crystal structure of the c-terminal domain of a flagellar hook-capping protein from xanthomonas...

doi:10.1016/j.jmb.2008.05.083 J. Mol. Biol. (2008) 381, 189–199

Available online at www.sciencedirect.com

Crystal Structure of the C-Terminal Domain of a FlagellarHook-Capping Protein from Xanthomonas campestris

Wei-Ting Kuo1, Ko-Hsin Chin1,2, Wen-Ting Lo1,Andrew H.-J. Wang3,4 and Shan-Ho Chou1,2*
1Institute of Biochemistry,National Chung-HsingUniversity, Taichung, 40227,Taiwan, Republic of China2National Chung HsingUniversity BiotechnologyCenter, National Chung-HsingUniversity, Taichung, 40227,Taiwan, Republic of China3Core Facility for ProteinCrystallography, AcademiaSinica, Nankang, Taipei,Taiwan, Republic of China4Institute of BiologicalChemistry, Academia Sinica,Nankang, Taipei, Taiwan,Republic of China
Received 29 February 2008;received in revised form29 May 2008;accepted 31 May 2008Available online7 June 2008

*Corresponding author.National ChBiotechnology Center, National ChuTaichung, 40227, Taiwan, Republic oE-mail address: [email protected] used: Se-Met, selen

fibronectin type III; PDB, Protein Da

0022-2836/$ - see front matter © 2008 E

The crystal structure of the C-terminal domain of a hook-capping proteinFlgD from the plant pathogen Xanthomonas campestris (Xc) has been deter-mined to a resolution of ca 2.5 Å using X-ray crystallography. The monomerof whole FlgD comprises 221 amino acidswith amolecular mass of 22.7 kDa,but the flexible N-terminus is cleaved for up to 75 residues during crystal-lization. The final structure of the C-terminal domain reveals a novel hybridcomprising a tudor-like domain interdigitated with a fibronectin type IIIdomain. The C-terminal domain of XcFlgD forms three types of dimers in thecrystal. In agreement with this, analytical ultracentrifugation and gel filtra-tion experiments reveal that they form a stable dimer in solution. From theseresults, we propose that the Xc flagellar hook cap protein FlgD comprisestwo individual domains, a flexible N-terminal domain that cannot be de-tected in the current study and a stable C-terminal domain that forms a stabledimer.

© 2008 Elsevier Ltd. All rights reserved.

Keywords: FlgD; hook capping; Xanthomonas campestris; Ig-like domain;Tudor-like domain
Edited by I. Wilson
Introduction

The bacterial flagellum is a very complicated na-nostructure assembled from more than 40 geneproducts.1–3 It comprises the basal body, hook, fila-ment, and other labile structures such as motor,switch, and export apparatus. The flagellum issequentially assembled from simpler to more com-plex structures.3 First, the MS (integral membrane

ung Hsing Universityng-Hsing University,f China..omethionine; Fn-III,ta Bank.

lsevier Ltd. All rights reserve

ring) ring complex is formed, with switch and exportapparatus later added. Basal body assembly thencontinues with the formation of rod and addition ofperiplasmic P and outer L rings. When rod assemblyis finished, hook assembly is then initiated. Finally,filament grows to a length of approximately 15 μm toform a functional flagellar structure, which providescell the capability to move to more favorable envi-ronments and to prevent stressful conditions.In a swimming Escherichia coli or Salmonella cell,

flagella rotate at high speed in the order of 300 Hz.4

However, it is important to note that flagella areattached on an immobile basal body. Hence, anintermediate substructure is necessary for a cell towithstand the enormous torsional force caused bythe highly rotating flagella. Such a “buffer” is servedby a hook substructure of around 55 nm.5,6 However,

d.

190 C-Terminal Domain Crystal Structure of Flagellar Hook-Capping Protein

to form the hook structure in vivo, a cap is necessaryto prevent the leakage of hook monomers into themedium. Furthermore, the hook cap should not bepermanently present and need to be removed whenhook structure is complete to continue the flagellarbiogenesis. The scaffolding protein FlgD is believedto be the protein to form the hook cap structure.7

Besides its obvious role as a cap, it also plays anactive role in productive hook monomer polymeri-zation7 and determination of correct hook length,with the help of FliK protein.8–10 Thus, althoughFlgD is not present in a mature flagellum, it playsdiverse roles in forming a functional hook structure.The importance of hook capping is clearly revealedin an flgD mutant, which cannot continue flagellarbiogenesis when basal body is completed.7 Howsuch a transient intermediate structure can performsomany critical functions during flagellar biogenesisis interesting to know.In the past decade, there have been considerable

progresses toward understanding the formation ofhook and filament substructures; the atomic struc-tures of hook FlgE and filament FliC subunits havebeen determined by X-ray crystallography, and theirpolymer structures have been solved by fitting thesingle crystal structures to the corresponding cryoe-lectron microscopy images.5,6,11 The coordinates ofthe crystal structures of two hook–filament junctionproteins, FlgK (2D4Y) and FlgL (2D4X), have alsobeen deposited. However, to date, no atomic capstructure of hook or filament type has beenpublished.We now report the first crystal structure of the C-terminal domain of a flagellar hook protein (XC1894)from the plant pathogen Xanthomonas campestris pv.campestris (Xc). Bioinformatics study indicates that itis a hook-capping protein. Final tertiary structure ofthe XC1894 C-terminal domain reveals that it adoptsa novel hybrid comprising a tudor-like domaininterdigitated with an FN-III domain. The monomeris highly enriched in β-strands and can interact withother subunits in three different ways.

