figure 1). To elucidate the functional significance of T4SS in Xal JG43, we generated six knockout mutants (indicated by red arrows) using double homologous recombination to delete the complete VirB/VirD locus from the JG43 genome (Figure 1). A putative open reading frame (ORF) containing 351 amino acids was identified in the genome sequence of the Xal JG43 strain (GenBank accession number GCA_024158005.1). This ORF, located from nucleotide 953,496 to 954,551 with the locus tag GE000765, exhibits a typical domain structure of traffic ATPases belonging to the large AAA+ secretion protein superfamily, characterized by the presence of an N‐terminal domain (NTD) and a C‐terminal domain (CTD). A BLASTp search revealed that this ORF shares 100% amino acid sequence identity with the bacterial secretion protein VirB11 in Xal GPE PC73, which is annotated as a T4SS protein. This suggests that the ORF in the Xal JG43 strain is also the T4SS protein VirB11.

FIGURE 1 Schematic diagram of conservation of virB/virD operon genes among type IV secretion systems. The Xanthomonas albilineans virB/virD4 operon, with subunit functions and associations ATPase, pilus, outer membrane (OM) protein, and intermembrane (IM) protein; genes chosen for deletion are indicated by red arrows.

Confirmation of the virB11‐deleted mutant strain (∆virB11) was performed through PCR amplification using specific primers (virB11L‐F/virB11R‐R), followed by sequencing of the PCR product (Table 1). The resulting PCR product was 1093 bp, containing 519 bp upstream and 574 bp downstream fragments of the virB11 gene. In contrast, the wild‐type strain JG43 produced a PCR product of 2149 bp, which included the full length of the 1056 bp virB11 gene and the same flanking fragments (Figure S1a–c). Similar confirmation was carried out for other genes (virB4, virB5, virB8, virB10, virD4) identified in the genome sequence of the Xal JG43 strain and the corresponding mutant strains (∆virB4, ∆virB5, ∆virB8, ∆virB10, ∆virD4) (Table 2; Figure 1).

TABLE 1 Primers used for construction of mutant and complemented strains.

Primers	Sequence (5–3′, restriction enzyme sites are underlined)	Restriction enzyme	Product of PCR (bp)	Description	
virB11‐L‐F	CGCGGATCC TATATTCCATCAACGGCAGACG	BamHI	519	Mutant of virB11	
virB11‐L‐R	TGCTCTAGA CATCAGATTCCTCAGGCGGTCA	XbaI	
virB11‐R‐F	TGCTCTAGA CGCTATATCACCGGTATCGATT	XbaI	574	
virB11‐R‐R	CCCAAGCTT AATTCCCGGAATAGTAGCAACT	HindIII	
virB11‐F	AGCGAATGGAAATACCGTCGTT	–	515	Inner primer of virB11	
virB11‐R	CGAGTAAAGCAAATGTACCGCA	–	
CMΔvirB11‐L	CGCGGATCCTATATTCCATCAACGGCAGACGC	BamHI	1574	Complementary primer ofvirB11	
CMΔvirB11‐R	CCCAAGCTTTCAATGAAAATCACTCATCCCGC	HindIII	
virB5‐L‐F	CGCGGATCCCACGGACGTCAACTTTGGTTCAG	BamHI	529	Mutant of virB5	
virB5‐L‐R	TGCTCTAGATCAGGAAGACCGAGCAAGCAACG	XbaI	
virB5‐R‐F	TGCTCTAGATGCAGGAACAACGGCGACGCGAA	XbaI	544	
virB5‐R‐R	CCCAAGCTT CGTGGCGAGCTTTGTGTTGCTGC	HindIII	
virB5‐L	AGATGACCAGCCAAATCAACCAG	–	576	Inner primer of virB5	
virB5‐R	AGATGACCAGCCAAATCAACCAG	–	
virB8‐L‐F	CGCGGATCC ATATATTTCGGCTGATCGCTAC	BamHI	547	Mutant of virB8	
virB8‐L‐R	TGCTCTAGA TGTTAACGCTCTCGCTCTCAAT	XbaI	
virB8‐R‐F	TGCTCTAGA TGGTAGTGCACCTGTTTATCCG	XbaI	570	
virB8‐R‐R	CCCAAGCTT ATCCAGCTGCGACTCATCCTTC	HindIII	
virB8‐L	GTGGGTCGCGTTCAGTGCAATT	–	520	Inner primer of virB8	
virB8‐R	ACGAAAACCCAGGATAGCGAAT	–	
virB10‐L‐F	CGCGGATCCATCCGGACTGCCACCCATTCCTATA	BamHI	555	Mutant of virB10	
virB10‐L‐R	TGCTCTAGA AACGCGGCTGACCCTGATCATG	XbaI	
virB10‐R‐F	TGCTCTAGA CACCGTGATCAATGTGTATGTTG	XbaI	568	
virB10‐R‐R	CCCAAGCTT CCTCAATTCAAGCAACTCAAGA	HindIII	
virB10‐L	GATTTTCTGGATCGTCGGTGGC	–	555	Inner primer of virB10	
virB10‐R	CCTGTCGGTTCTACTGCACCAT	–	
virB4‐L‐F	CGCGGATCC AGTATGCATAAGAACGTACTT	BamHI	524	Mutant of virB4	
virB4‐L‐R	TGCTCTAGA ATCGAAGTATCTGGGATGAACA	XbaI	
virB4‐R‐F	TGCTCTAGATGGTTGAGCGAATTCTATGCAAGGC	XbaI	576	
virB4‐R‐R	CCCAAGCTT CGAACTGATGGCGCTGATGGAG	HindIII	
virB4‐L	ATGGTGTGGTTTTCTCCGAAGC	–	505	Inner primer of virB4	
virB4‐R	GACAGTTCTGCACGCGTTGTGG	–	
virD4‐L‐F	CGCGGATCC ATCCGAGTCATGCGCTGTTCGAA	BamHI	550	Mutant of virD4	
virD4‐L‐R	TGCTCTAGA CGCGATGACGGCGGTAATGCCTT	XbaI	
virD4‐R‐F	TGCTCTAGAGCGGAGCGTAATCGCATGAGATTTT	XbaI	581	
virD4‐R‐R	CGCAAGCTT CCCAGATTGTTGTCGAAAACGGT	HindIII	
virD4‐L	GCATCCGTATGCGCCGAAAATCA	–	500	Inner primer of virD4	
virD4‐R	TCCTTGTACAGCGTGGAGGCGAT	–	

TABLE 2 Bacterial strains and plasmids used in the present study.

Strains or plasmids	Relevant characteristics	Reference	
Xanthomonas albilineans	
JG43	Wild type; Rifr	In this study	
ΔvirB11	virB11 in‐frame deletion mutant of JG43; Rifr	In this study	
CMΔvirB11	ΔvirB11 harbouring plasmid pL/virB11; Rifr; Tcr	In this study	
ΔvirB4	virB4 in‐frame deletion mutant of JG43; Rifr	In this study	
ΔvirB5	virB5 in‐frame deletion mutant of JG43; Rifr	In this study	
ΔvirB8	virB8 in‐frame deletion mutant of JG43; Rifr	In this study	
ΔvirB10	virB10 in‐frame deletion mutant of JG43; Rifr	In this study	
ΔvirD4	virD4 in‐frame deletion mutant of JG43; Rifr	In this study	
PΔvirB11	ΔvirB11 harbouring plasmid pLARF6; Rifr; Tcr	In this study	
Plasmids	
pLAFR6	Broad host range IncP cloning cosmid; Tcr	Jiang et al. (2018)	
pK18mobsacB	Suicide plasmid; Mob+ Tra−; Kanr	Guan et al. (2020)	
Escherichia coli	
DH5α	supE44 ΔlacU169 (Φ80lacZ ΔM15) hsdR17 recA1 endA1 gyrA96 thi‐1 relA1	Hanahan (1983)	
pK/virB11	pK18mobsacB containing the sequenced upstream and downstream of virB11, Kanr	In this study	
pK/virB4	pK18mobsacB containing the sequenced upstream and downstream of virB4, Kanr	In this study	
pK/virB5	pK18mobsacB containing the sequenced upstream and downstream of virB5, Kanr	In this study	
pK/virB8	pK18mobsacB containing the sequenced upstream and downstream of virB8, Kanr	In this study	
pK/virB10	pK18mobsacB containing the sequenced upstream and downstream of virB10, Kanr	In this study	
pK/virD4	pK18mobsacB containing the sequenced upstream and downstream of virD4, Kanr	In this study	
pL/virB11	pLAFR6 containing the sequenced whole ORF of virB11; Tcr	In this study	
Abbreviations: Kanr, kanamycin resistance; ORF, open reading frame; Rif,r, rifampicin resistance; Tcr, tetracycline resistance.

2.2 Virulence reduced in the virB11 deletion mutant of Xal

To investigate the impact of six deletion mutants (∆virB11, ∆virB4, ∆virB5, ∆virB8, ∆virB10, ∆virD4) on the virulence of Xal, we compared their disease index (DI) in the sugarcane susceptible cultivar GT46 with that of the wild‐type JG43 (Table 2). The DI induced by the wild‐type JG43 was 54.76. In contrast, the mutant strain (∆virB11) and its vector control (P∆virB11) showed significantly reduced DI values of 13.45 and 14.79, respectively (p < 0.01). The complemented strain CM∆virB11 partially restored the DI to 39.23 (Figure 2a,b). However, the DI values for the mutants ∆virB4, ∆virB5, ∆virB8, ∆virB10, and ∆virD4 were not significantly different from that of the wild‐type JG43 (Figure 2a). Growth analysis, determined by the optical density (OD600) of cell suspensions, showed that the growth rates of the deletion mutants (∆virB4, ∆virB5, ∆virB8, ∆virB10, ∆virD4, and ∆virB11) were comparable to that of the wild‐type JG43 (Figure 2c).

