JOURNAL OF BACTERIOLOGY, Sept. 1986, p. 968-9740021-9193/86/090968-07$02.00/0Copyright © 1986, American Society for Microbiology

Vol. 167, No. 3

Helical Structure of Bordetella pertussis FimbriaeALASDAIR C. STEVEN,'* MARGARET E. BISHER,1 BENES L. TRUS,2 DANIEL THOMAS,1 JIA MING ZHANG,3

AND JAMES L. COWELL3Laboratory of Cellular and Developmental Biology, National Institute of Diabetes and Digestive and Kidney Diseases,1Computer Systems Laboratory, Division of Computer Research Technology, National Institutes of Health,2 and Division

of Bacterial Products, Center for Drugs and Biologics, Food and Drug Administration,3 Bethesda, Maryland 20892

Received 20 December 1985/Accepted 13 May 1986

The helical structures of Bordetella pertussis fimbriae of serotypes 2 and 6 were determined by opticaldiffraction analysis of electron micrographs of negatively stained paracrystalline bundles of purified fimbriae.The fimbrial structure is based on an axial repeat of 13 nm that contains five repeating units in two completeturns of a single-start helix. This structure was confirmed by direct measurements of mass per unit length forindividual fimbriae performed by dark-field scanning transmission electron microscopy of unstained speci-mens. These data further established that the helically repeating unit is a monomer of fimbrial protein (Mr22,000 for type 2 and Mr = 21,500 for type 6). Radial density profiles calculated from the scanning transmissionelectron micrographs showed that the fimbria has peak density at its center, i.e., no axial channel, consistentwith the results of conventional negative-staining electron microscopy. The radial profile gives an outermostdiameter of -7.5 nm, although the peripheral density is, on average, diffuse, aUlowing sufficient intercalationbetween adjacent fimbriae to give a center-to-center spacing of -5.5 nm in the paracrystals. Despite serologicaland biochemical differences between type 2 and type 6 fimbriae, the packing arrangements of their fimbrialsubunits are identical. From this observation, we infer that the respective subunits may have in commonconserved regions whose packing dictates the helical geometry of the fimbria. It is plausible that a similarmechanism may underlie the phenomenon of phase variations in other systems of bacterial fimbriae.

Bacterial fimbriae (otherwise known as pili) are filamen-tous extracellular appendages composed of polymerizedprotein subunits. Several putative functions including rolesin motility have been ascribed to fimbriae from varioussources (13). However, recent attention has focused on theirproperties as adhesins that attach the bacteria to specificsites on susceptible cells to mediate infection. Similarly,fimbriae have attracted considerable interest as potentialvaccines on the premise that antifimbrial antibodies mayprevent effective adhesion and thus thwart the infectionprocess. At least six morphological classes of fimbriae havebeen distinguished (3), and molecular models have beenproposed for type I fimbriae of Escherichia coli (3), for theepisome-encoded F-pili that mediate conjugative transfer ofDNA between compatible strains of E. coli (5), and for thefimbriae of Pseudomonas aeruginosa (6, 24). However, themorphological diversity of fimbriae has yet to be fullycharacterized, as have the functional distinctions that thesestructural variations presumably reflect.

Recently, we have reported the purification of fimbriaefrom Bordetella pertussis 325 (26). These fimbriae werefound to consist of a single protein species with an apparentmolecular weight of -22,000 as determined by sodiumdodecyl sulfate-polyacrylamide gel electrophoresis, and tohave antigenic activity corresponding to a serotype 2 ag-glutinogen of B. pertussis. The purification method wasbased on pH-dependent precipitation of the fimbriae aftertheir detachment from cells by mechanical shearing. Elec-tron microscopy of negatively stained samples showed thatfimbriae in the dispersed state (i.e., at pH -9) are narrow(-5 to 6 nm in width) flexuous filaments (26). On the otherhand, precipitated fimbriae (i.e., at pH -6) were found toform well-aligned paracrystalline bundles of relatively

* Corresponding author.

straight fimbriae. Here we report an analysis of negativelystained paracrystals by optical diffraction and computerimage processing methods. These paracrystals yielddiffractograms that are sufficiently detailed to allow a con-clusive solution for the helical surface lattice on whichsubunits are packed. This structure was confirmed by directmeasurements of mass per unit length of individual fimbriaeperformed by analyzing dark-field scanning transmissionelectron micrographs (STEM) of unstained specimens.