Results

XC1894 exhibits N-terminal cleavage

When the full-length XC1894(221) was expressed,purified, and crystallized, good hexagonal-like crys-tals diffracted to a resolution of ca 2.5 Å were ob-tained after 14 days. However, when electrondensity map was calculated using these diffracteddata, only 138 amino acid residues could be clearlydefined, starting from the N-terminal Asp84 to theC-terminal Ser221. Many N-terminal residues wereinvisible in the map. The crystal was dissolved andsubmitted to matrix-assisted laser desorption/ioni-zation time of flight mass spectrometer for massanalysis to understand whether these residues aredisordered in crystal or cleaved during crystalformation. Surprisingly, no full-length mass peakwas detected at all. Instead, two peaks were found,one with a molecular mass of 14,612.24 Da and

another with a molecular mass of 14,997.71 Da (datanot shown), corresponding exactly to 146 and 150amino acid residues from the C-terminal end,respectively. It seems to be that full-length XC1894(221) has experienced cleavage from the N-terminus,with either 75 or 71 amino acids cleaved duringcrystallization. A time-course cleavage experimentwas performed to monitor the cleavage kinetics. Ascan be seen in lane a of Fig. 1, fresh XC1894(221) doeshave an intact molecular mass of roughly 23 kDa asexpected from the sequence when first purified.However, after 3 days, the intact protein has almostdisappeared, and a shorter fragment appeared atapproximately 20 kDa (lane b in Fig. 1). After 5 days,further degradation occurred, and new bands ataround 15 kDa appeared (lane c in Fig. 1). Finally,after 10 days, the bands at 23 or 20 kDa have entirelydisappeared and only bands at around 15 kDaremained (lane d in Fig. 1). A similar cleavagephenomenon occurs for shorter XC1894(187) proteinafter 10 days (lane e in Fig. 1). Interestingly, XC1894(150) protein seems to be immune to cleavage anddoes not exhibit any degradation even after 10 days(lane f in Fig. 1). These results indicate that the N-terminus of XC1894 before Asp84 is very mobile andcould be cleaved by an unknown mechanism.However, further deletion of the N-terminus resultsin protein instability, and no protein could beobtained when the shortest XC1894(137) gene wascloned and expressed. The data indicate that XC1894contains a stable core of around 150 amino acidresidues from the C-terminus and could be crystal-lized even when the disordered N-terminal sequenceis cleaved and degraded during crystallization.To investigate whether the cleavage is due to con-

tamination of a small amount of endogenous pro-teases during sample preparation,14 we have furtherperformed the cleavage experiment in the presence ofprotease inhibitor cocktail. The protease inhibitorcocktail is a mixture of inhibitors with broadspecificities toward serine proteases, cysteine pro-teases, acid proteases, metalloproteases, and amino-peptidases (Sigma, USA) and has been optimized toprevent protein degradation after induction duringprotein expression. Significantly, fresh FlgD wasagain truncated even in the presence of various inhi-bitors after 12 h treatment at 37 °C to a size of ap-proximately 15 kDa, as that happened in the absenceof the protease inhibitor cocktail (data not shown).However, it is still possible that the inhibitor cocktailis not complete and a very small amount of unknownendogenous protease may be present to execute thecleavage, but it is clear that the N-terminal region ofXcFlgD ismuchmore flexible andmore submissive tocleavage than the C-terminal region, as the cases inother flagellar axial proteins.15,16

XC1894 forms a stable dimer in solution

Analytical ultracentrifugation and gel filtrationexperiments have also been carried out to checkthe oligomeric state of the truncated XC1894(approximately 15 kDa) in solution. The analytical

Fig. 1. The time course ofXC1894auto-cleavage at 25 °C. The genefragments of different length wereprepared by PCR from genome ofX.campestris pv. campestris str. 17 usingprimers suitable for ligation-inde-pendent cloning approach.12,13 Thegene products were run on an SDS-PAGE to monitor the auto-cleavagekinetics. The first lane is frommarkerproteins. The auto-cleavage timecourse of XC1894(221) is shown inthe following lanes: (a) fresh protein,(b) after 3 days, (c) after 5 days, and(d) after 10 days. XC1894(187) alsoexhibits similar auto-cleavage phe-nomenon, but only the cleavage pro-duct after 10 days is shown in lane e.XC1894(150) remains unchangedafter 10days (lane f).On the contrary,XC1894(137) is very unstable, and noproduct could be obtained from thisclone.

191C-Terminal Domain Crystal Structure of Flagellar Hook-Capping Protein

ultracentrifugation experiments were performed atpH 8.0 and pH 5.5, Tris 20 mM, NaCl 80 mM bufferconditions. In both cases, a clear band at 30 kDa wasobserved. The gel filtration experiment was alsoperformed at a similar Tris buffer condition, and asimilar major band at 30 kDa position was alsorevealed. Both of these experiments indicate that theC-terminal domain of XC1894 forms a stable dimerin solution.

XC1894 is a flagellar hook-capping protein fromX. campestris

Figure 2 shows the multiple sequence alignment ofX. campestris XC1894 with the two well-studied FlgDsequences from E. coli17,18 and Salmonella typhi-murium.7 As can be seen from the alignment,XC1894 exhibits high sequence identity (36.9% and37.6%) and similarity (49.0% and 48.6%) with the E.coli and S. typhimurium FlgD sequences, respectively.The identical and similar residues also distributeevenly throughout the whole sequences (Fig. 2), in-dicating that XC1894 is very likely an FlgD protein.This conclusion is further strengthenedby the fact thatthe XC1894 gene is located in the middle of theflgBCDEF rod/hook protein operon, which is em-bedded in a gene cluster responsible for the flagellarfunction.21 Thorough BLAST sequence search againstthe UniProtKB/Swiss-Prot (http://us.expasy.org/sprot/) database returns XC1894 as a hook-cappingprotein also, with sequence identities of 39.7%,

41.0%, 63.0%, and 95.7% with the Q085P9, A4XW87,A1FR14, and Q3BU01 sequences, respectively. Fromthese bioinformatics studies, it is clear that XC1894 isa flagellar hook-capping protein from X. campestris.