FIGURE 2 Disease index and growth ability were measured in the wild type (JG43) and its mutant strains (ΔvirB4, ΔvirB5, ΔvirB8, ΔvirB10, ΔvirB11, ΔvirD4). (a) Disease index was measured in the wild type JG43, the deleted mutants of virB4 (ΔvirB4), virB5 (ΔvirB5), virB8 (ΔvirB8), virB10 (ΔvirB10), virB11 (ΔvirB11), virD4 (ΔvirD4), respectively, ΔvirB11 carrying either the empty vector pLAFR6 as vector control (PΔvirB11) or the recombinant vector pLvirB11, which had the full‐length virB11 gene cloned into pLAFR6 for complementation (CMΔvirB11). The bars represent standard errors of the means from three experiments. Each experiment contained three replicates for each strain. Statistical analysis was done using a t test, and statistical significance was denoted by **p < 0.01; ns, not significant (p > 0.05). (b) Appearance of representative sugarcane leaves inoculated with sterile water, the wild‐type strain of JG43, ∆virB11, P∆virB11, and complemented strain CM∆virB11 lesion areas of X. albilineans. Each strain (OD600 = 0.5) was inoculated with 15 sugarcane plants, three biological replicates were prepared of each strain. (c) Growth curve profiles of the wild type JG43 and its mutants in MW medium, optical density (OD600) of the cultured cell suspensions of the tested strains measured at time points of 0–114 h. The error bars represent the standard errors of the means from three experiments.

Given that ∆virB11 exhibited reduced virulence in Xal, we investigated its potential regulatory role through comparative transcriptome analysis of the virB11‐deleted mutant strain (∆virB11) and its wild‐type counterpart JG43. The results revealed 123 differentially expressed genes (DEGs), with 52 genes upregulated and 71 genes downregulated in the ∆virB11 strain (Table S1). The KEGG analysis identified ‘flagella assembly’ and ‘cellular chemotaxis’ as predominant terms (Figure 3).

FIGURE 3 KEGG pathway enrichment of ΔvirB11. KEGG pathway analysis showed significant enrichment in flagellar assembly and chemotaxis. KEGG enrichment was assessed using the Rich factor, false discovery rate (FDR), and the number of enriched genes. The Rich factor, the ratio of enriched to annotated differential genes, indicates enrichment degree, with higher values signifying more significant enrichment. FDR ranges from 0 to 1, with values nearer to zero denoting more significant enrichment.

2.3 virB11 deletion mutant of Xal regulates the flagellar assembly to cause swimming losses

Here, we present compelling evidence demonstrating the role of the virB11 gene in regulating flagellum‐dependent swimming motility. Our swimming motility assay showed that the mutant strain ∆virB11 failed to spread 4 days after inoculating a 10 μL bacterial suspension onto the centre of a soft agar plate (0.3% agar) at 28°C. In contrast, the wild‐type strain JG43 exhibited swimming with an average colony diameter of 1.93 cm, significantly larger than the ∆virB11 of 0.27 cm (p < 0.0001) and the P∆virB11 of 0.28 cm (p < 0.0001), but similar to the CM∆virB11 of 1.61 cm. This confirmed that ∆virB11 had lost its swimming ability. The other five mutants (∆virB4, ∆virB5, ∆virB8, ∆virB10, ∆virD4) showed swimming ability consistent with the wild‐type strain JG43 (Figure 4a,b). Furthermore, to determine the underlying cause of the observed loss of swimming motility, we conducted a comparative analysis of flagellar formation among Xal strains. Transmission electron microscopy (TEM) analysis confirmed the loss of flagella in the virB11‐deleted mutant (∆virB11), while flagella were restored in the complemented strain (CMΔvirB11) (Figure 5).

FIGURE 4 Swimming motility of JG43 and its mutant strains. (a)The swimming assay was conducted on JG43 and its mutants (ΔvirB4, ΔvirB5, ΔvirB8, ΔvirB10, ΔvirB11, ΔvirD4, virB11, complemented strains CMΔvirB11 and vector control strain PΔvirB11) for 4 days after inoculation of 10 μL each bacterial suspension to the centre of a medium with 0.3% specific agar (Aladdin) plate at 28°C. Experiments were repeated three times independently with similar results. (b) Swimming motility measured by colony diameters of each strain on swimming medium plate. The bars represent standard errors of the means from three experiments, and each experiment contained three replicates for each strain. Statistical analysis was done using a t test, and statistical significance was denoted by ****p < 0.0001; ns, not significant (p > 0.05).

FIGURE 5 The formation of polar flagellum under transmission electron microscopy of JG43, ΔvirB11, and the complemented CMΔvirB11 strains. Filament polar flagella (indicated by red arrows) were observed in the wild‐type strain JG43 and complemented strain CMΔvirB11 but not in the mutant strain ΔvirB11.

The bacterial flagellum is an extracellular tubular protein structure that acts as a propeller, enabling bacterial swimming motility. Compared to the wild‐type JG43, 28 DEGs were identified as involved in flagellar assembly, including 24 upregulated genes and 4 downregulated genes in the ∆virB11 (Table 3). The ∆virB11 regulon contained DEGs involved in flagellar assembly, encompassing components such as the filament, rod, hook, MS‐ring, C‐ring, stator unit, L‐ring, biosynthesis proteins, motor switch, flagellar chaperone, and assembly protein (Table 3; Figure 6a). RT‐qPCR analysis of 16 selected genes confirmed these findings, showing a strong positive correlation (R = 0.918, p < 0.0001) between the RT‐qPCR results and the transcriptome data (Figure 6b,c). The relative expression of 10 flagellar genes was measured in the wild‐type JG43 and the mutant‐complemented strain CM∆virB11. In CM∆virB11, the motA gene was downregulated, while five flagellar genes (flgH, flgK, flgF, flgC, fliL) were significantly upregulated, with fliL showing the highest expression, consistent with the ∆virB11 expression outcomes. Additionally, the relative expression of the flagellar biosynthesis genes flhB and flhF, the flagellar assembly gene fliH, the flagellar motor switch gene fliM, and the flagellin gene fliC in CM∆virB11 were similar to those in the wild‐type JG43 (Figure 6d).

TABLE 3 Differentially expressed genes involved in flagellar assembly in Xanthomonas albilineans mutant strain ΔvirB11 compared to its wild‐type strain JG43.

Gene ID	Gene	Gene function	Log2FC	p‐value	
GE002733	motA	Flagellar motor protein MotA	−1.12	0.008	
GE001210	motC	Flagellar motor protein MotC	1.56	0.043	
GE001204	flhF	Flagellar biosynthesis protein FlhF	1.65	0.033	
GE001203	flhA	Flagellar biosynthesis protein FlhA	2.94	0.001	
GE001202	flhB	Flagellar biosynthesis protein FlhB	1.47	0.025	
GE001198	fliO	Flagellar biosynthesis protein FliO	1.57	0.019	
GE001196	fliM	Flagellar motor switch protein FliM	2.07	0.005	
GE001195	fliL	Flagellar basal body FliL	3.01	0.001	
GE001194	fliK	Flagellar hook‐length control protein FliK	1.48	0.005	
GE001193	fliJ	Flagellar export protein FliJ	1.29	0.003	
GE001192	fliI	Flagellar assembly protein FliI	1.78	0.006	
GE001191	fliH	Fagellar assembly protein FliH	1.45	0.020	
GE001190	fliG	Flagellar motor switch protein FliG	1.63	0.006	
GE001189	fliF	Flagellar M‐ring protein FliF	1.92	0.002	
GE001188	fliE	Flagellar hook‐basal body protein FliE	2.78	0.001	
GE001171	fliS	Flagellar export protein FliS	−1.02	0.012	
GE001168	fliC	Flagellar filament hook protein FliC	−3.25	0.001	
GE001167	flgL	Flagellar hook‐associated protein FlgL	1.07	0.012	
GE001166	flgK	Flagellar hook‐associated protein FlgK	1.99	0.005	
GE001165	flgJ	Flagellar assembly protein FlgJ	1.89	0.011	
GE001164	flgI	Flagellar basal body P‐ring protein FlgI	1.47	0.024	
GE001163	flgH	Flagellar basal body L‐ring protein FlgH	1.71	0.019	
GE001162	flgG	Flagellar basal body rod protein FlgG	1.8	0.031	
GE001161	flgF	Flagellar basal body rod protein FlgF	1.93	0.015	
GE001160	flgE	Flagellar hook protein FlgE	1.59	0.033	
GE001158	flgC	Flagellar basal body rod protein FlgC	2.5	0.005	
GE001157	flgB	Flagellar basal body rod protein FlgB	3.07	0.001	
GE001154	flgM	Flagellar biosynthesis protein FlgM	−1.21	0.007	
Note: Gene ID refers to the locus_tag of DEGs X. albilineans ΔvirB11 strain compared to strain JG43, identified by BLASTn search against the strain JGT43 genome (GenBank accession number GCA_024158005.1). FC, fold‐change, |log2‐fold change| was derived from the mutant versus the wild‐type strain, |log2 fold‐change| > 1, or p‐value < 0.05.

