Physiology andEvolution ofSpirochetes · spirochetes indigenous to healthy humans or animals...

BACTzjIOwGICAL Ruviuws, Mar. 1977, p. 181-204Copyright C 1977 American Society for Microbiology

Vol. 41, No. 1Printed in U.S.A.

Physiology and Evolution of SpirochetesE. CANALE-PAROLA

Department of Microbiology, University of Massachusetts, Amherst, Massachusetts 01002

INTRODUCTION ........................................ 181METABOLISM OF SPIROCHETES ................................... 182

Cultivation of Spirochetes ................................... 182Anaerobic Energy-Yielding Metabolism ................................... 183Aerobic Dissimilatory Metabolism ................................... 187Physiology of T. pallidum ................................... 189Lipids and Fatty Acids ................................... 189Carotenoid Pigments ................................... 191Motility and Chemotaxis ................................... 191

EVOLUTION OF SPIROCHETES ................................... 194The Protospirochete Hypothesis ................................... 194Convergent Evolution Hypothesis ................................... 195Energy-Yielding Pathways of Spirochetes .................................... 195Development of Associations with Hosts ................................... 197Possible Relationship with Gliding Bacteria ................................... 197Spirochetes, Eucaryotic Flagella, and Cilia ................................... 197Concluding Remarks ................................... 198Summary ................................... 198

LITERATURE CITED................................... 199

INTRODUCTION

Since the discovery of the microbial world,morphological characteristics frequently havebeen assigned primary significance in the inter-pretation of "natural" or phylogenetic relation-ships among microorganisms. Thus, attemptsto construct classifications based on phylogeny,or to define microbial groups or taxa, have re-lied heavily upon morphological criteria.The group of bacteria called spirochetes (or-

der Spirochaetales) may be considered an ex-ample of a taxon that was proposed as a resultof the tendency to attribute phylogenetic ortaxonomic value to morphological features. Ac-cording to modern concepts, spirochetes are hel-ically shaped, heterotrophic bacteria (33). Theoutermost structure of the spirochetal cell is athin, three-layered membrane called "outersheath" or "outer cell envelope" (Fig. 1), possi-bly corresponding to the "outer membrane" ofgram-negative bacteria. This outer sheath com-pletely surrounds the body of the cell or proto-plasmic cylinder, which consists ofthe cytoplas-mic and nuclear regions enveloped by the cellmembrane and the cell wall. The helical proto-plasmic cylinder is wound together with a num-ber of filamentous structures, called "axial fi-brils" or "axial filaments," and both the proto-plasmic cylinder and the axial fibrils are en-closed by the outer sheath (Fig. 1). The numberof axial fibrils present per cell ranges from 2 tomore than 100, depending on the kind of spiro-chete. One end of each axial fibril is inserted

near one pole of the protoplasmic cylinder,whereas the other end is not inserted. Sinceindividual axial fibrils extend for most of thelength of the cell, axial fibrils inserted at oppo-site poles of the protoplasmic cylinder overlapin the central region of the cell.

In accordance with present taxonomic crite-ria, all bacteria that possess the morphologicalfeatures described above are spirochetes, ormembers of the order Spirochaetales (33). Prob-ably as a result of their unique cellular archi-tecture, spirochetes possess a type ofmovementnot observed in other bacteria. Thus, the spiro-chetal cell, which has no exoflagella, can loco-mote or swim in liquid environments withoutbeing in contact with solid surfaces. Further-more, spirochetes locomote by "creeping" or"crawling" on solid surfaces (23, 45). Because oftheir ultrastructural and chemical resem-blances to bacterial flagella, it has been sug-gested that axial fibrils play a role in the motil-ity of spirochetes (26).Although spirochetes have certain basic mor-

phological characteristics in common, they ex-hibit extreme phenotypic diversity. For exam-ple, they vary greatly in size, ranging fromvery small cells (e.g., 0.1 by 6 gum) to largeforms 0.75 to 3 /Am thick and 100 ,um in lengthor longer (33). Moreover, pronounced physiologi-cal differences exist among spirochetes. Thus,aerobic, facultatively anaerobic, as well as obli-gately anaerobic, species have been described.Almost total specialization in energy-yieldingmechanisms has been achieved by some spiro-