The tertiary structure of the XC1894 monomer

The tertiary structure of the XC1894 monomer wasdetermined using a selenomethionine (Se-Met)-sub-stituted sample to solve the phase problem. Except forthe 75 N-terminal residues that are cleaved, the elec-tron density can be successfully traced from Asp84 toSer221. The Ramachandran plot calculated from thePdbViewer22 shows that 93.5% of the residues are inthe most favored regions, with 3.6% of the residues inthe additional allowed regions. Four residues, includ-ing Asp107, Ser111, Ala178, and Ala215 (2.9%), alllocated in the loop turn regions, are found to deviatefrom the allowed torsional angles. Residues Asp107and Ser111 are located in theAsp107AlaThrGlySer111loop turn connecting theβAstrand and theβB strand.Asp107 engages in two unusual H-bonds, one bet-ween its side-chain atom and the backbone atom ofGly110, the other between its backbone atom and theside-chain atom of Arg162 in the E–F loop. Ala178 islocated in the Asp176ThrAlaGly179 loop turn con-necting the βF strand and the βG strand. Again, theAsp176 residue is involved in several H-bonds withthe turn amino acids. Firstly, an H-bond is foundbetween the Asp176 side-chain atom and the back-bone atom of Gly179. Secondly, two H-bonds are

Fig. 2. Multiple sequence alignment of XC1894 with E. coli FlgD and S. typhimurium FlgD using CLUSTAL W.19

Numbering system based on the XC1894 sequence is shown in the bottom in red. Identical residues are shown in red andboxed, while similar residues are shown in blue and boxed. The first 83 N-terminal residues invisible in the electrondensity map are shown in italics, with the first traceable aspartic residue annotated by a red triangle. The α-helix and β-strands 1, 2, 3, 4, and 5 of the tudor domain are plotted in blue cylinder and arrows, respectively, while the β-strands A, B,C, C′, E, F, and G of the Fn-III domain are plotted in orange arrows. The residues in the hydrophobic core of the tudordomain, Fn-III domain, and interface are marked in red circles, blue circles, and green circles, respectively. This alignmentfigure was generated using ALSCRIPT.20


found between the Asp176 backbone atom and thetwo backbone nitrogen atoms of Ala178 and Gly179.Finally, Ala215 is located in the Pro213LeuAlaAsn216loop connecting the β4 strand and the β5 strand, withthe backbone atom of Pro213 forming two H-bondswith the backbone atoms of Ala215 and Asn216.

XC1894 adopts a novel hybrid comprising atudor-like motif interdigitated with a fibronectintype III (Fn-III) motif

A DALI search using the coordinates of the coreXC1894 against the Protein Data Bank (PDB) return-


ed structures with only partial superimposition. Thefull structure is thus dissected into two separatedomains (Fig. 2), the first one combining segmentsfrom the N-terminal residues Asp84 to Pro100 andthe C-terminal residues from Thr186 to residue

Fig. 3. (a) The stereo pictures of XC1894 monomer crystal stthe N-terminus in blue to the C-terminus in red. The tutor dostrands (1, 2, 3, 4, and 5), while the Fn-III domain comprisesStrand 1 of the tudor domain is followed by strand A of the Fnby strand 2 of the tutor domain, forming a novel hybrid compr(b) The stereo pictures of XC1894 structure showing the consetudor domain are marked in red, those in the Fn-III domaincoloring system is similar to that in Fig. 2. Two salt bridges betwdomain and one H-bond between L222 from the tudor domaconnected by green dotted lines.

Ser221 (colored blue in Fig. 2), and the other fromthe central segment from residue Ser101 to residueAla185 (colored red in Fig. 2). They are searchedagainst PDB individually. Structures similar to thetudor-like domain were obtained from the first

ructure shown in ribbon. The structure is color coded frommain comprises an N-terminal α-helix followed by five β-seven β-strands forming two β-sheets (ABE and C′CEG).-III domain, and strand G of the Fn-III domain is followedising a tudor domain interdigitated with an Fn-III domain.rved hydrophobic residues. The hydrophobic residues inare in blue, and those in the interface are in green. Thiseen R232 from the tudor domain and E161 from the Fn-IIIin and K180 from the Fn-III domain are also shown and


search23–25 while structures similar to the Fn-III do-main were obtained from the second search.26–28 Thefirst domain of XC1894 is found to superimpose verywell with the spinal muscular atrophy tudor domain(1G5V) with an rmsd of 1.09 Å for 28 out of 53 Cα

atoms.23 It also superimposes well to a number ofchromo domain (a member of tudor family that caninteract with specific methylated lysines on his-tones),24,25 such as 1MHN, 2F5K, and 2G3R, withrmsd values of 1.25, 1.35, and 1.39 Å for 35, 41, and 41Cα atoms out of 53 Cα atoms, respectively. Likewise,the second domain of XC1894 superimposes wellwith a number of cell adhesion and heparin-bindingmolecules containing the Fn-III domain,26–28 such as1FNA, 2HAZ, and 2GEE, with rmsd values of 1.38,1.53, and 1.76 Å for 50, 40, and 53 Cα atoms out of 94Cα atoms, respectively. The final structure of themonomer is thus a novel hybrid comprising a tudor-like motif interdigitated with an Fn-III motif asshown in Fig. 3. It is highly enriched in β-strands(66%), as is also detected in other atomic flagellaraxial structures such as filament,11,29 hook,6 and fila-ment–hook junction proteins. Since most of the coreresidues in the tudor domain, the Fn-III domain, andthe interface domain are either identical or highlysimilar among X. campestris, E. coli, and S. typhimur-ium (Fig. 2), one can safely assume that the C-terminaldomain of hook caps of all these bacteria adoptssimilar structures.

The XC1894 tudor domain

The XC1894 tudor domain is rather typical exceptfor some minor change. As in a canonical tudor do-main, it comprises five β-strands forming a stronglybent antiparallel β-sheet. The five β-strands form abarrel-like fold that is padded by a long curved β2strand at the bottom, which is extended from the βGstand of the second Fn-III domain. The first β-strandis also preceded by a two-turn α-helix, not common-ly observed among the tudor domains published sofar. Many highly conserved hydrophobic residues,such as Leu92, Val97, Val192, Leu206, Leu214, andVal217, are present (red dots in Fig. 2) to stabilize thecore structure of the XC1894 tudor domain.