Abbreviation: DEGs, differentially expressed genes.

FIGURE 6 Differentially expressed genes (DEGs) involved in flagellar assembly in the regulons of ΔvirB11 and CMΔvirB11. (a) Schematic diagram of the DEGs involved in flagellar assembly, a flagellar assembly including flagellin (fliC), flagellar motor gene (motA), flagella L‐ring (flgH), and P‐ring (flgI), flagella biosynthesis (flhF, flhA, flhB, fliO, flgM), MS‐ring(fliF), flagella cap (fliD), hook (fliK, fliC, flgL, flgK, flgE) and basal‐body rod‐related genes (fliE, flgB, flgC, flgG, flgF). Flagellar chaperones (fliJ, fliS) assembly genes (fliH, fliI, flgJ) and flagellar export gene (fliS), upregulated genes are shaded blue, and down‐regulated genes are highlighted in red. (b) Flagellar DEGs with 16 selected genes (motA, flgM, fliS, fliC, flhF, flhA, flhB, flgH, flgK, flgC, flgG, flgF, fliL, fliG, fliM, fliE) reverse transcription‐quantitative PCR (RT‐qPCR) verification, all RT‐qPCRs were performed three or more times independently using primers listed in Table S2. (c) Flagellar DEGs RT‐qPCR analysis and transcriptome correlation analysis. (d) DEGs in flagellar genes (motA, flgH, flgK, flgF, flgC, fliL, flhB, flhF, fliH, fliM, fliC) involved in flagellar assembly in the regulons of wild‐type strain JG43 and complemented strain CMΔvirB11. Statistical analysis was done using a t test, and statistical significance was denoted by **p < 0.01, experiments were repeated three times independently results.

2.4 virB11 deletion mutant of Xal reduced the twitching ability resulting from the loss of pilus morphogenesis

Twitching motility is a type of solid surface translocation occurring in humid conditions on semisolid or solid surfaces. This process relies on retractile type IV pili and is not influenced by the presence of a flagellum (Merz et al., 2000; Wadhwa & Berg, 2022). The loss of the virB11 gene significantly impaired the twitching motility of the ∆virB11 strain, as evidenced by a notable decrease in bacterial colony diameter to only 0.2 cm compared to 2.0 cm for the wild‐type strain JG43 (p < 0.0001) at 4 days after inoculating 10 μL bacterial suspension onto the centre of an agar plate (1% agar) at 28°C. The mutants ∆virB11, P∆virB11 (0.21 cm), and CM∆virB11 (0.18 cm) exhibited a complete loss of twitching ability, remaining comparable to the mutant (p < 0.0001). Additionally, the twitching abilities of ∆virB4, ∆virB5, ∆virB8, ∆virB10, and ∆virD4 were consistent with the wild‐type strain JG43 (Figure 7a,b). Compared to the virB11‐deleted mutant (∆virB11), two DEGs (fimU and pilE) involved in type IV pilus (T4P) were upregulated in wild‐type JG43, which was further confirmed by RT‐qPCR analysis (Figure 7c). However, fimU and pilE genes were downregulated in the CMΔvirB11 compared to the wild‐type JG43 (Figure 7d).

FIGURE 7 Twitching motility, differentially expressed genes, and images of the pilus morphogenesis in the wild‐type JG43, ΔvirB11, and the complemented CMΔvirB11 strains obtained by transmission electron microscopy. (a)The twitching motility assay was conducted on JG43 and its mutants grown for 3 days after inoculating 10 μL each bacterial suspension to the centre of a medium with 1% agar plate in an inverted culture at 28°C. Experiments were repeated three times independently with similar results. (b) Twitching motility measured by colony diameters of each strain on twitching medium plate. (c) fimU and pilE genes upregulated in the regulons of ΔvirB11, all reverse transcription‐quantitative PCRs were performed three or more times independently using primers listed in Table S2. (d) fimU and pilE genes downregulated in CMΔvirB11. (e) Images of the pilus morphogenesis of JG43, ΔvirB11, and the CMΔvirB11 strains. The formation of pilus morphogenesis observed under transmission electron microscopy. Clear pilus morphogenesis was observed in the wild‐type strain JG43 but not in the mutant strain ΔvirB11 and complemented strain CMΔvirB11. The bars represent standard errors of the means from three experiments, and each experiment contained three replicates for each strain. Statistical analysis was done using a t test, and statistical significance was denoted by *p < 0.05, **p < 0.01, ****p < 0.0001; ns, not significant (p > 0.05).

To determine whether the loss of pilus morphogenesis contributed to the absence of twitching motility, we used TEM to examine the development of type IV pili in each tested bacterium, including the wild‐type JG43, its mutant strain ∆virB11, and the complemented strain CM∆virB11. The wild‐type JG43 displayed distinct pilus morphogenesis, whereas the mutant ∆virB11 and the complemented strain CM∆virB11 lacked such features (Figure 7e). These findings indicate that the deletion of the virB11 gene impairs type IV pilus formation, leading to the loss of twitching motility in the Xal strain.

2.5 VirB11 deletion mutant of Xal mediated the chemotaxis‐related gene to regulate swarming activity

Swarming motility is a coordinated movement where bacteria use both flagella and type IV pili to navigate on semisolid surfaces. Our swarming motility assay showed that the wild‐type JG43 initially exhibited a lag phase in colony growth, followed by a rhythmic cycle of coherent colony expansion, reaching a diameter of 1.66 cm at 5 days after inoculating 2 μL of bacterial suspension onto the centre of a semisolid agar plate (0.6% agar) at 28°C. The swarming ability of the ∆virB4, ∆virB5, ∆virB8, ∆virB10, and ∆virD4 mutants was similar to that of the wild‐type JG43. On the other hand, the swarming motility of the ∆virB11 mutant was significantly reduced, with a diameter of 1.03 cm, and P∆virB11 was 1.14 cm (p < 0.0001) compared to both the wild‐type strain JG43 and the complementation strain CM∆virB11 (Figure 8a,b). This reduction is due to the ∆virB11 mutant losing both flagella and pilus morphogenesis, essential for bacterial swarming on semisolid surfaces. The DEGs related to bacterial chemotaxis in the KEGG pathway indicated the regulatory role of virB11 in the chemotaxis of Xal JG43. RNA‐seq analysis revealed 11 DEGs involved in chemotaxis, including one upregulated gene and 10 downregulated genes (Tables 4 and S1). In the chemotaxis signalling pathway, the ∆virB11 mutant showed downregulation of essential genes, such as cheA, cheW, cheY, cheR, and cheB (Figure 8c). The downregulation of these five chemotaxis‐related genes in the ∆virB11 mutant was confirmed by RT‐qPCR analysis (Figure 8d). These findings indicated that the absence of the virB11 gene affected the chemotaxis system, potentially impairing the swarming ability of the mutant strain.

FIGURE 8 Swarming motility of JG43 and its mutants, and schematic diagram of the differentially expressed genes (DEGs) involved in chemotaxis in the regulons of ΔvirB11. (a) The swarming assay was conducted on the JG43 and its mutants (ΔvirB4, ΔvirB5, ΔvirB8, ΔvirB10, ΔvirB11, ΔvirD4, virB11‐complemented strain CMΔvirB11 and vector control strain PΔvirB11) grown in MW medium for 5–6 days after inoculation of 2 μL each bacterial suspension to the centre of a medium with 0.6% agar and specific agar (Aladdin) at 28°C. Experiments were repeated three times independently with similar results. (b) Swarming motility measured by colony diameters of each strain on swarming medium plate. The bars represent standard errors of the means from three experiments, and each experiment contained three replicates for each strain. Statistical analysis was done using a t test, and statistical significance was denoted by **p < 0.01, ****p < 0.0001; ns, not significant p > 0.05. (c) Schematic diagram of the DEGs involved in chemotaxis. (d) Chemotaxis DEGs with five selected genes reverse transcription‐quantitative PCR (RT‐qPCR) verification, all RT‐qPCRs were performed three or more times independently using primers listed in Table S2.

TABLE 4 Differentially expressed genes involved in chemotaxis in Xanthomonas albilineans mutant strain ΔvirB11 compared to its wild‐type strain JG43.

Gene ID	Gene	Gene function	Log2FC	p‐value	
GE001228	cheB	Chemotaxis response regulator protein CheB	−1.03	0.0025	
GE001226	cheR	Chemotaxis protein CheR	−2.14	0.0001	
GE001225	–	MCP four helix bundle domain‐containing protein	−1.07	0.0057	
GE001223	cheW	Chemotaxis protein CheW	−1.72	0.0011	
GE001220	–	MCP four helix bundle domain‐containing protein	−1.94	0.0010	
GE001217	cheM	CHASE3 domain‐containing protein CheM	−3.00	0.0001	
GE001216	cheA	Chemotaxis protein CheA	−1.55	0.0057	
GE001215	cheY	Chemotaxis protein CheY	−1.77	0.0002	
GE001209	cheA	Chemotaxis protein CheA	1.39	0.0231	
GE001145	–	Chemotaxis protein	−1.18	0.0022	
GE000460	–	MCP four helix bundle domain‐containing protein	−2.65	0.0001	
Note: Gene ID refers to the locus_tag of DEGs X. albilineans ΔvirB11 strain compared to strain JG43, identified by BLASTn search against the strain JGT43 genome (GenBank accession number GCA_024158005.1). FC, fold‐change, |log2‐fold change| was derived from the mutant versus the wild‐type strain, |log2 fold‐change| > 1, or p‐value < 0.05.