181

on January 21, 2021 by guesthttp://m

mbr.asm

.org/D

ownloaded from

http://mmbr.asm.org/

FIG. 1. Schematic representation of a spirochete. The broken line indicates the outer sheath (outer cellenvelope). The area delimited by the thick solid line, adjacent to the broken line, represents the protoplasmiccylinder. The circles near the ends ofthe protoplasmic cylinder indicate the insertion points ofthe axial fibrils.The solid thin lines, wound around the protoplasmic cylinder, are the axial fibrils.

chetes, which are limited to dissimilating onlycertain long-chain fatty acids and a few long-chain alcohols (98). Other spirochetes deriveenergy exclusively by fermenting sugars (37,38, 77, 78, 80), whereas others possess remarka-ble metabolic versatility, being able to catabol-ize a variety of amino acids and carbohydrates(79; E. Canale-Parola and R. P. Blakemore,Abstr. Annu. Meet. Am. Soc. Microbiol., 1975,K52, p. 155). Furthermore, spirochetes are pres-ent in a wide spectrum of natural habitats.Many occur, grow, and persist as free-livingforms in bodies of marine and fresh water andin mud (33, 36). Others are part of the normalmicroflora present on or in eucaryotic hosts.Many of these host-associated spirochetes havebecome adapted to life within specialized habi-tats, such as the crystalline style of molluscs(26), the gingival crevice of humans (26), thecolon of mammals, where they are attached toepithelial cells (127, 164), or the body surface ofprotozoa (24, 26, 42, 153). A relatively smallnumber of host-associated spirochetes have theproperty of pathogenicity and are causativeagents of diseases, such as relapsing fever, lep-tospirosis, syphilis, and other treponematoses(47).The multifarious phenotypic manifestations

of spirochetes reflect marked differences in thegenotypes of these bacteria. In fact, it has beenfound that spirochetes differ greatly in deoxyri-bonucleic acid base composition, with guanineplus cytosine contents ranging from 36 to 66mol% (33). Physiological differences, as well asecological, morphological, and other considera-tions, have been used in classifying the spiro-chetes into five genera: Spirochaeta, Cristis-pira, Treponema, Borrelia, and Leptospira(33). Table 1, included for the purpose of clarify-ing the nomenclature used in this article, sum-marizes some of the characteristics of these fivegenera.My main objective in writing this review ar-

ticle is to examine and discuss recent literatureon the physiology of spirochetes, although lessrecent publications pertinent to the presenta-tion are also considered. Also, I shall reviewand analyze published data that seem to havesignificance with regard to the evolutionary

history of spirochetes. My ultimate intention isto stimulate the interest of microbiologists inthis diverse and intriguing group ofbacteria.

METABOLISM OF SPIROCHETESCultivation of Spirochetes

There have been relatively few extensivestudies on the metabolism of spirochetes. Thislack of effort may be ascribed to the fact that formany years after their discovery (50), spiro-chetes have been an ill-defined group of micro-organisms, attracting the attention of a limitednumber of microbial physiologists. Further-more, many spirochetes, especially many of theanaerobic host-associated forms, are not readilymass cultured inasmuch as they are nutrition-ally fastidious. Generally, the cultivable anaer-obic host-associated spirochetes are grown incomplex media supplemented with blood se-rum, serum components, ascitic fluid, or rumenfluid. Chemically defined growth media, someof which contain over 50 medium components,have been described for a few host-associatedanaerobic spirochetes (151, 152).Growth yields of spirochetes vary consider-

ably. The oral spirochete Treponema denticolagrows in complex media to a density of 5 x 108cells per ml and has doubling times of 12 to 14 h(79). T. vincentii, another oral spirochete, hasbeen grown to a density of 5 x 107 cells per ml inmedia containing ascitic fluid (129). Among thepathogens, the leptospires reach densities of 108to 4 x 108 cells per ml in media to which serumor an albumin-fatty acid supplement has beenadded (6). Cultivation techniques by whichhigh yields of virulent treponemes or borreliaemay be obtained have not been developed. Aculture medium suitable for the growth of sev-eral species of relapsing fever borreliae hasbeen described (106, 137), but this medium sup-ports maximum cell yields of only 5 x 107 cellsper ml after 7 days of incubation (106). Similaror somewhat higher yields are obtained whenT. hyodysenteriae, the primary etiologicalagent of swine dysentery (85), is grown in brothcultures (110). Various other virulent trepo-nemes and borreliae have not been grown inpure culture in vitro. Among these is T. palli-