The XC1894 Fn-III domain

The XC1894 Fn-III domain is also rather canonicaland is formed from seven β-strands labeled ABCC′EFG from N to C according to the standard topolo-gical labeling of Ig-like proteins.30 These sevenstrands form two β-sheets that are packed to forma sandwich-like structure enclosing a very hydro-phobic core comprising residues from both sheets.Sheet 1 comprises strandsABE and sheet 2 comprisesstrands C′CFG. The three hairpins between strandsA–B, C–C′, and F–G are connected by regular β-turns, while the other three between strands B–C,C′–E, and E–F are looped across sheet 1 and sheet 2 toform the so-called “crossover” loops. As in the tudordomain, many hydrophobic residues such as Val104,Ile106, Val116, Val124, Phe126, Ile128, Leu138,

Val148, Phe151, Trp153, Ile170, and Ala172 (bluedots in Fig. 2) are located in the structural core region(Fig. 3b). Such a highly hydrophobic core is char-acteristic of an Ig-like domain.31,32 Furthermore, inour hybrid structure comprising a tudor and an Fn-III domain, the hydrophobic core is found to extendfrom the tudor domain to the Fn-III domain throughan interfacial hydrophobic core comprising Val99,Val188, and Leu209 (green dots in Figs. 2 and 3b).Two salt bridges (between the side-chain atom ofArg219 in the tudor domain and the side-chain atomof Glu148 in the Fn-III domain) and one H-bond(between the backbone atom of Leu209 in the tudordomain and the side-chain atom of Lys167 in the Fn-III domain) are also found, which further stabilizethe connection between the tudor and the Fn-III do-mains (Fig. 3b).

Three different interfaces for XC1894 monomerinteractions

Tudor domain is usually found to adopt a tandemversion33,34 or interact with other Sm-like motifthrough β-strand 4 or 5.23 Similarly, Fn-III motifalso exists in tandem repeats but connected bylinker.26,27 The crystal packing of the XC1894 mono-mer showed that XC1894 monomers form threedifferent interfaces, different from all those found ina pure tudor or Fn-III domain, to form a pentagondecamer. As shown in Fig. 4, three types of mono-meric interfaces are observed: type I is formedthrough interactions between residues mainly fromthe Fn-III domain, type II from the Fn-III domain only,and type III from the tudor domain only. In type Iinterface, interactions are largely due to residuessituated in the loop 1–A, loop C–C′, strand F, andstrand G from the Fn-III domain (Fig. 4a). Two H-bonds between the Gln103 side-chain atom of strandA and the backbone atom of Lys183 from strandG arealso observed. Someminor interactions from residuesin the loop 4–5 of the tudor domains are also present.In type II interface, interactions are due only fromresidues situated in strands C and C′ and loops EFand FG of the Fn-III domain (Fig. 4b). Four H-bonds,two between the side-chain atom of Asn125 fromstrand C and the side-chain atom of Ser140 fromstrand C′ and two between the side-chain atom ofAsn159 from loop EF and the backbone atom ofPhe135 from strand C′, are observed. Such interac-tions have never been observed between Fn-IIIdomains published so far,26–28 while in type III inter-face, only residues from the tudor domain contributeto the interactions. Thus, the backbone atoms ofresidues Val195, Thr196, Ile197, and Gly198 fromstrand 2 form H-bonds with the backbone atoms ofresidues Gly198′, Ile197′, Thr196′, and Val195′ fromstrand 2′, respectively, to form a continuous antipar-allel six-stranded β-sheet of 4322′3′4′ topology (Fig.4c). Besides these four backbone H-bonds, one H-bond between the side-chain atoms of Thr196 andThr196′ is also detected. Some minor interactionsderiving from residues situated in the N-terminalresidues are also observed. The β2–β2′ strand inter-

Fig. 4. The three types of interfaces between the XC1894 monomer structures in crystal. (a) Type I interface is formedmainly from residues located in the Fn-III domain, with minor contribution from residues located in the tudor domain.Two H-bonds between the side-chain oxygen atoms of Gln103 of strand A and the backbone nitrogen atoms of Lys183from strand G are shown in ball-and-stick representation and connected by green dotted lines. (b) Type II interface isformed from residues situated in strands C and C′ and loops EF and FG of the Fn-III domain only. Two salt bridges andtwo H-bonds in the interface are shown in ball-and-stick representation and connected by green dotted lines. (c) Type IIIinterface is formed from residues situated in the tudor domain only. Four H-bonds between the backbone atoms ofstrands 2 and 2′ extend the three-stranded 432 β-sheet into a six-stranded 4322′3′4 β-sheet. One extra H-bond between theside-chain atoms of Thr196 is also observed. These H-bonds are connected by green dotted lines.


action in XC1894 has never been observed in anypublished interactions in the tudor domains or Sm-like domains, because in these domains, usually

strands β4 and β5 of one domain interacts withstrands β5′ and β4′ of the neighboring domain conse-cutively to form either a hexamer or a heptamer.35–39


Discussion

From the determined structure, it is clear whythe N-terminus of XC1894 can be cleaved for up to75 amino acids, while the C-terminus remains in-tact. We believe that the main reason is the necessi-ty to maintain an intact tudor domain; the last fiveC-terminal residues of XC1894 comprise strand 5of the tudor domain, and any cleavage from the C-terminus can destabilize the tudor domain to agreat extent, while the N-terminal sequence prece-ding the α-helix is outside the tudor domain andcan be spared without causing its destabilization.Flagellar axial proteins usually contain unstable

terminal sequences.15,40 In fact, both flagellin andhook proteins were found to contain disordered ter-minal regions that span approximately the first 70and the last 30 amino acid residues.15,16,41 The time-course cleavage data for XC1894 are consistent withthe published result, but only the 75 N-terminalamino acids were cleaved; no C-terminal cleavagewas observed. Such results have raised the questionregarding the possible function of terminal disor-dered regions of flagellar axial proteins. The labile N-terminal residues have been found to be the signalsfor exporting17,42 and are necessary for controllingthe self-assembly of flagellin and hook protein.15 Arecent study of FlgD fromE. colidoes indicate that thefirst 71 N-terminal residues are the signal for exportinto the flagellar channel.17 This number is verysimilar to the number of residues that are cleaved inour XcFlgD protein.The crystal structure of the C-terminal domain of