Abbreviation: DEGs, differentially expressed genes.

3 DISCUSSION

The findings presented herein demonstrated that the deletion of the virB11 gene in Xal resulted in reduced swarming and virulence, as well as a complete loss of swimming and twitching motility due to the inability to develop flagella and type IV pilus structures. However, the growth ability of the mutant strains was similar to that of the wild‐type strain JG43.

The T4SS is crucial for bacterial conjugation and the transport of virulence factors by pathogenic bacteria. The proteins that make up the T4SS are conserved in both plant and animal pathogens, such as Agrobacterium tumefaciens and Legionella pneumophila (Cascales & Christie, 2003; Jeong et al., 2015). However, the T4SS in Xanthomonas has not been reported to secrete virulence factors into host cells. The T4SS evolved from ancestral conjugation systems for specialized purposes related to bacterial colonization or infection. For instance, Agrobacterium tumefaciens uses the VirB/VirD4 T4SS to deliver oncogenic DNA (T‐DNA) and effector proteins to plant cells, resulting in the tumorous disease known as crown gall (Gordon & Christie, 2014; Li & Christie, 2018; Zupan et al., 2000). Conjugation in gram‐negative bacteria is mediated by the T4SS, a complex macromolecular structure involved in substrate transport and pilus biogenesis. VirB11 interacts with the coupling protein VirD4 during plasmid and protein transfer by assisting TrwB of the type IV coupling protein (T4CP) in transporting the nucleoprotein substrate (Ripoll‐Rozada et al., 2013). The DNA substrate is initially recruited to the T4SS by T4CP of VirD4, a member of the ATP‐binding subunit family associated with bacterial conjugation systems (Llosa et al., 2002). VirD4 then transfers the substrate to VirB11, an ATPase localized at the cytoplasmic face of the inner membrane. Subsequently, the substrate is delivered to the inner membrane subunits VirB6 and VirB8, with VirB4 facilitating the transfer to these subunits. Finally, VirB6 and VirB8 collaborate to deliver the substrate to the periplasmic and outer membrane‐associated proteins VirB2 and VirB9 for passage to the cell exterior (Cascales & Christie, 2003). VirB11 works as a switch for DNA transport in T4SSs. DNA transfer requires specific interactions that prevent the replacement of VirB11, even with closely related homologues (Cascales et al., 2013). Recent findings support a model in which the early stages of DNA transfer are triggered by DNA ligand binding to VirD4 and VirB11, leading to ATP hydrolysis and subsequent conformational changes in the T4SS channel (Cascales et al., 2013). A tripartite interaction (TrIP) assay also confirmed that mutations in ∆virD4, ∆virB11, and ∆virB4 completely blocked substrate binding to the T4CP and substrate transfer to VirB6 and VirB8, respectively (Atmakuri et al., 2004), suggesting that these subunits energize the transfer reaction through an ATP‐dependent mechanism (Atmakuri et al., 2004).

The coupling of ATP binding and hydrolysis to macromolecular secretion systems is essential for the pathogenicity of gram‐negative bacteria (Ripoll‐Rozada et al., 2013). Our results showed that only the virB11‐deletion mutant (ΔvirB11) failed to develop flagella and type IV pili, resulting in reduced swarming, swimming, twitching motility, and decreased virulence. Deleting other virB/virD4 genes (∆virB4, ∆virB5, ∆virB8, ∆virB10, and ∆virD4) did not affect swarming, swimming, or twitching motility in Xal. In conclusion, the irreplaceability of VirB11 means that its removal hinders DNA transfer and disrupts ATP binding/hydrolysis coupling, which blocks substrate binding to the T4CP and substrate transfer to VirB6 and VirB8. This disruption affects the energy balance within the cell and the function of the T4SS.

Twitching is a crucial process that allows bacterial cells to adhere to surfaces and colonize hosts (Mattick, 2002). This process involves single cells undergoing sequential extension, attachment, and retraction of type IV pilus (Kühn & Talà, 2021). Type IV pilus‐like systems are essential for virulence in many bacterial pathogens (McCallum, Benlekbir, et al., 2019). The type IV pilus (T4P) is a flexible surface filament with a diameter of 4–7 nm and a length of several micrometres that can extend, adhere to surfaces, and retract (Dunger et al., 2016). The polymerization and depolymerization of pilin subunits, powered by motor ATPases from the PilT/VirB11‐like family, drive the functioning of the pilus (McCallum, Benlekbir et al., 2019). The pilus consists of pilin subunits, a pre‐pilin peptidase, and a motor (McCallum, Burrows, et al., 2019). Additionally, the pilus polymerization aids the extraction of the columnar protein from the membrane by using a cytoplasmic motor and ATP hydrolysis energy, while the motor is made up of two well‐conserved components with PilT/VirB11‐like ATPase and PilC‐like inner‐membrane protein (Chiang et al., 2008; LaPointe & Taylor, 2000; McCallum, Benlekbir, et al., 2019). Consequently, PilT/VirB11‐like proteins rotate PilC‐like proteins to extract columnar proteins from the membrane and insert them into the growing pilus polymer, thus promoting the polymerization of columnar proteins (Chang et al., 2016; McCallum et al., 2017). The PilT/VirB11‐like ATPase is responsible for T4P depolymerization, which depolymerizes quickly at the base, leading to fibre retraction (Merz et al., 2000). VirB11 proteins are traffic ATPases localized at the cytoplasmic face of the inner membrane (Atmakuri et al., 2004; Planet et al., 2001). These cytoplasmic hexametric proteins, belonging to the AAA+ ATPase family, govern the processes of pilus polymerization and depolymerization (Atmakuri et al., 2004; Hare et al., 2006; Planet et al., 2001; Ripoll‐Rozada et al., 2013). The absence of v irB11 significantly affects the pilus polymerization and depolymerization processes, consequently affecting twitching motility and virulence. In this study, TEM analysis demonstrated that the virB11‐deleted mutant could not form pili. Deleting the virB11 gene led to upregulation of the fimU and pilE genes in the mutant, which are predicted to produce minor pilins and contain a component set required for pilus assembly (Taguchi & Ichinose, 2011). The assembly and function of T4P requires the presence of these two minor subunits, and any loss of these subunits abolishes the pilus assembly in the ∆virB11 mutant strain (Nguyen et al., 2015). Pilus assembly is essential for T4P‐mediated bacterial motility and is driven by a cytoplasmic ATPase (Collins et al., 2018; Dye & Yang, 2020). As an ATPase, the deletion of virB11 prevented the ATP hydrolysis necessary for pilus morphogenesis, polymerization, and depolymerization. Interestingly, the CMΔvirB11 strain did not restore T4P morphogenesis but significantly downregulated fimU or pilE genes. These findings imply that the concentration of intracellular ATP might regulate T4P assembly and activity in vivo, making this process unexpectedly complex. This complementation failure suggests that the absence of virB11 has a profound and possibly irreversible impact on the assembly or function of the type IV pili, which are essential for twitching motility. The inability to recover the twitching phenotype upon complementation indicates that other regulatory or structural components influenced by VirB11 might be irreparably disrupted in the mutant, highlighting the critical and complex role of VirB11 in maintaining type IV pilus functionality and bacterial motility.

Flagella are crucial for the swimming motility of gram‐negative bacteria. They are complex nanomachines powered by ions and follow specific assembly pathways for efficient synthesis (Sanchez et al., 2023; Wadhwa & Berg, 2022). The flagellar assembly process involves coordinated steps, starting with the motor and export gate formation, followed by the construction of the extracellular propeller structure (Rossi et al., 2023). The bacterial flagellar motor, a supramolecular protein machine, propels flagellum rotation, enabling bacterial motility and survival in various environments while influencing pathogenicity (Tan et al., 2021). A reversible motor facilitates the rotation of flagellar filaments. The membrane‐embedded MS‐ring motor (MotA and MotB) surrounds the rotor, inducing rotation (Wadhwa & Berg, 2022). The C‐ring determines the direction of rotation by linking to the rotor element. A flexible hook connects the rod to the filament, transmitting the rotation, whereas the P and L rings help hold the rod within the cell envelope, facilitating flagellar filament rotation (Johnson & Furlong, 2021; Tan et al., 2021). These components are essential for flagellar assembly and rotation.

Swimming, accomplished through the rotation of rigid helical filaments, generates thrust and counteracts the adhesive force acting on the cell body, propelling the cell forward (Wadhwa & Berg, 2022). The flagellar filaments primarily consist of flagellin, which provides bacteria with motility and chemotactic abilities (Luo et al., 2023). Despite being just 20 nm in diameter, which is too small to be observed using an ordinary light microscope, stained and tagged flagellar filaments become visible under dark‐field observation due to scattered light (Wadhwa & Berg, 2022). In this study, we stained the bacterial flagella and observed the loss of flagella in the ∆virB11 mutant. Our findings are consistent with previous studies showing that the loss of flagellar function, such as in the fliC‐null flagellar filament, is associated with the absence of swimming ability, compromised adhesion, and biofilm formation (Newton et al., 1991; Wu et al., 2022). Mutations in genes like flhF, which controls the number and position of polar flagella in Vibrio species, and flhB, which encodes a flagellar protein component, have been shown to result in the loss of flagellar formation (Kondo et al., 2018; Kovtunov et al., 2013). Similarly, mutation in the motor switch gene fliM leads to abnormalities in flagellar formation and rotation (Homma et al., 2022), and the deletion of the flagella assembly gene fliH causes a loss of swimming motility (Dutta & Lee, 2022). In the current research, we observed the upregulation of flhF, flhB, fliC, fliM, and fliH genes in the ∆virB11 mutant, indicating an attempt to regulate flagellar assembly. However, the deletion of the virB11 gene caused irregular flagellar assembly, ultimately leading to the failure to form flagella. Swimming by the rotation of flagellar filaments is the most extensively studied mode of bacterial motility (Niu & Zhang, 2023; Wadhwa & Berg, 2022; Wu et al., 2022). The ∆virB11 mutant of Xal lost its swimming ability due to the absence of flagella. However, the expression patterns of flhF, flhB, fliC, fliM, and fliH genes in CM∆virB11 were consistent with those observed in the wild‐type JG43, suggesting that the lack of flagella resulted from defects in flagellin assembly.