182 CANALE-PAROLA BACTRIMOL. Rzv.


mbr.asm

.org/D

ownloaded from


PHYSIOLOGY AND EVOLUTION OF SPIROCHETES 183

TABLE 1. Summary of spirochete classification and nomenclaturea

CharacteristicsFree-living in aquatic environments. The genus includes three obligately anaerobicspecies (S. stenostrepta, S. zuelzerae, S. litoralis), one facultatively anaerobic species (S.aurantia), and a species (S. plicatilis) that has not been isolated. A red-pigmentedhalophilic, facultative anaerobe, referred to as spirochete RS1 (67), is a member of thisgenus and the name S. halophila RS1 has been proposed for it (Greenberg and Canale-Parola, in press). Spirochete RS1 was isolated from a solar lake and requires 0.75 MNaCl, 0.2 M MgSO., and 0.01 M CaCl2 for optimum growth. Species ofSpirochaeta havea G + C content of DNA ranging from 50 to 66 mol% (buoyant density) (33).

Large spirochetes usually found in the digestive tract of many marine and freshwatermolluscs (33). Not grown in pure culture. See review by Breznak (26).

Present in the mouth, intestinal tract, and genital areas ofhumans and animals. Manyare members of the normal microflora of the healthy human and animal body. Somespecies (e.g., T. pallidum, T. pertenue, T. carateum) are pathogenic. Species that havebeen cultivated in pure culture in vitro are obligate anaerobes. Certain nonvirulenttreponemes reportedly isolated from syphilitic lesions or other pathological processes areknown by the trivial names of Reiter treponeme, Kazan treponeme, Nichols nonpatho-genic treponeme, and Noguchi treponeme (174). In Bergey's Manual (33) the Reiter andKazan treponemes are designated T. phagedenis, whereas the Nichols nonpathogenicand the Noguchi treponemes are designated T. refringens. Some workers (71) do notbelieve that the available evidence is sufficient to justify the use of the name T.phagedenis for the Reiter treponeme. In this review I refer to the four above-namedtreponemes by their trivial names. Species of Treponema have a G + C content of theDNA ranging from 37 to 46 mol% (33, 38).

Cause relapsing fever in humans and similar diseases in animals (107, 151). Transmittedby lice or ticks. Anaerobic (33) or microaerophilic (107).

Small, obligately aerobic spirochetes, generally with one or both ends of the cells bent orhooked. Found free-living in surface waters or soil and in association with animals andhumans (33, 76, 166). Host-associated leptospires can cause disease (leptospirosis) inhumans and other mammals. The G + C content of the DNA is 36 to 39 mol% (33).

a See Bergey's Manual (33) for detailed classification and for micrographs illustrating the overall mor-

phology and size ofrepresentative spirochetes. G + C, guanine plus cytosine; DNA, deoxyribonucleic acid.

dum, the causative agent of syphilis, which isgenerally maintained in a reproducing state byinoculation in living animals, i.e., rabbits. Re-cently, a report has been published describingthe cultivation of virulent T. pallidum in cul-tures containing baby hamster kidney tissuecells (99). In vitro cultivation has not beenachieved with certain relapsing fever borreliae(70), such as Borrelia duttonii, which has beengrown in experimental animals by investiga-tors who studied its metabolism (60). Manyspirochetes indigenous to healthy humans oranimals have never been cultivated. These in-clude various spirochetes that are inhabitantsof the gut of termites, the crystalline style ofmolluscs, and the intestine of humans.

In contrast to the host-associated forms, free-living spirochetes grow abundantly in serum-free, readily prepared media, reaching high celldensities which, in some cases, approach 1010cells per ml (36, 77). Thus, these organismsserve as useful tools in investigations for which

large amounts of cell material are desirable. Asa consequence, much of our present knowledgeon the metabolism of spirochetes has been ac-quired through the study of free-living forms.Various spirochetes have been isolated from

natural habitats using selective enrichmentmethods. Such methods, which have been de-scribed in detail elsewhere (36), are based onthe ability of spirochetes to pass through filtersof small pore diameter (0.2 to 0.45 ,um, averagediameter) and on the property of spirochetes tomigrate more readily than many other bacteriathrough agar gels. These enrichment tech-niques have been used successfully for the iso-lation of thin spirochetes (0.5 gm or less indiameter). Selective isolation methods for spi-rochetes of larger cell diameter have not beendeveloped.