the hook-capping protein adopts a novel hybridstructure comprising a tudor domain and an Fn-IIIdomain, both of which have been found in a numberof protein structures with important biologicalfunctions. For example, the tudor domain is presentin the survival of motor neuron protein that plays acrucial role in the assembly of spliceosomal uridine-rich small nuclear ribonucleoprotein complexes,23 inthe fragile X mental retardation protein family,25 andin the DNA repair proteins recognizing specifichistone lysine methylation for transcriptional regu-lation. The tudor domains always pair by strandsβ4–β5′, never by strands β2–β2′. The Fn-III domainis mainly present in human fibronectin proteins thatcan recognize RGD loop,27 bind heparin and inte-grin,26 or serve as a basic module for extracellularmatrix formation.43 Fn-III domains are linked by alinker and have never been observed to interact di-rectly. In this novel hook cap structure comprising atudor domain and an Fn-III domain, many newinteraction interfaces are found. Since the Fn-III do-main is commonly observed in the eukaryotic king-dom and only rarely observed in the bacterialkingdom, it has been proposed that the Fn-III domainin bacteria is horizontally transferred directly fromthe eukaryotic kingdom.44Currently, three lower-resolution electron micros-

copy pictures regarding the filament cap structure,that is, an isolated cap,45 a cap dimer,46 and a cap–filament complex,46 are available. In the isolated cap

picture, 10 molecules of FliD were found to self-assemble into an annular structure of approximately100 Å in dimension that consists of five substruc-tures arranged in a pentagonal shape.45 In the capdimer picture, two pentamers are arranged in abipolar arrangement. Two characteristic domainsare found in each pentamer: a top plate domain ofpseudo 5-fold symmetry and five bottom leg-likeanchor domains.46 The two pentamer caps are con-nected with a disordered domain at the center. In thecap-filament picture, the leg domains appear to pluginto the cavity of the filament end (probably domainD0 of flagellin) to anchor the cap domain.46 It is alsoproposed that the insertion of a flagellin subunit intothe double indentation of the filament end willrotate the cap domain into the next stable position.Hence, the synergistic interactions between the capdomain, the leg domain, and the incoming flagellinmonomer are necessary to act coordinately to self-assemble a flagellum.46 A similar situation may alsoexist in the hook cap structure, since in crystal, the C-terminal domain of XcFlgD was also found to form apentamer of dimers (data not published). However,since the N-terminal domain of XcFlgD was invi-sible and, in solution, only a dimer was found for theC-terminal domain of XcFlgD, whether XcFlgDadopts a similar structure containing a leg domainand a plate domain of pentamer of dimers remainsto be seen.Interestingly, a genetic study published by Kutsu-

kake and Doi in 1994 claimed that the 86 N-terminalresidues of the S. typhimurium hook cap protein FlgDare sufficient to complement all the flgD mutationsexamined.47 The N-terminal domain of S. typhimur-ium FlgDmay form the leg-like domain as the FliD inthe filament cap, and its interaction with the flagellinsubunit is indeed critical for self-assembling a func-tional flagellum. More experiments are necessary toestablish the roles of the FlgD N-terminal and C-terminal domains in X. campestris.

Materials and Methods

Cloning and purification

The XC1894 gene fragments with 221, 187, 150, and 137amino acid residues were PCR amplified directly from theplant pathogen X. campestris pv. campestris str. 17 (Xcc)using a fixed backward 5′-TTATCCACTTCCAATGT-CAGCTGACGCGGAGCACGTTGGC primer and differ-ent forward primers: (1) 5′-TACTTCCAATCCAATGCT-ATGAGCACGATCGGCAGTGACCTTTA-3′, (2) 5′-TACT-TCCAATCCAATGCTTTGATGACCGAGCAGCTC-CAGC-3 ′ , (3) 5 ′-TACTTCCAATCCAATGCTAC-CAAGGTCGGCAATTTCTCCG-3′, and (4) 5′-TACTT-CCAATCCAATGCTGTGCTCAAGGGTGCCGCGCT-3′,for the 1894(221), 1894(187), 1894(150), and 1894(137)genes, respectively, for forming the fragments of requiredlength. All PCR fragments have correct sizes in an SDS-PAGE experiment and were confirmed by DNA sequen-cing. A ligation-independent cloning approach12,13 wasused to obtain the desired constructs. The final constructscode for an N-terminal His6 tag, a 17-amino-acid linker,and various XC1894 targets under the control of a T7

Table 1. Statistics of data collection and structuralrefinement of Se-Met XC1894

Space group P6122Unit-cell parameters (Å) a=b=116.75, c=71.85Mosaicity (°) 0.52Wavelength (Å)

High remote 0.964215Inflection 0.979467

Resolution range (Å)High remote 30–2.51 (2.61–2.51)Inflection 30–2.55 (2.65–2.55)

Data cutoff (σF)High remote 2.0Inflection 2.0

Number of measured reflectionsHigh remote 132,632 (12,604)Inflection 126,783 (12,023)

Number of unique reflectionsHigh remote 11,527 (1096)Inflection 10,985 (1064)

RedundancyHigh remote 11.5 (11.5)Inflection 11.5 (11.3)

Completeness (%)High remote 99.7 (100)Inflection 99.9 (100)

Rmerge(%)High remote 5.9 (68)Inflection 6.0 (56)

Mean I/σ(I)High remote 33.9 (3.3)Inflection 31.3 (3.9)

Rfree test set size (%) 5R/Rfree (%) 22.5/24.1Solvent content (%) 65Matthew coefficient (Å3/Da) 3.55Average B value