In addition, we applied a modified purification scheme toobtain fimbriae of serotype 6 from B. pertussis. Despitedifferences from serotype 2 fimbriae with respect to molec-ular weight, antigenic specificity, and propensity to aggre-gate as a function of pH (J. L. Cowell, J. M. Zhang and A.Urisu, manuscript in preparation), similar electron micro-scopic analyses showed their supramolecular structure to beindistinguishable from that of serotype 2 fimbriae.

MATERIALS AND METHODSGrowth of cells for isolation of fimbriae. B. pertussis 325

(serotype 1.2.4) was used for the isolation of type 2 fimbriae,and B. pertussis 114 (serotype 1.3.6) was used for theisolation of type 6 fimbriae. The bacteria were grown inliquid culture with shaking as previously described (26),except that modified Stainer-Scholte medium (7) was usedfor the growth of strain 114.

Isolation and purification of fimbriae. Type 2 fimbriae weredetached by mechanical shearing and purified by successiveprecipitations with ammonium sulfate, phosphate buffer (pH6.0), and magnesium chloride as described previously (26).For the isolation and purification of type 6 fimbriae, all stepswere conducted at 4 to 8°C. Bacteria were collected bycentrifugation for 30 min at 5,000 x g. The bacterial pelletwas suspended to 20% (wt/vol) in 0.125 M ethanolamine (pH10.5) and blended in a Sorvall Omni-mixer as describedpreviously (26). Ethanolamine at pH 10.5 was used to keep

968

STRUCTURE OF BORDETELLA PERTUSSIS FIMBRIAE 969

c d

50nmFIG. 1. Electron micrographs of B. pertussis fimbriae negatively stained with 1% uranyl acetate. (a) Serotype 2 fimbriae in the dispersed

state (adsorbed to substrate at pH 9.5); (b) serotype 6 fimbriae in the dispersed state (pH 10.5); (c) serotype 2 fimbriae in the paracrystallinestate (pH 6.0); (d) serotype 6 fimbriae in the paracrystalline state (pH 7.5). Optical diffraction patterns of the areas windowed in panels c andd are shown in Fig. 2b and c, respectively.

the fimbriae in a nonaggregrated suspension. Cells werepelleted twice by centrifugation for 20 min at 10,000 x g. Thesupernatant was precipitated over 30 min with 1.76 g ofammonium sulfate per 10 ml. After centrifugation (27,000 xg for 20 min), the pellet was suspended in 0.125 M ethanol-amine (pH 10.5) with 0.1 of the volume of the supernatantand, after blending, was dialyzed for 16 h against 1,000volumes of the same buffer. Subsequent centrifugationswere done at 27,000 x g for 20 min. After dialysis, insolublematerial was removed by centrifugation. The supernatantwas adjusted to pH 7.4 and dialyzed for 16 h against 1,000volumes of 0.01 M sodium phosphate-buffered saline, pH7.4. The precipitate was collected by centrifugation, sus-pended in 4 M urea in 0.05 M Tris buffer (pH 7.0) with 0.1 ofthe volume of the original supernatant and stirred for 16 h at8°C. Urea was included to solubilize contaminants. Aftercentrifugation, the pellet was again extracted with 4 M ureaas described above. The fimbriae were collected by centrif-ugation and solubilized with -3 ml of 0.125 M ethanolamine(pH 10.5) by stirring for 2 h. The solubilized fimbriae were

dialyzed against 0.05 M Tris hydrochloride buffer (pH 8.5)containing 0.8% NaCl and stored at -20°C.