Anaerobic Energy-Yielding MetabolismAll strains of anaerobic and facultatively an-

aerobic free-living spirochetes that have been

ClawsificationSpirochaeta

Cristispira

Treponema

Borrelia

Leptospira

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


184 CANALE-PAROLA

isolated are carbohydrate fermenters. None hasbeen shown to ferment amino acids, but it ispossible that such strains will be obtained byusing appropriate selective enrichment tech-niques.Under anaerobic conditions, all fermentative

free-living spirochetes tested form acetate,CO2, and H2 as major products of carbohydratedissimilation (Table 2). Ethyl alcohol is anothermajor product, except that one species (Spi-rochaeta zuelzerae) does not produce it and spi-rochete Z4 only forms small amounts of it (Ta-ble 2). The latter two spirochetes maintaincharge balance not only by producing H2, butalso by reducing greater amounts of pyruvateto lactate than those spirochetes that formethyl alcohol as a major product. Furthermore,both S. zuelzerae and spirochete Z4 producesuccinate (Table 2).

The pathways of carbohydrate fermentationof S. aurantia, S. litoralis, and S. stenostreptahave been investigated by me and my co-workers(27, 28, 37, 38, 77, 78, 80, 91). Assays of enzy-

matic activities in cell extracts and determina-tions of radioactivity distribution in productsformed from 14C-labeled substrates showed thatthese spirochetes ferment glucose via the Emb-den-Meyerhof (EM) pathway. Cells or extractsof S. aurantia and S. stenostrepta ferment py-ruvate to the same major products formed fromglucose, except that acetoin is produced by cellsuspensions or cell extracts of both spirochetesand diacetyl is produced by S. aurantia ex-

tracts. Whole cells or cell extracts of S. auran-

tia, S. litoralis, and S. stenostrepta exhibit a

coenzyme A-dependent C02-pyruvate ex-

change. No formate-pyruvate exchange was de-tected in whole cells or extracts, and the datashowed that free formate is not involved in C02and H2 production. It was concluded that thethree organisms utilize a clostridial-type clasticsystem to metabolize pyruvate to acetyl-coen-zyme A, C02, and H2 (Fig. 2). Enzymatic assaysof cell extracts showed that acetyl-coenzyme Ais converted to ethyl alcohol by nicotinamideadenine dinucleotide-dependent acetaldehydeand alcohol* dehydrogenase activities (EC1.2.1.10 and EC 1.1.1.1, respectively). Further-more, phosphotransacetylase (EC 2.3.1.8), ace-

tate kinase (EC 2.7.2.1), lactate dehydrogenase(EC 1.1.1.27), and hydrogenase activities were

|CARBOHYDRATE S

ATP- |PATHWAY

PYRUVATE LACTATE

CLOSTRIDIAL- TYPECLEAVAGE

CO2 H2

ACETALDEHYDE=, ACETYL - COA .ACACETYL-PO4

g 3_~~~~~~~~~ATPETHANOL ACETATE

FIG. 2. Pathways for anaerobic dissimilation ofcarbohydrates by free-living spirochetes. The brokenarrow indicates a minor pathway.

TABLE 2. Fermentation products offree-living spirochetesAmt of products (gtmol/100 ,umol of glucose fermented)a

Productsb S. aurantia S. litoralis S. stenostrepta S. zuel- Spirochete Spirochete

1 2 1 2 1 2 zerae (1) Z4 (1) RS1(1)

Acetate 69.2 50.3 37.5 57.0 93 20.4 82 94.8 52.4Ethyl alcohol 151.0 78.4 109.5 140.5 84 146.2 ND 10.5 132.0CO2 165.3 128.2 127.5 201.8 140 187.5 68 72.7 176.1H2 107.2 79.5 74.0 74.4 180 27.2 164 186.9 130.3Lactate 1.0 17.2 6.5 Trace 10 8.2 87 56.8 1.8Formate 5.2 NR 2.8 Trace Trace NR ND 10.7 NDPyruvate NR 3.1 0.3 Trace NR NR NR NR NDSuccinate ND NR ND ND ND ND 13 26.3 NDGlycerol NR 4.4 NR NR NR NR ND NR NRAcetoin, di- Trace NR ND ND ND NR ND ND ND

acetyla 1, Products of growing cells; 2, products of cell suspensions. ND, Not detected; NR, not reported.b Data for S. aurantia are from Breznak and Canale-Parola (27, 28), for S. litoralis from Hespell and

Canale-Parola (78), for S. stenostrepta from Canale-Parola et al. (37) and Hespell and Canale-Parola (77), forS. zuelzerae from Veldkamp (171), for spirochete Z4 from Canale-Parola et al. (38), and for spirochete RS1from Greenberg and Canale-Parola (in press).