Chemotaxis is also crucial in microbial motility and virulence (Porter et al., 2011). Both motility and chemotaxis play significant roles in the collective behaviours of bacteria, including swarming (Colin et al., 2021). Chemotaxis involves continuously monitoring specific chemical concentrations by methyl‐accepting chemotaxis proteins (MCPs) on motile cells. Transcriptome analysis revealed that DEGs encoding transmembrane chemoreceptors are involved in chemotaxis signal transduction. MCPs can detect various chemical signals, including attractants and repellents (Huang et al., 2019; Porter et al., 2011). The biochemical network determines motility behaviour by regulating the probability of flagellar motor rotation in either a counterclockwise or clockwise direction through the phosphorylation of a small protein called CheY. When phosphorylated CheY binds to the FliM motor, the bidirectional rotation of the flagellar motor is driven by the unidirectional rotation of MotA around MotB (Porter et al., 2011).

MCP‐promoted bacterial swarming and nutrient consumption depend on the synchronization of cell elongation and surfactant secretion, which enables coordinated swarming (Kearns, 2010). This suggests enhancing chemotaxis and motility might improve survival without flagella and pili. Bacterial swarming can occur without flagella through a reversal or reorientation frequency adjustments during surface motility (Kearns, 2010; Kearns & Shimkets, 1998). However, the role of chemotaxis signal transduction proteins in swarming is not primarily to direct movement. Although these proteins are essential for swarming, they do not facilitate the typical chemotactic orientation characterized by running and tumbling behaviours necessary for directed movement (Kearns, 2010). Therefore, the modulation of the transcriptome of chemotaxis‐related genes consistently occurred in the ∆virB11 mutant, resulting in reduced swarming motility. Our findings revealed that the loss of swimming and twitching abilities in the virB11‐deletion mutant is due to the absence of flagella‐driven swimming and T4P‐mediated twitching. The mutant relies on chemotaxis for swarming, but this ability was reduced compared to the wild‐type JG43. Additionally, we found that virulence factors such as protease, lipases, rhamnolipids, and motility play significant roles in bacterial virulence (Craig et al., 2004; Persat et al., 2015; Wadhwa & Berg, 2022; Xu et al., 2023). In summary, the virB11‐deletion mutant (∆virB11) lost swimming and twitching abilities, reduced swarming behaviour, and decreased overall virulence. Notably, the impact on motility did not affect the growth of the ∆virB11 mutant, as motility and colony expansion are primarily independent, and the presence of a surfactant significantly influences colony growth (Be'er et al., 2009).

4 EXPERIMENTAL PROCEDURES

4.1 Bacterial strains culturing

Xanthomonas albilineans JG43, its mutant strains (∆virB11, ∆virB4, ∆virB5, ∆virB8, ∆virB10, ∆virD4), ∆virB11 complemented strain CMΔvirB11, vector control strain PΔvirB11 were grown on an MW medium with 1 L consisting of peptone (5 g), Mg2SO4·7H2O (0.25 g), KH2PO4 (0.5 g), sucrose (10 g), Na2SO3 (0.05 g) and agar powder (10 g), pH 6.8–7.0, supplemented with 50 μg/mL of rifampicin (Rif).

4.2 Construction of the virB11 mutant and its complemented strain CMΔvirB11

The PCR primer pairs used to amplify the mutant strain ∆virB11 and the complementation strain CM∆virB11 were designed based on the upstream and downstream sequences of the wild‐type JG43 genome (GenBank accession number CP000512.1) (Tables 1 and S2). All primers for this study were designed using the Vector NTI software (Li, Li, et al., 2022).

PCR amplification for constructing the ∆virB11 mutant involved obtaining a 519‐bp upstream fragment (virB11L‐F/virB11L‐R) and a 574‐bp downstream fragment (virB11R‐F/virB11R‐R) using the JG43 genome as a template (Table 1). After confirmation by sequencing, the upstream (BamHI/XbaI) and downstream (XbaI/HindIII) PCR fragments were digested with the appropriate restriction enzymes and ligated into the pK18mobsacB (kanamycin‐resistant) vector, resulting in the recombinant plasmid. This plasmid was then introduced into competent JG43 cells via electrotransfer, generating the recombinant strain (pK/virB11) (Table 2). Confirmation of the recombinant pK/virB11 was performed by screening on MW medium supplemented with 10% sucrose and rifampicin (Rif) and kanamycin (Kan) antibiotics, followed by PCR verification (virB11L‐F/virB11R‐R). The amplified PCR product lacking the virB11 gene was verified by the absence of amplification by the primer pair virB11F/virB11R, which spans from 1 to 515 bp of the virB11 gene (Table 1). The construction process for the other five mutants (∆virB4, ∆virB5, ∆virB8, ∆virB10, ∆virD4) followed the same procedure as that for ∆virB11.

The complemented strain CMΔvirB11 was constructed by introducing the virB11 ORF and 300–500 bp upstream sequences. A 1056‐bp segment encompassing the virB11 gene and a 518‐bp upstream region were amplified by PCR using primers CMΔvirB11‐L and CMΔvirB11‐R (Table 1). After sequencing confirmation, the PCR fragments (BamHI/HindIII) were digested with appropriate restriction enzymes and ligated into the pLAFR6 (tetracycline‐resistant) vector to create the recombinant plasmid (pL/virB11) (Table 2). The pL/virB11 plasmid was screened on MW medium supplemented with 10% sucrose and Rif and tetracycline (Tc) and confirmed through PCR using CMΔvirB 11‐L and CMΔvirB11‐R primers, then the pL/virB11 plasmid was introduced into the ΔvirB11 mutant via electroporation, resulting in the complemented strain CMΔvirB11. Colonies showing resistance to Tc on MW agar supplemented with Tc indicated successful complementation, confirmed by PCR amplification of the 1574 bp fragment. The ∆virB11 strain was also transformed with the empty vector pLAFR6 by electroporation to construct the vector control strain P∆virB11 (Guan et al., 2020; Jiang et al., 2018) (Table 2).

4.3 Preparation of competent cells

Xanthomonas albilineans JG43 and ∆virB11 strains were grown in 15 mL of liquid medium containing 50 μg/mL Rif until they reached an OD600 of 0.6–0.8. The cultures were then cooled in the refrigerator for 30 min. After cooling, the cultures were divided into EP tubes and centrifuged at 12,000 g for 1 min at 4°C. The supernatant was removed, and each tube was filled with 500 μL of distilled water. The contents of the tubes were combined and centrifuged again under the same conditions. The resulting pellet of Xal competent cells was resuspended in 1 mL of distilled water, centrifuged, and washed three times with distilled water. Finally, 100 μL of distilled water was added to the pellet, which was kept on ice for plasmid transduction. All pipette tips and distilled water were prechilled to maintain low temperatures (Wang & Tseng, 1992).

4.4 Inoculation and virulence analysis

The JG43, ΔvirB11, and CMΔvirB11 strains were diluted in MW medium containing 50 μg/mL Rif, and their OD600 was adjusted to 0.5 for inoculation. The virulence of the wild‐type JG43 and its mutants (ΔvirB11 and CMΔvirB11) was tested on 6‐week‐old sugarcane seedlings. Leaf scald symptoms were recorded on emerging leaves after being cut off above the sugarcane growing point (Champoiseau et al., 2006). The virulence assay was conducted, and the disease index (DI) was evaluated 90 days after inoculation using a scale from 0 to 5, as described by Hong et al. (2021). The disease severity scale was as follows:

0: healthy leaves without white streaks; 1: white line streaks on the leaves; 2: two or more white line streaks on the leaves; 3: yellow or white stems and leaves; 4: plant death or abnormal sprouting of lateral buds; and 5: complete plant death.

The DI was calculated using the following formula (Hong et al., 2021): Disease index=∑Number of diseased plants in each grade×value of each grade/Total number of plants investigated×highest grade value.

4.5 Growth assay

The wild‐type strain JG43 and its mutants were cultivated on MW medium to assess their in vitro growth ability. Each Xal strain was inoculated into 300 mL of fresh MW medium with a 3 mL cell suspension (OD600 = 1.0), representing 1% of the total volume. The cultures were incubated at 28°C with shaking at 210 rpm. The OD600 of the suspension was measured every 6 h for 114 h using a microplate reader (Thermo Fisher Scientific Inc.). The growth curves were plotted using Origin 2022 software. The x‐axis represented the time in hours, and the y‐axis represented OD600 on a log10 scale (https://ea‐origin.en.softonic.com/) (Artier et al., 2018; Roy et al., 2022; Soares et al., 2010).