Conventional transmission electron microscopy. Specimenswere observed in a Philips EM400T electron microscopeoperating at 80 kV, with a liquid-nitrogen anticontaminationdevice in routine use. Magnifications, nominally x36,000 orx46,000, were calibrated relative to the 2.49-nm periodicityof crystallites of the dye olive-T (8). Negative staining wasdone by standard procedures (see, e.g., reference 19). Al-though a number of different stains including phosphotung-stic acid and ammonium molybdate were used, the mostsatisfactory results (as judged by optical diffraction), and allthose reported in this communication, were obtained with1% uranyl acetate.STEM. All STEM and attendant specimen preparation

procedures were carried out at the STEM facility operatedas a National Institutes of Health Biotechnology Resource atBrookhaven National Laboratory (23). After adsorption to athin carbon film substrate, the fimbriae were washed andfreeze-dried as described previously (9, 20). Images were

VOL. 167, 1986

970 STEVEN ET AL.

d

,.. N M .

FIG. 2. Optical diffraction patterns from iegatively stained paracrystalline bundles of B. pertussis fimbriae. Panel a is from a large areaof a serotype 2 paracrystal, and panels b and c derive from smaller, well-ordered areas of paracrystals of fimbriae of serotypes 2 and 6,respectively. The patterns exhibit systems of equally spaced layer lines (I = 1 to 5), ofwhich the fifth has a meridional reflection correspondingto an axial spacing of 2.6 nm. The weaker layer lines, I = 1 and 4, are discernible in panels b and c only. Panels d and e show, at half-scale,computed diffraction patterns of model helices which include five repeating units in two turns and in one turn of the helix, respectively. Thearrowhead in panel a indicates the equatorial reflection whose spacing is a measure of the center-to-center spacing of the packed fimbriae.

recorded directly in digital form with sampling by the fo-cused electron probe (diameter, -0.25 nm) at intervals of 0.5or 1.0 nm. Mass-per-unit-length analyses were performed asdescribed previously (21), working with the dark-field signalrecorded by the high-angle annular detector (9, 20). Simi-larly, radial reconstructions (17) were done with the PICprogram (22) implemented on a VAX 11/780 computer andaccompanying image-processing hardware.

Optical diffraction and image processing. Images (originalelectron micrograph negatives) were analyzed in a converg-ing-beam diffractometer (19) calibrated with a 200-lines-per-inch grating.Computations of the projection images of model helices

and calculations of their diffraction patterns were performedwith the PIC program (22), incorporating some routines fromthe earlier MDPP system (14).

REStJLTSpH-dependent paracrystallization of fimbriae. At pH 9 or

above, serotype 2 fimbriae of B. pertussis are dispersed insolution. Negative-staining electron microscopy revealedsinuous, rather thin filaments (5 to 6 nm in width) of variablelength (Fig. la) which show no evidence of a hollow (stain-penetrable) axial channel. Apart from occasional indicationsof penrpheral serrations (Fig. la), these images reveal fewimmediate indications of substructure. On account of theirlow contrast and also because of their pronounced curva-ture, such images of individual fimbriae are not well suitedfor diffraction analysis. At pH 6 or below, paracrystallineaggregation of fimbriae takes place (Fig. lc), in which lateralpacking produces bundles of aligned fimbriae. With fimbriaeof serotype 6 (Fig. lb and d), essentially the same phenom-enon is observed except that the transitional pH value forparacrystallization (-7.5) is significantly higher. In theparacrystalline formations, the fimbriae are relativelystraight, and these bundles exhibit conspicuous striationsupon negative staining; as such, they are readily amenable toanalysis by optical diffraction.