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



detected in cell extracts of all three species. Thedata indicated that in S. aurantia, S. litoralis,and S. stenostrepta the acetyl-coenzyme Aformed via the pyruvate clastic system is con-verted to acetate in reactions catalyzed by phos-photransacetylase and acetate kihase and toethanol through a double reduction involvingaldehyde and alcohol dehydrogenase activities(Fig. 2). A small proportion of pyruvate is notcleaved but is reduced to lactate (Fig. 2). Theproposed pathways (Fig. 2) indicate that in ad-dition to the net yield of 2 mol of adenosinetriphosphate (ATP) per mol of glucose metabo-lized via the EM pathway, the spirochetes de-rive additional ATP through reactions leadingto acetate formation from pyruvate.

Anaerobic molar growth yield determina-tions were conducted with spirochete RS1,which has fermentation pathways indistin-guishable from those of S. aurantia, S. litor-alis, and S. stenostrepta (E. P. Greenberg andE. Canale-Parola, Arch. Microbiol., in press).Results of these studies indicated that the ATPyield predicted by the proposed pathways (Fig.2) would support anaerobic molar growth yieldsof the magnitude observed. This finding, aswell as studies with the three species of Spiro-chaeta previously discussed (28, 77, 80), indi-cate that the pathways outlined in Fig. 2 consti-tute the major anaerobic energy-yielding mech-anisms utilized by these free-living spirochetes.

In nature, obligately anaerobic species ofSpirochaeta commonly occur in sulfide-contain-ing mud or water. Thus, it was thought possiblethat these spirochetes carried out anaerobicrespiration, using sulfate as the electron accep-tor. However, experiments conducted with S.stenostrepta (77) showed that H2S was notevolved by this spirochete growing in variousmedia to which were added ammonium sulfateand possible electron donors such as pyruvate,lactate, malate, or glucose. A small amount ofH2S was evolved when the organism was cul-tured in media including sodium thioglycolate,but the gas probably originated from thesulfhydryl group of this compound.

Iron-sulfur proteins, such as rubredoxins andferredoxins, participate in the metabolic activi-ties of many anaerobic bacteria. Ferredoxinsfimction as electron carriers in various oxida-tion-reduction reactions (32). Rubredoxins ofanaerobic bacteria are known to participate inelectron transfer (119), but the specific physio-logical role(s) of these proteins has not beenfound. A rubredoxin isolated from the aerobicPseudomonas oleovorans is required for the hy-droxylation of fatty acids and hydrocarbons(135, 136). Since iron-sulfur proteins are com-

monly present in H2-producing organisms pos-sessing a clostridial-type pyruvate clastic sys-tem, investigators have searched for these pro-teins in S. aurantia, S. stenostrepta, and S.litoralis (28, 77, 80, 91). Rubredoxins were iso-lated from cell extracts of these three species.Johnson and Canale-Parola (91) reported thatrubredoxins from S. aurantia and S. stenos-trepta contained one atom of iron per moleculeand no inorganic sulfide. The proteins' molecu-lar weight (ca. 6,000), spectral properties, andamino acid composition were similar to those ofrubredoxins obtained from other anaerobic bac-teria. The same authors (91) isolated an unsta-ble ferredoxin from cell extracts of anaerobi-cally grown S. aurantia. This ferredoxin hadspectral characteristics and amino acid compo-sition typical of other bacterial ferredoxina. Itcontained four atoms of iron and four acid-la-bile sulfide residues per molecule, and its mo-lecular weight was near 6,000. Purified spiro-chetal ferredoxin stimulated acetyl phosphateformation from pyruvate by clostridial extractsdepleted of iron-sulfur proteins by passagethrough diethylaminoethyl-cellulose columns(91). Stimulation of acetyl phosphate formationfrom pyruvate was also observed when Clos-tridium butyricum ferredoxin was added to di-ethylaminoethyl-cellulose-treated S. stenos-trepta extracts (77). These results indicate thatferredoxin participates in the anaerobic pyru-vate metabolism of spirochetes. Apparently,spirochetal rubredoxin has a different role,since it did not stimulate acetyl phosphate for-mation from pyruvate by diethylaminoethyl-cellulose-treated C. butyricum extracts (91).Johnson and Canale-Parola (91) reported