4.6 Motility test

All Xal strains were incubated in MW medium for swimming motility testing until they reached an OD600 of 1.0. A toothpick was used to inoculate each cell suspension onto the centre of a plate containing tryptone (0.3 g/L), yeast extract (0.3 g/L), and agar (0.3%), with a pH of 7.0. The plates were then incubated at 28°C for 4 days without shaking. After the incubation period, the colony diameter was measured following the method described by Guan et al. (2020).

Each Xal strain was adjusted to an OD600 of 0.2 for swarming motility testing. Then, 2 μL of this suspension was streaked onto the surface of an MW medium plate containing 0.6% agar. The plates were incubated at 28°C for 5–6 days without shaking. After the incubation period, the colony diameter was measured according to the method described by Doshi and Shaw (2023).

Twitching motility was investigated by incubating individual colonies of Xal strains overnight and adjusting their OD600 to 1.0. Sterile toothpicks were used to inoculate each colony at the interface between the plate and the twitching medium, which contained tryptone (3 g/L), yeast extract (1.5 g/L), NaCl (3 g/L), and agar (10 g/L). The plates were incubated upside down at 28°C for 4 days. The colony shape at the inoculation point was then observed. After removing the agar and washing off non‐adherent bacteria with deionized water, the plates were stained with a 2% crystal violet solution for 30 min. The unbound crystal violet dye was rinsed off with deionized water. Finally, the diameter of the bacterial colonies was measured as described by Kilmury and Burrows (2018), McCallum, Benlekbir, et al. (2019), and Ye et al. (2022).

4.7 Flagella and pilus observations by TEM

Formvar‐coated copper grids were suspended in a diluted bacterial solution (50–100 μL) for 3–5 min for flagella observations. The grids were then rinsed with distilled water, cleaned, and dried with filter paper. Afterward, the grids were immersed in a 2% uranyl solution for 30 s, followed by a 2–3 min treatment with uranium acetate. To observe pili, the Formvar‐coated copper grids were soaked in the phosphotungstic acid dye for 30 s and then washed two or three times with sterile water (McGroarty, 1994; Müller & Klug, 1979). Excess liquid was removed with filter paper, and the grids were air‐dried for 3–5 min. The samples were then examined using a TEM apparatus (HT‐7700) operating at an acceleration voltage of 80 kV (Guan et al., 2020).

4.8 Bacterial culture, RNA isolation and RT‐qPCR analysis

Wild‐type JG43 and the deleted mutant ΔvirB11 were grown in MW medium with Rif at 28°C and 210 rpm overnight until they reached an OD600 of 1.0. Three biological replicates were prepared for each strain. Bacterial cells (2 mL) were collected and centrifuged for 1 min at 12,000 g at 4°C. This process was repeated with another 1 mL of cells. After centrifugation, the supernatants were carefully removed, and the cell pellets were rapidly frozen in liquid nitrogen for 3 min and then stored at −80°C. Bacterial RNA was extracted using a bacterial RNA kit (Promega (Beijing) Biotech Co., Ltd.) according to the manufacturer's instructions. cDNA synthesis and qPCR detection were performed using a FastQuant RT kit (TianGen), following the manufacturer's instructions. RT‐qPCR analysis used specific primers designed for the DEGs (Table S1). The RT‐qPCR assay was conducted using a Roche LightCycler 96 instrument (Guan et al., 2020).

4.9 RNA‐Seq data analysis

The sequencing data was analysed by Shanghai Personal Bio‐Technology Co., Ltd., in Shanghai, China. DEGs between the wild‐type JG43 and its mutant strain ΔvirB11 were analysed using the Agilent 2100 Bioanalyzer. The raw reads were mapped to the genomic reference sequence of the Xal strain JG43 using Bowtie2 (http://bowtie‐bio.sourceforge.net/index.shtml). DESeq was used to identify DEGs based on p‐values < 0.05. The fold change of reads per kilobase per million mapped reads (RPKM) between the mutant and the wild type was calculated. A threshold of log2 fold change > |1.0| and p < 0.05 was applied for identifying significant DEGs.

CONFLICT OF INTEREST STATEMENT

The authors declare that they have no competing interests.

Supporting information

Figure S1. (a) Verification of upstream and downstream fragments of the virB11 gene. M: 1 kb marker, lane 1: verification of upstream fragments of the virB11 gene, lane 2: verification of downstream fragments of the virB11 gene. (b) Verification of the reconstructed plasmid pL‐virB11. M: 5 kb marker, lanes 1–5: restriction digestion and PCR were used to verify the reconstructed plasmid of pK18mobsacB containing virB11. (c) Verification of ΔvirB11 inter (virB11‐L/ virB11‐R) and outer primers (virB11‐L‐L/ virB11‐R‐R). M: 2 kb marker, lane 1: validation of internal primers using ΔvirB11 genomic DNA as template, lane 2: validation of internal primers using JG43 genomic DNA as template, lane 3: Validation of outer primers using ΔvirB11 genomic DNA as template, lane 4: validation of outer primers using JG43 genomic DNA as template, lane 5: validation of internal primers using water as template.

Table S1. ΔvirB11 regulated genes in the transcriptome.

Table S2. The sequence of quantitative PCR primers was used in the present study.

ACKNOWLEDGEMENTS

This work was supported by the China Agricultural Research System of MFA and MARA (CARS 170109) and the Guangxi Science and Technology Major Project (Guike AD20207020).

DATA AVAILABILITY STATEMENT

The sugarcane materials (Saccharum officinarum) used in the experiment were supplied by the sugarcane clonal germplasm repository of Guangxi University. The data that supports the findings of this study are available in the supplementary material of this article.
==== Refs
REFERENCES