Optical diffraction analysis. Diffraction patterns from largeareas of paracrystals (e.g., Fig. 2a) typically show thefollowing intense but somewhat diffuse features: ameridional reflection at 0.38 nm-1 correspotiding to an axial

repeat of 2.6 ± 0.1 nm and layer lines at meridional spacingsof 0.15 and 0.23 nm-1. These spacings are, respectively,fractions of 2/5 and 3/5 that of the meridional reflection. Thefirst of these (at 0.15 nm'1) is invariably the stronger of thetwo. These reflections tend to be relatively diffuse becauseof disorder within large paracrystals. Sharper diffractionpatterns from relatively small, but more coherent, areas ofparacrystal (Fig. 2b) show, albeit faintly, two additionallayer lines at spacings of 1/5 and 4/5 that of the meridionalreflection. The patterns generated by paracrystals ofserotype 6 fimbriae show exactly the same features (Fig. 2c).These diffractograms exhibit five equally spaced layer

lines (I = 1 to 5), of which the fifth is a meridional.Accordingly, they represent a structure that has an exactaxial repeat of 13 nm, containing five helical steps. The onlyremaining ambiguity in interpretation of the pattern (apartfrom handedness of the helix) is whether the surface latticeis such that the 13-nm axial repeat contains one or two turnsof the basic helix. (Structures with three and four turns peraxial repeat are, respectively, the same as two- and one-turnstructures apart from an inversion of handedness.) Comput-er-generated diffraction patterns corresponding to both alter-natives are shown in Fig. 2d and e. With one turn per 13-nmpitch (Fig. 2e), the reflections on layer-lines 2 and 3 aresituated further from the meridian than those of layer-lines 1and 4, and vice versa for the two-turn model (Fig. 2d).Referring to Fig. 2b and c, it is clear that the experimentalpatterns conform to the latter structure (cf. Fig. 2d).Thus, the surface lattice of B. pertussis fimbriae is a

single-start helix of 6.5-nm pitch with 2.5 repeating units perturn. A radial projection of this surface lattice is drawn inFig. 3a, and its cylindrical folding is represented in tPig. 3b.It is noteworthy (see Discussion) that the strongest feature ofthe paracrystal diffractograms is invariably the second layer-line. This reflection is a first-order Bessel function: it corre-sponds to a single-start helix of 6.5-nm pitch and is presum-ably generated by a set of relatively prominent stain-penetrable helical grooves on the fimbrial surface.

Determination of fimbrial mass per unit length by STEM.The biochemical evidence (1, 26) has shown that B. pertussisfimbriae contain a single species of protein subunit, with anestimated Mr of 22,000 in the case of serotype 2. Thehelically repeating unit should contain an integral number of

J. BACTERIOL.

STRUCTURE OF BORDETELLA PERTUSSIS FIMBRIAE 971

-.0.l ..3

-

-O

0 ,.,

0

0

0 /

c ;

\,7

.j_

FIG. 3. Schematic drawings representing the helical surface lattice of B. pertussis fimbriae; the handedness of the helix has not yet beendetermined, and an arbitrarily chosen right-handed structure is shown here. (a) Radial projection of the surface lattice. The set of lattice linesdrawn marks the single-start helix that contributes the prominent layer line (1 = 2) in the diffraction patterns (Fig. 2). (b) Helical structuregenerated by cylindrical closure of the surface lattice. (c) Model of fimbrial structure based on the determined surface lattice and radialdistribution of density.

copies of this protein. To confirm our deduced surfacelattice, and also to determine whether the repeating unit is amonomer, a dimer, or a trimer, etc., we determined the massper unit length of type 2 fimbriae by quantitative STEM (9,20, 23). The dark-field mode of this microscope gives a signal

that is proportional to the mass in each portion of specimensampled by the STEM electron probe. Appropriate two-dimensional integration of the density distribution recordedin such images therefore affords a direct determination ofmass per unit length for individual particles (20, 21, 23). A