(main-chain/side-chain atoms)59.1/60.4

Model contentProtein residues/Water 137/67

Deviation from ideal geometryBond lengths (Å) 0.009Bond angles (°) 1.400Chiral 0.102

Ramachandran plot (%)a

Residues in most favored regions 93.5Residues in additionally allowedregions

3.6

Values in parentheses are for the outermost shell while thepreceding values refer to all data.

a Four residues (2.9%) are located in the loop turn regions andfound to deviate from the allowed torsional angles (see the maintext for explanation).


promoter. Overexpression of the His6-tag target proteinswere induced by the addition of 0.5 mM IPTG at 293 K for20 h. The XC1894(137) protein is not stable enough to becollected during purification. Other three target proteinswere purified by immobilized metal affinity chromato-graphy on a nickel column (Sigma). TheHis6 tag and linkerwere cleaved from XC1894 by tobacco etch virus proteaseat 289 K for 20 h. For crystallization, XC1894 proteins ofdifferent length were further purified on an anion ex-changer column (AKTA, Pharmacia Inc.). The final freshtarget proteins exhibit purities greater than 99% andcontain only an extra tripeptide (SNA) at the N-terminalend. However, they experienced cleavage from the N-ter-minus; thus, only fresh samples were used for crystal-lization screening. Se-Met-labeled XC1894was prepared ina similar way, except that it was produced using a non-auxotroph E. coli strain BL21 (DE3) as host in the absence ofmethionine but with ample amounts of Se-Met (100 mg/l).The M9 medium consists of 1 g of NH4Cl, 3 g of KH2PO4,and 6 g of Na2HPO4 supplemented with 20% (W/V) ofglucose, 0.3% (W/V) ofMgSO4, and 10mgof FeSO4 in 1 l ofdouble-distilled water. The induction was conducted at289 K for 24 h by the addition of 0.8 ml of 0.5 mM IPTG.Purification of the Se-Met-labeled XC1894 protein wasperformed using the protocols as established for the nativeproteins.

Effects of protease inhibitors on FlgD cleavage

The time-course cleavage experiments of Xcc FlgD wereperformed in the absence or presence of protease inhibitorcocktail. The inhibitor cocktail comprises AEBSF [4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride] forinhibiting serine proteases such as trypsin, chymotrypsin,and so forth; bestatin hydrochloride for inhibitingaminopeptidases such as leucine aminopeptidase andalanyl aminopeptidase; E64 [N-(trans-epoxysuccinyl)-L-leucine 4-guanidinobutylamide] for inhibiting cysteineproteases such as calpain, cathepsin B, and so forth;ethylenediaminetetraacetic acid for inhibiting metallopro-teases; pepstatin A for inhibiting acid proteases such aspepsin, rennin, and many microbial aspartic proteases;and PMSF inhibitor (Sigma). The fresh Xcc FlgD proteinsof various lengths were incubated at 25 °C for theindicated time. The inhibitor cocktail was applied at therecommended concentration of 0.2 g/ml. The cleavageexperiment in the presence of protease inhibitor cocktailwas performed at 37 °C for 12 h.

Crystallization

For crystallization, the native protein was concentratedto 25 mg/ml in 20 mM sodium chloride and 80 mM Tris,pH 8.0, using an Amicon Ultra-10 (Millipore). Screeningfor crystallization condition was performed using sitting-drop vapor diffusion in 96-well plates (HamptonResearch)at 297 K by mixing 0.5 μl protein solution with 0.5 μlreagent solution. Initial screens including the Hamptonsparse matrix Crystal Screens 1 and 2, a systematic PEG(polyethylene glycol)–pH screen, and a PEG/Ion screenwere performed using the Gilson C240 crystallizationworkstation. Hexagonal-like crystals appeared in 14 daysfrom a reservoir solution comprising 1.7 M sodium acetateand 0.1 M sodium cacodylate. Crystals suitable fordiffraction experiments were grown by mixing 1.5 μlprotein solution with 1.5 μl reagent solution at room tem-perature and reached dimensions of 0.1 mm×0.1 mm×0.3 mm. Se-Met-labeled XC1894 was crystallized in the

same way, but under different crystallization conditions;similar hexagonal-like crystals appeared in 14 days from areservoir solution comprising 2.5M sodium chloride, 0.1MTris (pH 7.0) buffer, and 0.2 M magnesium chloride.

Data collection

Crystals were flash-cooled at 100 K under a stream of coldnitrogen. X-ray diffraction data were collected using theNational Synchrotron Radiation Research Center (NSRRC)beamline 13B1 in Taiwan.However, native Xcc FlgD crystalssuffered extensive radiation damage during data collection,and the resulting diffraction data are not sufficient forstructural determination. Only the diffraction data collectedat twodifferent wavelengths (high remote and inflection) onthe Se-Met-substituted crystals were found to be of good


quality (up to ca 2.50 Å resolution) for the phase andstructural determination. The data were indexed andintegrated using HKL2000 processing software,48 givingdata sets that were approximately 99% complete with anoverall Rmerge of 5.9–6.6% on intensities. The refinement ofselenium atom positions, phase calculation, and densitymodification were performed using the program SOLVE/RESOLVE.49 The model was manually adjusted using theXtalView/Xfit package. CNS50was then used for refinementto a final Rcryst of 22.5% and Rfree of 24.1%. The crystalsbelong to the P6122 space group. The data and refinementstatistics are summarized in Table 1.

Accession number

The coordinates of the XC1894 monomer and itsstructure factors have been deposited in the PDB (accessionnumber: 3C12).

Acknowledgements

This work is supported by the Academic Excel-lence Pursuit grant from the Ministry of Educationand by the National Science Council, Taiwan, ROC(grants 94-2113-M005-003 and 95-2113-M005-018) toS.-H. Chou. We thank the Core Facilities for ProteinX-ray Crystallography in the Academia Sinica,Taiwan for help in crystal screening, the NationalSynchrotron Radiation Research Center (NSRRC) inTaiwan, and the SPring-8 Synchrotron facility inJapan for assistance of X-ray data collection. TheNSRRC is a user facility supported by the NationalScience Council, Taiwan, ROC, and the ProteinCrystallography Facility is supported by theNational Research Program for Genomic Medicine,Taiwan, ROC.