that rubredoxin was present in both aerobicallyand anaerobically grown cells of S. aurantia,whereas ferredoxin was not detected in aerobi-cally grown cells. Thus, ferredoxin does nottake part in the metabolism ofS. aurantia cellsgrowing in air. Rubredoxin preparations fromaerobically or anaerobically grown S. aurantiahad identical spectral characteristics, molecu-lar weight, and iron content. These findingsshow that rubredoxin from aerobically grownS. aurantia differs from P. oleovorans rubre-doxin, which has a molecular weight of 19,000and binds either one or two atoms of iron (168).It would be interesting to determine whetherthe rubredoxin occurring in aerobically grownS. aurantia functions in hydroxylation mecha-nisms similar to those present in P. oleovorans.

Quantitative information has been reportedon the anaerobic dissimilatory pathways of afew host-associated spirochetes. In one of thesestudies, B. duttonii cells were separated from

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


186 CANALE-PAROLA

the blood of heavily infected rats by a methodinvolving differential centrifugation (60). Sus-pensions of B. duttonii cells incubated in thepresence of glucose converted this sugar mainlyto lactate, without C02 evolution or 02 uptake.The molar ratio of glucose utilized to lactateformed was close to 1:2. Fluoride and iodoace-tate inhibited acid production from glucose,whereas azide had little or no effect. Finally,enzymes of the EM pathway were detected incell extracts (154-156). The results indicate thatB. duttonii obtains energy by a homolactic fer-mentation in which carbohydrate is metabo-lized to pyruvate via the EM pathway; pyruvateacts as the terminal electron acceptor and isreduced to lactate. Aerobic respiration was notdetected in the organism.Another series of studies dealt with T. denti-

cola, an anaerobic spirochete that is a commoninhabitant of the gingival sulcus and gingivalcrevice regions of the human mouth. The sul-cus-crevice regions, which include the furrowand fissure present between the gingiva andthe tooth enamel (34), are inhabited by an enor-mous variety and number of microorganisms(157), many of which have not been cultivated.Anaerobes and facultative anaerobes are preva-

lent in these regions (54, 157), with the latterorganisms probably serving as scavengers ofmolecular oxygen to generate anaerobiosis. Thephysiological interactions between the sulcus-crevice microorganisms and the host, andamong members of the microflora residing inthis habitat, constitute complex but fascinatingproblems of ecology. However, understandingof such interactions cannot be achieved untilextensive information becomes available on thephysiology of microorganisms residing in thesulcus-crevice regions. Such information was

sought by investigators who studied the metab-olism of T. denticola (79; Canale-Parola andBlakemore, Abstr. Annu. Meet. Am. Soc. Mi-crobiol., 1975, K52, p. 155; unpublished data).This organism was grown in complex mediacontaining serum, yeast extract, Trypticase,glucose, thiamine pyrophosphate, and othercomponents. To identify the substrates dissimi-lated by T. denticola, 14C-labeled compoundswere added to these media, and the amount ofradioactivity in the fermentation products wasmeasured. Furthermore, amino acid analysesof growth medium supernatant liquid were car-ried out before, .during, and at the end ofgrowth of the treponemes. These experimentswere complemented by amino acid analyses ofT. denticola cells, by determination of productsformed from various substrates by cell suspen-sions, and by measurements of growth stimula-