0a,

._

.E 1'co

c

.0

E

zo

0

Mass-per-Unit-Length (kDa/nm)

"00to" ID O . iE0

"' 0

c o

c 5_w _%o co

0.2>~._

;0~*_ _

In000.-

5 nm 0 1 2 3 e 5Radius (nm)

FIG. 4. (a) Dark-field STEM image of unstained fimbriae prepared by freeze-drying; also shown is a calibration particle of tobacco mosaicvirus (T). (b) Histogram of average mass-per-unit-length values measured for 43 individual fimbriae from such STEM images. (c) Radial profileof cylindrically averaged (dry) density in fimbriae, also calculated directly from these micrographs. Experimental uncertainty in such profilesis greatest close to the axis (17); accordingly, the apparent blip of density at the axis (r < 0.4 nm) should be interpreted with caution.Nevertheless, we feel confident that these data demonstrate the absence of an axial channel on any significant scale.

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972 STEVEN ET AL.

STEM dark-field image of unstained fimbriae prepared byfreeze-drying is shown in Fig. 4a. The global average lineardensity was 8.90 kilodaltons (kDa)/nm for measurementsmade on 43 fimbriae amounting to a cumulative length of8.65 ,um. The standard deviation of the set of measurementsfor the individual fimbriae was 0.77 kDalnm, and the stan-dard error of their mean was 0.11 kDalnm. These data arehistogrammed in Fig. 4b. The determined value is in goodagreement with the value of 8.46 kDalnm predicted from thesurface lattice and a subunit molecular weight of 22,000,although it should be noted that errors at the level of 10 to15% at least may be incurred in molecular weight valuesestimated by sodium dodecyl sulfate-polyacrylamide gelelectrophoresis (25). The observed agreement corroboratesour deduced surface lattice and establishes that the repeatingunit is a monomer of fimbrial protein.Diameter and radial density profile of fimbriae. In addition

to quantitating the average linear density along a filament,the STEM data can also be used to map the average densitydistribution in the radial dimension (17). The transverseprofile of projected density averaged over short straightpieces of fimbria was calculated, and the radial densityprofile calculated from this projection is given in Fig. 4c. Itshows the highest density at the center, with density taperingoff to an outer radial limit of -3.8 nm. This corresponds to adiameter somewhat in excess of the center-to-center spacing

\ / \ - D -,

Wa-t \{o _ _-atO\ B

FIG. 5. Diagram illustration of how layered aggregates may beformed as crystals of intercrossing fimbriae which have the helicalstructure determined in this study. The prominent ridges of fimbriaA are parallel to and fit snugly into the corresponding grooves offimbria B when the two are fitted together at a crossing angle (2a) of-40° (see Discussion). For instance, ridge cc' on the upper side offimbria B fits into the groove between ridges aa' and bb' on the lowerside of fimbria A. An identical contact is made with B by filamentA', not only in terms of groove/ridge fitting but also in terms of theprecise configuration of the subunits that make contact between therespective fimbriae, because A' is parallel to A and is displacedalong B by one complete axial repeat (D) of 13 nm. The arrangementshown gives a maximum amount of groove/ridge overlap; arrange-ments that have identical lattice parameters, but which would bedifferent at the molecular level (although indistinguishable in termsof currently available information), may be generated by slidingfimbriae A, A', etc., along the direction c'c. For instance, such ashift by -3 nm would generate an equivalent arrangement withfimbriae A and B related by a two-fold screw axis in the midplane,oriented in a direction that bisects their crossing angle.

of 5.5 nm measured from the paracrystals and indicates thatwhen they are packed into the paracrystals, there is appre-ciable intercalation between neighboring fimbriae. As calcu-lated from the STEM density profile, only 19% of the totalfimbrial mass is located at radii greater than 2.7 nm.