References

1. Macnab, R. M. (1999). The bacterial flagellum: re-versible rotary propellor and type III export appara-tus. J. Bacteriol. 181, 7149–7153.

2. Pallen, M. J., Penn, C. W. & Chaudhuri, R. R. (2005).Bacterial flagellar diversity in the post-genomic era.Trends Microbiol. 13, 143–149.

3. Macnab, R. M. (2003). How bacteria assemble flagella.Annu. Rev. Microbiol. 57, 77–100.

4. Lowe, G., Meiser, M. & Berg, H. C. (1987). Rapidrotation of flagellar bundles in swimming bacteria.Nature, 325, 637–640.

5. Samatey, F.A.,Matsunami,H., Imada, K.,Nagashima, S.,Shaikh, T. R., Thomas, D. R. et al. (2004). Structure of thebacterial flagellar hook and implication for themolecularuniversal joint mechanism. Nature, 431, 1062–1068.

6. Shaikh, T. R., Thomas, D. R., Chen, J. Z., Samatey, F. A.,Matsunami, H., Imada, K. et al. (2005). A partial atomicstructure for the flagellar hook of Salmonella typhimu-rium. Proc. Natl Acad. Sci. USA, 102, 1023–1028.

7. Ohnishi, K., Ohto, Y., Aizawa, S.-I., Macnab, R. M. &Iino, T. (1994). FlgD is a scaffolding protein neededfor flagellar hook assembly in Salmonella typhimu-rium. J. Bacteriol. 176, 2272–2281.

8. Moriya, N., Minamino, T., Hughes, K. T., Macnab,R. M. & Namba, K. (2006). The type III flagellarexport specificity switch is dependent on FliK rulerand a molecular clock. J. Mol. Biol. 359, 466–477.

9. Waters, R. C., O'Toole, P. W. & Ryan, K. A. (2007). TheFliK protein and flagellar hook-length control. ProteinSci. 16, 769–780.

10. Minamino, T., Saijo-Hamano, Y., Furukawa, Y., Gonza-lez-Pedrajo, B., Macnab, R. M. & Namba, K. (2004).Domain organization and function of Salmonella FliK, aflagellar hook-length control protein. J. Mol. Biol. 341,491–502.

11. Yonekura, K., Maki-Yonekura, S. & Namba, K. (2003).Complete atomic model of the bacterial flagellar fila-ment by electron cryomicroscopy. Nature, 424, 643–650.

12. Stols, L., Gu,M., Dieckman, L., Raffen, R., Collart, F. R.& Donnelly, M. I. (2001). A new vector for high-throughput, ligation-independent cloning encoding atobacco etch virus protease cleavage site. ProteinExpression Purif. 25, 8–15.

13. Aslanidis, C. & de Jong, P. J. (1990). Ligation-independent cloning of PCR products (LIC-PCR).Nucleic Acids Res. 18, 6069–6074.

14. Saijo-Hamano, Y., Namba, K. & Oosawa, K. (2000). Anew purification method for overproduced proteinssensitive to endogenous proteases. J. Struct. Biol. 132,142–146.

15. Vonderviszt, F., Ishima, R., Akasaka, K. &Aizawa, S.-I.(1992). Terminal disorder: a common structural featureof the axial proteins of bacterial flagellum? J. Mol. Biol.226, 575–579.

16. Vonderviszt, F., Kanto, S., Aizawa, S. I. & Namba, K.(1989). Terminal regions of flagellin are disordered insolution. J. Mol. Biol. 209, 127–133.

17. Weber-Sparenberg, C., Poplau, P., Brookman, H.,Rochon, M., Mockel, C., Nietschke, M. & Jung, H.(2006). Characterization of the type III export signal ofthe flagellar hook scaffolding protein FlgD of Esche-richia coli. Arch. Microbiol. 186, 307–316.

18. Kagawa, H., Aizawa, S. I., Yamaguchi, S. & Ishizu, J. I.(1979). Isolation and characterization of bacterialflagellar hook proteins from salmonellae and Esche-richia coli. J. Bacteriol. 138, 235–240.

19. Thompson, J. D., Higgins, D. G. & Gibson, T. J. (1994).CLUSTALW: improving the sensitivity of progressivemultiple sequence alignment through sequenceweighting, position specific gap penalties and weightmatrix choice. Nucleic Acids Res. 22, 4673–4680.

20. Barton, G. J. (1993). ALSCRIPT: a tool to format mul-tiple sequence alignments. Protein Eng. 6, 37–40.

21. Hu, R.-M., Yang, T.-C., Yang, S.-H. & Tseng, Y.-H.(2005). Deduction of upstream sequences of Xantho-monas campestris flagellar genes responding to trans-cription activation by FleQ. Biochem. Biophys. Res.Commun. 335, 1035–1043.

22. Guex, N. & Peitsch, M. C. (1997). SWISS-MODEL andthe Swiss-PdbViewer: an environment for compara-tive protein modeling. Electrophoresis, 18, 2714–2723.

23. Selenko, P., Sprangers, R., Stier, G., Buhler, D., Fischer,U. & Sattler, M. (2001). SMN tudor domain structureand its interaction with the Sm proteins. Nat. Struct.Biol. 8, 27–31.

24. Corsini, L. & Sattler, M. (2007). Tudor hooks up withDNA repair. Nat. Struct. Biol. 14, 98–99.

25. Maurer-stroh, S., Dickens, N. J., Hughes-Davies, L.,Kouzarides, T., Eisenhaber, F. & Ponting, C. P. (2003).The Tudor domain ‘Royal family’: Tudor, plant Age-net, Chromo, PWWP, and MBT domains. Trends Bio-chem. Sci. 28, 69–74.