Alvarez‐Martinez, C.E. & Christie, P.J. (2009) Biological diversity of prokaryotic type IV secretion. Microbiology and Molecular Biology Reviews, 73 , 775–808.19946141
Alvarez‐Martinez, C.E. , Sgro, G.G. , Araujo, G.G. , Paiva, M.R.N. , Matsuyama, B.Y. , Guzzo, C.R. et al. (2021) Secrete or perish: the role of secretion systems in Xanthomonas biology. Computational and Structural Biotechnology Journal, 19 , 279–302.33425257
Artier, J. , Silva, F. , Souza, F.M. , Pauletti, B.A. , Leme, A.F.P. , Carnielli, C.M. et al. (2018) Comparative proteomic analysis of Xanthomonas citri ssp. citri periplasmic proteins reveal changes in cellular envelope metabolism during in vitro pathogenicity induction. Molecular Plant Pathology, 19 , 143–157.27798950
Atmakuri, K. , Cascales, E. & Christie, P.J. (2004) Energetic components VirD4, VirB11, and VirB4 mediate early DNA transfer reactions required for bacterial type IV secretion. Molecular Microbiology, 54 , 1199–1211.15554962
Be'er, A. , Smith, R.S. , Zhang, H.P. , Florin, E.L. , Payne, S.M. & Swinney, H.L. (2009) Paenibacillus dendritiformis bacterial colony growth depends on surfactant but not on bacterial motion. Journal of Bacteriology, 191 , 5758–5764.19617369
Cascales, E. , Atmakuri, K. , Sarkar, M.K. & Christie, P.J. (2013) DNA substrate‐induced activation of the Agrobacterium VirB/VirD4 type IV secretion system. Journal of Bacteriology, 195 , 2691–2704.23564169
Cascales, E. & Christie, P.J. (2003) The versatile bacterial type IV secretion systems. Nature Reviews Microbiology, 1 , 137–149.15035043
Champoiseau, P. , Daugrois, J.H. , Pieretti, I. , Cociancich, S. , Royer, M. & Rott, P. (2006) High variation in pathogenicity of genetically closely related strains of Xanthomonas albilineans, the sugarcane leaf scald pathogen, in Guadeloupe. Phytopathology, 96 , 1081–1091.18943496
Chang, Y.W. , Rettberg, L.A. , Treuner‐Lange, A. , Iwasa, J. , Søgaard‐Andersen, L. & Jensen, G.J. (2016) Architecture of the type IVa pilus machine. Science, 351 , 2001.
Chiang, P. , Sampaleanu, L.M. , Ayers, M. , Pahuta, M. , Howell, P.L. & Burrows, L.L. (2008) Functional role of conserved residues in the characteristic secretion NTPase motifs of the Pseudomonas aeruginosa type IV pilus motor proteins PilB, PilT and PilU. Microbiology, 154 , 114–126.18174131
Christie, P.J. (2016) The mosaic type IV secretion systems. EcoSal Plus, 7 , 10.
Christie, P.J. , Whitaker, N. & González‐Rivera, C. (2014) Mechanism and structure of the bacterial type IV secretion systems. Biochimica et Biophysica Acta, Bioenergetics, 1843 , 1578–1591.
Colin, R. , Ni, B. , Laganenka, L. & Sourjik, V. (2021) Multiple functions of flagellar motility and chemotaxis in bacterial physiology. FEMS Microbiology Reviews, 23 , 45.
Collins, R. , Karuppiah, V. , Siebert, C.A. , Dajani, R. , Thistlethwaite, A. & Derrick, J.P. (2018) Structural cycle of the Thermus thermophilus PilF ATPase: the powering of type IVa pilus assembly. Scientific Reports, 8 , 14022.30232337
Costa, T.R.D. , Harb, L. , Khara, P. & Zeng, L. (2021) Type IV secretion systems: advances in structure, function, and activation. Molecular Microbiology, 115 , 436–452.33326642
Craig, L. & Forest, K.T. (2019) Type IV pili: dynamics, biophysics and functional consequences. Nature Reviews Microbiology, 17 , 429–440.30988511
Craig, L. , Pique, M.E. & Tainer, J.A. (2004) Type IV pilus structure and bacterial pathogenicity. Nature Reviews Microbiology, 2 , 363–378.15100690
Doshi, A. & Shaw, M. (2023) Engineered bacterial swarm patterns as spatial records of environmental inputs. Nature Chemical Biology, 19 , 878–886.37142806
Dufrêne, Y.F. & Persat, A. (2020) Mechanomicrobiology: how bacteria sense and respond to forces. Nature Reviews Microbiology, 18 , 227–240.31959911
Dunger, G. , Llontop, E. , Guzzo, C.R. & Farah, C.S. (2016) The Xanthomonas type IV pilus. Current Opinion in Microbiology, 30 , 88–97.26874963
Dutta, S. & Lee, Y.H. (2022) High‐throughput identification of genes influencing the competitive ability to obtain nutrients and performance of biocontrol in Pseudomonas putida JBC17. Scientific Reports, 12 , 872.35042886
Dye, K.J. & Yang, Z. (2020) Cyclic‐di‐GMP and ADP bind to separate domains of PilB as mutual allosteric effectors. Biochemical Journal, 477 , 213–226.31868878
Gordon, J.E. & Christie, P.J. (2014) The Agrobacterium Ti plasmids. Microbiology Spectrum, 2 , 10.
Guan, W. , Wang, T. , Huang, Q. , Tian, E. , Liu, B. , Yang, Y. et al. (2020) A LuxR‐type regulator, AcrR, regulates flagellar assembly and contributes to virulence, motility, biofilm formation, and growth ability of Acidovorax citrulli . Molecular Plant Pathology, 21 , 489–501.31943660
Ha, D.G. , Kuchma, S.L. & Toole, G.A. (2014) Plate‐based assay for swimming motility in Pseudomonas aeruginosa . Methods in Molecular Biology, 1149 , 59–65.24818897
Hanahan, D. (1983) Studies on transformation of Escherichia coli with plasmids. Journal of Molecular Biology, 166 , 557–580.6345791
Hare, S. , Bayliss, R. , Baron, C. & Waksman, G. (2006) A large domain swap in the VirB11 ATPase of Brucella suis leaves the hexameric assembly intact. Journal of Molecular Biology, 360 , 56–66.16730027
Homma, M. , Takekawa, N. , Fujiwara, K. , Hao, Y. , Onoue, Y. & Kojima, S. (2022) Formation of multiple flagella caused by a mutation of the flagellar rotor protein FliM in Vibrio alginolyticus . Genes to Cells, 27 , 568–578.35842835
Hong, D.K. , Talha, J. , Yao, Y. , Zou, Z.Y. & Wang, J.D. (2021) Silicon enhancement for endorsement of Xanthomonas albilineans infection in sugarcane. Ecotoxicology and Environmental Safety, 220 , 112380.34058676
Huang, L. , Lin, L.X. , Su, Y. , Qin, Y. , Kong, W. , Zhao, L. et al. (2017) MCP, aer, cheB, and cheV contribute to the regulation of Vibrio alginolyticus (ND‐01) adhesion under gradients of environmental factors. Microbiology Open, 6 , 00257.
Huang, Z. , Pan, X. , Xu, N. & Guo, M. (2019) Bacterial chemotaxis coupling protein: structure, function and diversity. Microbiological Research, 219 , 40–48.30642465
Jeckel, H. , Jelli, E. , Hartmann, R. , Singh, P.K. , Mok, R. , Totz, J.F. et al. (2019) Learning the space‐time phase diagram of bacterial swarm expansion. Proceedings of the National Academy of Sciences of the United States of America, 116 , 1489–1494.30635422
Jeong, K.C. , Sexton, J.A. & Vogel, J.P. (2015) Spatiotemporal regulation of a Legionella pneumophila T4SS substrate by the meta‐effector SidJ. PLoS Pathogens, 11 , 1004695.
Jiang, B.L. , Jiang, G.F. , Liu, W. , Yang, L.C. , Yang, L.Y. , Wang, L. et al. (2018) RpfC regulates the expression of the vital regulator hrpX of the hrp/T3SS system in Xanthomonas campestris pv. campestris . BMC Microbiology, 18 , 103.30176800
Johnson, K.S. & Ottemann, K.M. (2018) Colonization, localization, and inflammation: the roles of H. pylori chemotaxis in vivo. Current Opinion in Microbiology, 41 , 51–57.29202336
Johnson, S. & Furlong, E.J. (2021) Molecular structure of the intact bacterial flagellar basal body. Nature Microbiology, 6 , 712–721.
Kearns, D.B. (2010) A field guide to bacterial swarming motility. Nature Reviews Microbiology, 8 , 634–644.20694026
Kearns, D.B. & Losick, R. (2003) Swarming motility in undomesticated Bacillus subtilis . Molecular Microbiology, 49 , 581–590.12864845
Kearns, D.B. & Shimkets, L.J. (1998) Chemotaxis in a gliding bacterium. Proceedings of the National Academy of Sciences of the United States of America, 95 , 11957–11962.9751772
Kerr, J.E. & Christie, P.J. (2010) Evidence for VirB4‐mediated dislocation of membrane‐integrated VirB2 pilin during biogenesis of the Agrobacterium VirB/VirD4 type IV secretion system. Journal of Bacteriology, 192 , 4923–4934.20656905
Kilmury, S.L.N. & Burrows, L.L. (2018) The Pseudomonas aeruginosa PilSR two‐component system regulates both twitching and swimming motilities. mBio, 9 , 1310–1318.
Kondo, S. , Imura, Y. , Mizuno, A. , Homma, M. & Kojima, S. (2018) Biochemical analysis of GTPase FlhF which controls the number and position of flagellar formation in marine Vibrio . Scientific Reports, 8 , 12115.30108243
Kovtunov, E.A. , Petrova, L.P. , Shelud'ko, A.V. & Katsy, E.I. (2013) Insertional mutagenesis of a chromosomal copy of flhB gene is concurrent with defects in the formation of polar and lateral flagella in bacterium Azospirillum brasilense Sp245. Genetika, 49 , 1013–1016.25474889
Kuchma, S.L. , Delalez, N.J. , Filkins, L.M. , Snavely, E.A. , Armitage, J.P. & Toole, G.A. (2015) Cyclic di‐GMP‐mediated repression of swarming motility by Pseudomonas aeruginosa PA14 requires the MotAB stator. Journal of Bacteriology, 197 , 420–430.25349157
Kühn, M.J. & Talà, L. (2021) Mechanotaxis directs Pseudomonas aeruginosa twitching motility. Proceedings of the National Academy of Sciences of the United States of America, 118 , e2101759118.34301869
LaPointe, C.F. & Taylor, R.K. (2000) The type 4 prepilin peptidases comprise a novel family of aspartic acid proteases. Journal of Biological Chemistry, 275 , 1502–1510.10625704
Li, M. , Bao, Y. , Li, Y. , Akbar, S. , Wu, G. , Du, J. et al. (2022) Comparative genome analysis unravels pathogenicity of Xanthomonas albilineans causing sugarcane leaf scald disease. BMC Genomics, 23 , 671.36162999