DISCUSSION

Helical geometry of B. pertussis fimbriae. From opticaldiffraction analysis of electron micrographs of negativelystained paracrystals, we determined that the packing ofsubunits in these fimbriae conforms to helical symmetry,based on a single-start helix of 6.5-nm pitch with 2.5 repeat-ing units per turn. This structure was confirmed by directmeasurements of mass per unit length of individual fimbriaeperformed by analyzing STEM images of unstained speci-mens. These data fulfilled the linear density predicted by thededuced helical parameters, given the molecular weight ofthe fimbrial protein subunit (22,000), and established that therepeating unit is a monomer of this protein.The center-to-center spacing of adjacent fimbriae in the

paracrystal was measured by diffraction to be -5.5 nm, andthis figure is consistent with the apparent projected widths ofindividual negatively stained fimbriae (5 to 6 nm). However,radial density profiles derived from the unstained STEMdata indicate an outer diameter of -7.5 nm. To reconcile thisdisparity, we infer that, in the paracrystals, there is interca-lation between the peripheries of adjacent fimbriae so thatthe center-to-center spacing in the paracrystal is consider-ably less than the outer diameter of the fimbria. Most likely,the slightly narrower appearance (5 to 6 nm) of individualfimbriae in negative stain is the result of coincidental positivestaining of their peripheries or of ineffectual exclusion ofnegative stain. Both the negatively stained conventionaltransmission electron microscope images and the radialdensity profile derived from unstained STEM images concurthat fimbriae of this type have relatively high density at theircenters, i.e., they do not have hollow axial channels. More-over, the radial density profile (Fig. 4c) indicates that theoutermost density is, on average, diffuse, only -10% of thetotal mass lying external to a radius of 3.0 nm. We note thata similar intercalation phenomenon has previously beendocumented in the actin system, where the center-to-centerspacing in actin paracrystals is -6.8 nm (15), whereas theouter diameter of F-actin is considerably higher, possibly asmuch as 10.0 nm (4).A three-dimensional model for the packing of subunits in

the fimbria, based on the deduced surface lattice and on theradial density map, is shown in Fig. 3c.

Layered aggregates of fimbriae. In an earlier study, Blomet al. (2) reported another type of paracrystalline aggregateconsisting of layers of parallel filaments spaced -9 nm apart,with successive layers oriented at a crossing angle of -40°.On the basis of optical filtering of electron micrographs, theyproposed a structural model in which these filaments wereenvisaged to consist of trinodular subunits, 13 nm in length,arranged end-to-end, which they interpreted in terms of thefilamentous hemagglutinin of B. pertussis. It has since beenestablished on both antigenic and biochemical grounds thatthe filamentous hemagglutinin is an entirely different mole-cule from fimbrial protein (1, 26). For instance, the filamen-tous hemagglutinin comprises several proteins in the 100- to160-kDa range, whereas serotype 2 fimbrial protein is ahomogeneous species of 22 kDa. Nevertheless, we considerit likely that the layered aggregates described by Blom et al.(2) were indeed packings of fimbriae, because their geomet-