tion by different compounds added to media.The results of the experiments indicated thatT. denticola catabolized L-cysteine, glycine, L-serine, -alanine, L-arginine, L-citrulline, andL-histidine, as well as carbohydrates. The fer-mentation products included acetate, lactate,succinate, formate, pyruvate, ethanol, C02,H2S, and NH3. Acetate was the major non-gaseous fermentation product recovered. Underthe growth conditions used, the productsformed from glucose constituted a small portionof the total products that accumulated in cul-tures. Amino acid analyses of culture superna-tant fluid showed that growing cells of T. denti-cola dissimilated significant amounts of gluta-mate and aspartate as well. This observationwas in apparent contradiction with results ofprevious experiments, which indicated that cellsuspensions did not catabolize either of thesetwo amino acids and that only very low levels ofradioactivity were recovered in fermentationproducts of growing cells when L-[14C]glutamate was added to complex culturemedia. However, further experimentation in-volving amino acid analyses indicated that glu-tamate was dissimilated when it was added tocell suspensions in the bound form, that is, as acomponent of peptides or proteins. A possibleexplanation for these results is that T. denti-cola cells do not have a transport system forfree glutamate but are able to transfer gluta-mate-containing peptides across the cell mem-brane. Aspartate may be transported in a simi-lar way. This behavior would be analogous tothat of other bacteria, such as Bacteroides rum-inicola (139) and Fusiformis necrophorus (173),which do not have the ability to transport intothe cell certain free amino acids but apparentlytransport peptides, which function as carriersof amino acids.Assays of enzymatic activities in T. denticola

cell extracts indicated that this spirochete de-grades glucose via the EM pathway (79). Likethe anaerobic free-living spirochetes previouslymentioned, T. denticola possesses a coenzymeA-dependent C02-pyruvate exchange activityassociated with a clostridial-type clastic systemfor pyruvate metabolism (Fig. 3). Pyruvate isdecarboxylated via this system to acetyl-coen-zyme A, which is converted to acetate throughthe action ofphosphotransacetylase and acetatekinase. Hydrogen gas is not formed by growingcells, and hydrogenase activity has not beendetected in cell extracts. Mechanisms to main-tain charge balance include the formation ofethanol, succinate, and lactate and the conver-sion of glycine to acetate. A rubredoxin or rub-redoxin-like protein was detected in cell ex-

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from


PHYSIOLOGY AND EVOLUTION OF SPIROCHETES 187EM CLOSTRIDIAL-TYPE

PATHWAY CLEAVAGE CO2[CARBOHYDRATES c =R PYRUVATEA

24 S ACETATE + ATP

IIPKH2S X ETHANOL

02 ~~~LACTATE>NH3L- CYSTEINEL- SERINEL-ALANINE

|L-ARGININE | L-CITRULLINEi ORNITHINE + PHOSPHATE

NH3 J ATP

CO2 NH3

FIG. 3. Energy-yielding pathways ofT. denticola. In addition to those indicated in this figure, other aminoacids are catabolized (see text).

tracts of T. denticola (91).T. denticola ferments some amino acids, such

as L-cysteine, i-serine, and L-alanine, by path-ways that involve pyruvate as an intermediate(Fig. 3). The pyruvate formed is decarboxyl-ated, with formation mainly of acetate. Elec-trons generated during pyruvate cleavage areused to form lactate and ethanol. Other aminoacids are dissimilated by pathways that do notinvolve pyruvate as an intermediate. Thus, T.denticola catabolizes i-arginine by a pathwaysimilar to that present in other bacteria (48;Fig. 3). This pathway involves conversion of L-arginine to citrulline, which is then cleaved toornithine and carbamoyl phosphate. The latteris then degraded to NH3 and C02, with forma-tion of ATP. Exogenously supplied L-citrullineis dissimilated through the same pathway(Canale-Parola and Blakemore, Abstr. Annu.Meet. Am. Soc. Microbiol., 1975, K52, p. 155).The picture that emerges from these meta-

bolic studies with T. denticola provides a bio-chemical interpretation for some of the ecologi-cal properties of a treponeme, which is a com-mon inhabitant of the human mouth. This an-aerobe possesses an abundance of diverse dis-similatory pathways that enable it to deriveenergy for growth from a relatively wide spec-trum of substrates. The metabolic flexibilitythat T. denticola possesses is probably one ofthe factors that enables it to overcome competi-tion by faster-growing microorganisms in itsnatural habitat and allows it to survive andthrive in an environment where differentgrowth substrates may become available or un-available in rapid succession to the indigenousmicroflora.The fermentation of glucose by an anaerobic

spirochete isolated from bovine rumen contentswas studied by Bryant (31). Products of this

fermentation were CO2, ethanol, succinic, lac-tic, acetic, and formic acids. The amount ofsuccinic acid formed accounted for 41% of thecarbon in the fermented glucose. H2 was notproduced. Another rumen spirochete producedformic, acetic, butyric, lactic, and succinic acidsfrom glucose (59). Ziolecki et al. (182) foundthat succinic and acetic acids were major prod-ucts of glucose fermentation by eight strains ofspirochetes isolated from the bovine rumen.Allen et al. (7) reported experiments indicat-

ing that glucose is a major carbon and energysource for the Reiter treponeme and suggestingthat arginine, histidine, serine, threonine, andglutamic acid may serve as energy sources forthis treponeme.