Li, Y. , Li, M. , Du, J. & Zhang, M. (2022) Ultrastructure sugarcane leaves and stems tissue infected by Xanthomonas albilineans . Acta Phytopathologica Sinica, 52 , 763–768.
Li, Y.G. & Christie, P.J. (2018) The Agrobacterium VirB/VirD4 T4SS: mechanism and architecture defined through in vivo mutagenesis and chimeric systems. Current Topics in Microbiology and Immunology, 418 , 233–260.29808338
Li, Y.G. , Hu, B. & Christie, P.J. (2019) Biological and structural diversity of type IV secretion systems. Microbiology Spectrum, 7 , 10.
Llosa, M. , Gomis‐Rüth, F.X. , Coll, M. & de la Cruz, F.F. (2002) Bacterial conjugation: a two‐step mechanism for DNA transport. Molecular Microbiology, 45 , 1–8.12100543
Luo, Y. , Wang, J. , Gu, Y.L. , Zhang, L.Q. & Wei, H.L. (2023) Duplicated flagellins in pseudomonas divergently contribute to motility and plant immune elicitation. Microbiology Spectrum, 14 , 11.
Malamud, F. , Torres, P.S. , Roeschlin, R. , Rigano, L.A. , Enrique, R. , Bonomi, H.R. et al. (2011) The Xanthomonas axonopodis pv. citri flagellum is required for mature biofilm and canker development. Microbiology, 157 , 819–829.21109564
Mary, C. , Fouillen, A. , Bessette, B. , Nanci, A. & Baron, C. (2018) Interaction via the N terminus of the type IV secretion system (T4SS) protein VirB6 with VirB10 is required for VirB2 and VirB5 incorporation into T‐pili and for T4SS function. Journal of Biological Chemistry, 293 , 13415–13426.29976757
Mattick, J.S. (2002) Type IV pili and twitching motility. Annual Review of Microbiology, 56 , 289–314.
McCallum, M. , Benlekbir, S. , Nguyen, S. , Tammam, S. & Rubinstein, J.L. (2019) Multiple conformations facilitate PilT function in the type IV pilus. Nature Communications, 10 , 5198.
McCallum, M. , Burrows, L.L. & Howell, P.L. (2019) The dynamic structures of the type IV pilus. Microbiology Spectrum, 7 , 2.
McCallum, M. , Tammam, S. , Khan, A. , Burrows, L.L. & Howell, P.L. (2017) The molecular mechanism of the type IVa pilus motors. Nature Communications, 5 , 8.
McGroarty, J.A. (1994) Cell surface appendages of lactobacilli. FEMS Microbiology Letters, 124 , 405–409.7851748
Merz, A.J. , So, M. & Sheetz, M.P. (2000) Pilus retraction powers bacterial twitching motility. Nature, 407 , 98–102.10993081
Müller, G. & Klug, H. (1979) Zur Problematik der elektronenmikroskopischen Darstellung der Pilistruktur von Neisseria gonorrhoeae [Electron microscopic representation of the pili structure of Neisseria gonorrhoeae]. Dermatologische Monatsschrift, 165 , 18–27.
Newton, S.M. , Wasley, R.D. , Wilson, A. , Rosenberg, L.T. , Miller, J.F. & Stocker, B.A. (1991) Segment IV of a Salmonella flagellin gene specifies flagellar antigen epitopes. Molecular Microbiology, 5 , 419–425.1710314
Nguyen, Y. , Sugiman‐Marangos, S. , Harvey, H. , Bell, S.D. , Charlton, C.L. , Junop, M.S. et al. (2015) Pseudomonas aeruginosa minor pilins prime type IVa pilus assembly and promote surface display of the PilY1 adhesin. Journal of Biological Chemistry, 290 , 601–611.25389296
Niu, Y. & Zhang, R. (2023) Flagellar motors of swimming bacteria contain an incomplete set of stator units to ensure robust motility. Science Advances, 9 , 6724.
Oliveira, N.M. & Foster, K.R. (2016) Single‐cell twitching chemotaxis in developing biofilms. Proceedings of the National Academy of Sciences of the United States of America, 113 , 6532–6537.27222583
Persat, A. , Inclan, Y.F. , Engel, J.N. , Stone, H.A. & Gitai, Z. (2015) Type IV pili mechanochemically regulate virulence factors in Pseudomonas aeruginosa . Proceedings of the National Academy of Sciences of the United States of America, 112 , 7563–7568.26041805
Pieretti, I. , Pesic, A. , Petras, D. , Royer, M. , Süssmuth, R.D. & Cociancich, S. (2015) What makes Xanthomonas albilineans unique amongst xanthomonads? Frontiers in Plant Science, 6 , 289.25964795
Planet, P.J. , Kachlany, S.C. , DeSalle, R. & Figurski, D.H. (2001) Phylogeny of genes for secretion ATPases: identification of the widespread tadA subfamily and development of a diagnostic key for gene classification. Proceedings of the National Academy of Sciences of the United States of America, 98 , 2503–2508.11226268
Porter, S.L. , Wadhams, G.H. & Armitage, J.P. (2011) Signal processing in complex chemotaxis pathways. Nature Reviews Microbiology, 9 , 153–165.21283116
Raina, J.B. , Fernandez, V. , Lambert, B. , Stocker, R. & Seymour, J.R. (2019) The role of microbial motility and chemotaxis in symbiosis. Nature Reviews Microbiology, 17 , 284–294.30923350
Ripoll‐Rozada, J. , Peña, A. , Rivas, S. , Moro, F. , Cruz, F. , Cabezón, E. et al. (2012) Regulation of the type IV secretion ATPase TrwD by magnesium: implications for catalytic mechanism of the secretion ATPase superfamily. Journal of Biological Chemistry, 287 , 17408–17414.22467878
Ripoll‐Rozada, J. , Zunzunegui, S. , Cruz, F. , Arechaga, I. & Cabezón, E. (2013) Functional interactions of VirB11 traffic ATPases with VirB4 and VirD4 molecular motors in type IV secretion systems. Journal of Bacteriology, 195 , 4195–4201.23852869
Rossi, P. , Xing, Q. , Bini, E. , Portaliou, A.G. , Clay, M.C. , Warren, E.M. et al. (2023) Chaperone recycling in late‐stage flagellar assembly. Journal of Molecular Biology, 435 , 167954.37330284
Roy, S. , Mittal, P. , Tayi, L. , Bondada, S. , Ray, M.K. , Patel, H.K. et al. (2022) Xanthomonas oryzae pv. oryzae exoribonuclease R is required for complete virulence in rice, optimal motility, and growth under stress. Phytopathology, 112 , 501–510.34384245
Sanchez, J.C. , Montemayor, E.J. , Ploscariu, N.T. , Parrell, D. , Baumgardt, J.K. , Yang, J.E. et al. (2023) Atomic‐level architecture of Caulobacter crescentus flagellar filaments provide evidence for multi‐flagellin filament stabilization. bioRxiv. 10.1101/2023.07.10.548443. [Preprint].
Scharf, B.E. , Hynes, M.F. & Alexandre, G.M. (2016) Chemotaxis signaling systems in model beneficial plant–bacteria associations. Plant Molecular Biology, 90 , 549–559.26797793
Sgro, G.G. , Oka, G.U. , Souza, D.P. , Cenens, W. , Bayer‐Santos, E. , Matsuyama, B.Y. et al. (2019) Bacteria‐killing type IV secretion systems. Frontiers in Microbiology, 10 , 1078.31164878
Soares, M.R. , Facincani, A.P. , Ferreira, R.M. , Moreira, L.M. , Oliveira, J.C. , Ferro, J.A. et al. (2010) Proteome of the phytopathogen Xanthomonas citri subsp. citri: a global expression profile. Proteome Science, 8 , 55.21062441
Souza, D.P. , Oka, G.U. , Alvarez‐Martinez, C.E. , Bisson‐Filho, A.W. , Dunger, G. , Hobeika, L. et al. (2015) Bacterial killing via a type IV secretion system. Nature Communications, 6 , 6453.
Suaste‐Olmos, F. , Domenzain, C. , Mireles‐Rodríguez, J.C. , Poggio, S. , Osorio, A. , Dreyfus, G. et al. (2010) The flagellar protein FliL is essential for swimming in Rhodobacter sphaeroides . Journal of Bacteriology, 192 , 6230–6239.20889747
Taguchi, F. & Ichinose, Y. (2011) Role of type IV pili in virulence of Pseudomonas syringae pv. tabaci 6605: correlation of motility, multidrug resistance, and HR‐inducing activity on a nonhost plant. Molecular Plant–Microbe Interactions, 24 , 1001–1011.21615203
Tan, J. , Zhang, X. , Wang, X. , Xu, C. , Chang, S. , Wu, H. et al. (2021) Structural basis of assembly and torque transmission of the bacterial flagellar motor. Cell, 184 , 2665–2679.33882274
Thomas, N.A. , Mueller, S. , Klein, A. & Jarrell, K.F. (2002) Mutants in flaI and flaJ of the archaeon Methanococcus voltae are deficient in flagellum assembly. Molecular Microbiology, 46 , 879–887.12410843
Wadhwa, N. & Berg, H.C. (2022) Bacterial motility: machinery and mechanisms. Nature Reviews Microbiology, 20 , 161–173.34548639
Wang, T.W. & Tseng, Y.H. (1992) Electrotransformation of Xanthomonas campestris by RF DNA of filamentous phage phi Lf. Letters in Applied Microbiology, 14 , 65–68.1367905
Wu, C.M. , Huang, H.H. , Li, L.H. , Lin, Y.T. & Yang, T.C. (2022) Molecular characterization of three tandemly located flagellin genes of Stenotrophomonas maltophilia . International Journal of Molecular Sciences, 31 , 23.
Wu, H.Y. , Chen, C.Y. & Lai, E.M. (2014) Expression and functional characterization of the Agrobacterium VirB2 amino acid substitution variants in T‐pilus biogenesis, virulence, and transient transformation efficiency. PLoS One, 27 , 9.
Xu, K.Z. , Xiang, S.L. , Wang, Y.J. , Wang, B. & Jia, A.Q. (2023) Methyl gallate isolated from partridge tea (Mallotus oblongifolius Miq. Müll. Arg.) inhibits the biofilms and virulence factors of Burkholderia thailandensis . Journal of Ethnopharmacology, 10 , 320.
Ye, Z. , Ye, L. , Li, D. , Lin, S. , Deng, W. , Zhang, L. et al. (2022) Effects of daphnetin on biofilm formation and motility of Pseudomonas aeruginosa . Frontiers in Cellular and Infection Microbiology, 18 , 12.
Yuan, Q. , Carle, A. , Gao, C. , Sivanesan, D. , Aly, K.A. , Höppner, C. et al. (2005) Identification of the VirB4‐VirB8‐VirB5‐VirB2 pilus assembly sequence of type IV secretion systems. Journal of Biological Chemistry, 280 , 26349–26359.15901731
Zhu, D. , Wang, S. & Sun, X. (2021) FliW and CsrA govern flagellin (FliC) synthesis and play pleiotropic roles in virulence and physiology of Clostridioides difficile R20291. Frontiers in Microbiology, 5 , 12.
Zupan, J. , Muth, T.R. , Draper, O. & Zambryski, P. (2000) The transfer of DNA from Agrobacterium tumefaciens into plants: a feast of fundamental insights. The Plant Journal, 23 , 11–28.10929098