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STRUCTURE OF BORDETELLA PERTUSSIS FIMBRIAE 973

rical parameters (interfilament spacing and crossing angle)are explained in a straightforward way in terms of the helicalstructure that we determined for the fimbria. Its axial repeatdistance is 13 nm, and the predominant feature of its surfacerelief is a set of alternating ridges and grooves with a pitch of6.5 nm, i.e., the features that, when stained, give rise to thestrong 1 = 2 layer line in the diffraction patterns of ourparacrystals (Fig. 2). This family of helices has a pitch angle(a) of 210 at a radius of 2.7 nm (i.e., half the interaxis spacingbetween two close-packed fimbriae). Thus, if (as illustratedin Fig. 5) one fimbria (A) is rotated relative to a second (B)so that its ridges are parallel to, and fit into, the correspond-ing grooves of fimbria B (and vice versa), their crossing-angle is 2ac = 42°. Now, if a third fimbria (A') is to cross Bparallel to A, and interact with B via precisely the same setof contacts as A, its crossing point should be 13 nm furtheralong B (or some integral number of axial repeats). Thus, theinterfilament spacing (d) is related to the axial repeat (D) andthe crossing-angle (2a) by the relationship d = D cos(2a),and the value predicted for d, 9.6 nm, comes out very closeto the observed value (17). According to this construction,the layered aggregates are held together only by directinteractions between successive layers and not betweenadjacent fimbriae within a layer, because their spacing (9.0 to9.6 nm) substantially exceeds the outer diameter of thefimbria (-7.5 nm). Moreover, these aggregates may beconsidered as true crystals, based on a single specific type ofinterfimbrial contact, although crystals generated by thiscontact alone are restricted to two layers in thickness. Forfimbriae in either layer of a two-layer aggregate, the helicalsymmetry of the individual fimbria precludes the presenceon the outer surface of sites that could engage a third layerby the same interaction.To summarize, the axial repeat (13.0 nm) determined here

for purified fimbriae is in essential aggreement with the12.6-nm repeat reported by Blom et al. (2) for their "layeredaggregate" filaments. However, we propose that the latterare composed of the fimbrial protein, not filamentous hemag-glutinin, and that their axial repeat contains five equivalentsubunits, not three "nodules." Their publisheddiffractogram (Fig. 14 of reference 2) indexes on a two-dimensional reciprocal lattice with a rhombic unit cell, suchas would be generated by the crossing of two parallel arraysof uniform cylinders, appropriately spaced. We detect noother reflections in the pattern that could correspond tosampling of helical layer lines and which would conveysubstructure of a single filament. Thus, we consider that theinferred trinodular repeating element (2) most likely repre-sents an artifact of the optical filtering.Comparison with proposed helical structures of other

fimbriae. Specific surface lattices have been attributed tothree other kinds of fimbriae. (i) E. coli type I fimbriae havebeen described in terms of a single-start helix of 2.3-nm pitchwith 3.125 subunits per turn (3); (ii) a single-start helix of4.1-nm pitch with about four (6) or five (24) subunits per turnhas been assigned to the polar fimbriae of P. aeruginosa; (iii)a four-start helix of 12.8-nm pitch with -8.7 subunits perturn in each strand has been deduced for the F-pili of E. coli(5). There are no obvious interrelationships among thesesurface lattices, nor do any of them resemble that estab-lished here for B. pertussis fimbriae. Moreover, unlike B.pertussis fimbriae, all three are thought to be hollow, and allare substantially denser, corresponding to linear densitiesthat are two- to threefold higher than the 8.9-kDa/nm valueof type 2 fimbriae.

Alteration without change (11). In contrast to the diversity

of fimbrial structures discussed above, it is noteworthy thatthe helical structures of type 2 and type 6 fimbriae areidentical, notwithstanding the antigenic and molecularweight differences between their respective subunits. Thisuniformity of supramolecular structure would be readilyexplained if the differences between the respective polypep-tide chains were confined to residues that occupy peripherallocations in the fimbria and if its backbone consisted ofconserved portions of the fimbrial subunits which, in partic-ular, should mediate the contacts between the polymerizedmolecules. Such a partitioning of conserved and variableregions would admit alterations in the antigenic character ofthe fimbria while preserving its assembly properties, asreflected in the helical structure.A similar mechanism has recently been proposed to rec-