26. Sharma, A., Askari, J. A., Humphries, M. J., Jones, E. Y.& Stuart, D. I. (1999). Crystal structure of a heparin-and integrin-binding segment of human fibronectin.EMBO J. 18, 1468–1479.

27. Leahy, D. J., Aukhil, I. & Erickson, H. P. (1996). 2.0 Åcrystal structure of a four-domain segment of humanfibronectin encompassing the RGD loop and synergyregion. Cell, 84, 155–164.

28. Dickinson, C. D., Veerapandian, B., Dai, X.-P., Hamlin,R. C., Xuong, N.-H., Ruoslahti, E. & Ely, K. R. (1994).Crystal structure of the tenth type III cell adhesion mo-dule of human fibronectin. J. Mol. Biol. 236, 1079–1092.

29. Samatey, F. A., Imada, K., Nagashima, S., Vonder-viszt, F., Kumasaka, T., Yamamoto, M. & Namba, K.(2001). Structure of the bacterial flagellar protofila-ment and implications for a switch for supercoiling.Nature, 410, 331–337.

30. Williams, A. F. & Barclay, A. N. (1988). The immuno-globulin superfamily: domains for cell surface recog-nition. Annu. Rev. Immunol. 6, 381–405.

31. Hamill, S. J., Steward, A. & Clarke, J. (2000). Thefolding of an immunoglobulin-like Greek key proteinis defined by a common-core nucleus and regionsconstrained by topology. J. Mol. Biol. 297, 165–178.

32. Lappalainen, I., Hurley, M. G. & Clarke, J. (2008).Plasticity within the obligatory folding nucleus of animmunoglobulin-like domain. J. Mol. Biol. 375, 547–559.

33. Botuyan, M. V., Lee, J., Ward, I. M., Kim, J.-E.,Thompson, J. R., Chen, J. Z. & Mer, G. (2006). Struc-tural basis for the methylation state-specific recogni-tion of histone H4-K20 by 53BP1 and Crb2 in DNArepair. Cell, 127, 1361–1373.

34. Huang, Y., Fang, J., Bedford, M. T., Zhang, Y. & Xu,R.-M. (2006). Recognition of histone H3 lysine-4methylation by the double tudor domain of JMJD2A.Science, 312, 748–751.

35. Collins, B. M., Harrop, S. J., Kornfeld, G. D., Dawes,I. W., Curmi, P. M. G. & Mabbutt, B. C. (2001). Crystalstructure of a heptameric Sm-like protein complexfrom archaea: implications for the structure andevolution of snRNPs. J. Mol. Biol. 309, 915–923.

36. Sauter, C., Basquin, J. & Suck,D. (2003). Sm-like proteinsin Eubacteria: the crystal structure of the Hfq proteinfrom Escherichia coli. Nucleic Acids Res. 31, 4091–4098.

37. Schumacher, M. A., Perarson, R. F., Moller, T.,Valentin-Hansen, P. & Brennan, R. G. (2002). Struc-tures of the pleiotropic translational regulator Hfq andan Hfq–RNA complex: a bacterial Sm-like protein.EMBO J. 21, 3546–3556.

38. Pannone, B. K. & Wolin, S. L. (2000). RNA degrada-tion: Sm-like proteins wRING the neck of mRNA.Curr. Biol. 10, R478–R481.

39. Seraphin, B. (1995). Sm and Sm-like proteins belong toa large family: identification of proteins of the U6 aswell as the U1, U2, U4 and U5 snRNPs. EMBO J. 14,2089–2098.

40. Namba, K. (2001). Roles of partly unfolded conforma-tions in macromolecular self-assembly. Genes Cells, 6,1–12.

41. Vonderviszt, F., Aizawa, S.-I. & Namba, K. (1991). Roleof thedisordered terminal regions of flagellin in filamentformation and stability. J. Mol. Biol. 221, 1461–1474.

42. Kuwajima, G., Kawagishi, I., Homma, M., Asaka, J.-I.,Kondo, E. & Macnab, R. M. (1989). Export of an N-terminal fragment of Escherichia coli flagellin by aflagellum-specific pathway. Proc. Natl Acad. Sci. USA,86, 4953–4957.

43. Lundell, A., Olin, A. I., Morgelin, M., al-Karadaghi, S.,Aspberg, A. & Logan, D. T. (2004). Structural basis forinteractions between tenascins and lectican C-typelectin domains: evidence for a crosslinking role fortenascins. Structure, 12, 1495–1506.

44. Bork, P. & Doolittle, R. F. (1992). Proposed acquisitionof an animal protein domain by bacteria. Proc. NatlAcad. Sci. USA, 89, 8990–8994.

45. Ikeda, T., Oosawa, K. & Hotani, H. (1996). Self-assembly of the filament capping protein, FliD, ofbacterial flagella into an annular structure. J. Mol. Biol.259, 679–686.

46. Yonekura, K., Maki, S., Morgan, D. G., DeRosier, D. J.,Vonderviszt, F., Imada, K. & Namba, K. (2000). Thebacterial flagellar cap as the rotary promoter offlagellin self-assembly. Science, 290, 2148–2152.

47. Kutsukake, K. & Doi, H. (1994). Nucleotide sequenceof the flgD gene of Salmonella typhimurium which isessential for flagellar hook formation. Biochim. Bio-phys. Acta, 1218, 443–446.

48. Otwinowski, Z. & Minor, W. (1997). Processing of theX-ray diffraction data collected in oscillation mode.Methods Enzymol. 276, 307–326.

49. Terwilliger, T. C. & Berendzen, J. (1999). AutomatedMAD and MIR structure solution. Acta Crystallogr.,Sect. D: Biol. Crystallogr. 55, 849–861.

50. Brunger, A. T., Adams, P. D., Clore, G. M., DeLano,W. L., Gros, P., Grosse-Kunstleve, R. W. et al. (1998).Crystallography & NMR system: a new softwaresuite for macromolecular structure determination.Acta Crystallogr., Sect. D: Biol. Crystallogr. 54, 905–921.

crystal structure of the c-terminal domain of a flagellar hook-capping protein from xanthomonas...

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