oncile the range of different protein subunits found in variousintermediate filaments of eucaryotic cells with their seem-ingly common (or at least closely related) supramolecularstructures (16, 18). A precedent of conservation ofsupramolecular structure between related fimbriae is indi-cated by the close resemblance between the respectiveX-ray fiber diffraction patterns of PAO and PAK fimbriae ofP. aeruginosa (6), which are antigenically distinct and whoseamino-terminal sequences are closely homologous whiletheir carboxy-terminal sequences are considerably less so(12). Furthermore, it has been discovered that the molecularbasis of phase changes in gonococcal fimbriae resides in thevariable C-terminal domains of this otherwise conservativefamily of polypeptide chains (10). Although the higher-orderstructures of these fimbriae have not yet been determined,extension to them of the present hypothesis would allow thatthe supramolecular organization of gonococcal fimbriae maywell be invariant upon phase change, as the packing ofsubunits would be expected to be determined by the con-served domain(s) of their fimbrial protein molecules. How-ever, further experimental evidence is certainly required toclarify the status of this hypothesis, as applied both to B.pertussis fimbriae and to those of other bacterial systems.

ACKNOWLEDGMENTS

We gratefully acknowledge J. Wall and J. Hainfeld of the Depart-ment of Biology, Brookhaven National Laboratory, for provision ofSTEM data and thank Kristin Elmore for specimen preparation andFrank Kito for operation of the microscope. We also thank R. D. B.Fraser for a helpful discussion concerning the crystalline packing ofhelices and Todd Simpson for contributing to the computer analysis.

LITERATURE CITED1. Ashworth, L. A. E., L. I. Irons, and A. B. Dowsett. 1982.

Antigenic relationship between serotype-specific agglutinogenand fimbriae of Bordetella pertussis. Infect. Immun.37:1278-1281.

2. Blom, J., G. A. Hansen, and F. M. Poulsen. 1983. Morphology ofcells and hemagglutinogens of Bordetella species: resolution ofsubstructural units in fimbriae of Bordetella pertussis. Infect.Immun. 42:308-317.

3. Brinton, C. C. 1965. The structure, function, synthesis andgenetic control of bacterial pili and a molecular model for DNAand RNA transport in Gram-negative bacteria. Trans. N.Y.Acad. Sci. 27:1003-1054.

4. Egelman E. H., and R. Padron. 1984. X-ray evidence that actinis a 10OX filament. Nature (London) 307:56-58.

5. Folkhard, W., K. R. Leonard, S. Malsey, D. A. Marvin, J.Dubochet, A. Engel, M. Achtman, and R. Helmuth. 1979. X-raydiffraction and electron microscopic studies on the structure ofbacterial F-pili. J. Mol. Biol. 130:145-160.

6. Folkhard, W., D. A. Martin, T. H. Watts, and W. Paranchych.1981. Structure of polar pili from Pseudomonas aeruginosa

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974 STEVEN ET AL.

strains K and 0. J. Mol. Biol. 149:79-93.7. Hewlett, E. L., and J. Wolff. 1976. Soluble adenylate cyclase

from the culture medium of Bordetella pertussis. J. Bacteriol.127:890-898.

8. Labaw, L. W., E. A. Padlan, D. M. Sehgal, and D. R. Davies.1975. An EM study of phosphorylcholine-binding Fab' immu-noglobulin fragment crystals. J. Ultrastruct. Res. 51:326-339.

9. Mosesson, M. W., J. Hainfeld, R. H. Haschemeyer, and J. Wall.1981. Identification and mass analysis of human fibrinogenmolecules and their domains by scanning transmission electronmicroscopy. J. Mol. Biol. 153:695-718.

10. Schoolnik, G. K., R. Fernandez, J. Y. Tai, J. Rothbard, andE. C. Gotschlich. 1984. Gonococcal pili: primary structure andreceptor binding domain. J. Exp. Med. 159:1351-1370.

11. Sharpe, T. 1974. Porterhouse blue, p. 25. Martin, Secker andWarburg, London.

12. Shastry, P. A., B. B. Finlay, B. L. Paslosky, W. Paranchych,J. R. Pearlstone, and L. B. Smillie. 1985. Comparative se-quences of the amino acid and nucleotide sequences of pilinderived from Pseudomonas aeruginosa PAK and PAO. J.Bacteriol. 164:571-577.

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