Other reports dealing with the dissimilatorymetabolism of anaerobic spirochetes describethe demonstration of various enzymes in trepo-nemes and S. zuelzerae (5, 8, 143, 165), thedeamination of amino acids and transamina-tion reactions in the Reiter treponeme (8, 9),gas evolution by treponemes and S. zuelzerae(114), cytochromes in the Reiter treponeme(105), and criteria for the biochemical differen-tiation of certain oral spirochetes (158).

Aerobic Dissimilatory MetabolismS. aurantia and the halophilic spirochete

RS1 (67), which are facultative anaerobes, andthe leptospires, which are obligate aerobes,couple oxidation of substrates with reduction ofmolecular oxygen. Breznak and Canale-Parola(27, 29) reported that cells of S. aurantia grow-ing in the presence of air performed an incom-plete oxidation of carbohydrates, producing pri-marily CO2, acetate, and pyruvate from glu-cose. From 20 to 30% of the total sugar carbonused was evolved as C02, and approximately50% was incorporated into cell material.

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


188 CANALE-PAROLA

Slightly more than one-third of the assimilatedglucose carbon was used for the synthesis of celllipids, which constituted 29 to 36% of the dryweight of S. aurantia. Apparently, a relativelylarge proportion of the acetyl-coenzyme A de-rived from carbohydrate oxidation was utilizedfor lipogenesis. A tricarboxylic acid cycle eitherwas not present or served in a minor cataboliccapacity.Aerobic and anaerobic molar growth yield

determinations showed that S. aurantia de-rived more energy from the aerobic oxidation ofsugars than from their fermentation andstrongly suggested the presence of oxidativephosphorylation mechanisms in this bacterium(29). A search for cytochromes revealed thepresence of cytochrome b558 and cytochrome o,which were associated primarily with a particu-late fraction of S. aurantia cell extracts. Thisfraction also contained 02-dependent reducednicotinamide adenine dinucleotide (NADH2)dehydrogenase activity. Cytochromes of the aor c type were not found in this spirochete.Protoheme, but not heme a or mesoheme, wasdetected. A scheme for terminal electron trans-port in S. aurantia was suggested (29):NADH2 -* flavoprotein -- [cytochrome b,, -

cytochrome o] -- 02

The brackets indicate that cytochrome b558 andcytochrome o may be one and the same hemo-protein, as has been observed in other bacteria(22). According to this scheme, the aerobic elec-tron transport in S. aurantia is a relativelyprimitive one and may constitute an early evo-lutionary attempt by spirochetes at synthesiz-ing functional hemoproteins (29). The aerobicenergy-yielding metabolism of halophilic spiro-chete RS1 is similar to that of S. aurantia(Greenberg and Canale-Parola, in press).The leptospires, which, as previously men-

tioned, are obligate aerobes, are quite distinctmetabolically from spirochetes capable of an-aerobic growth. Early attempts at studying themetabolism of leptospires established that res-piration of whole cells was stimulated by rabbitblood serum or crude phospholipid prepara-tions, but not by carbohydrates or amino acids(61, 123). It was suggested that the blood se-rum, which was usually included in growthmedia for leptospires, contained a constitu-ent(s) that served as respiratory substrate orotherwise stimulated respiration of leptospires(61, 123). It was later shown that long-chainfatty acids present in blood serum were respon-sible for the observed stimulation of respiratoryactivity (73). Furthermore, serum proteinserved as a "detoxifier" by removing inhibitory

effects caused by the fatty acids (73). As a resultof other studies (10, 51, 52, 74, 89, 93, 95, 98,148, 160, 161, 170, 172, 178), it was establishedthat fatty acids such as palmitate, stearate, oroleate serve as major carbon and energysources for leptospires. With the exception oflong-chain fatty alcohols (e.g., palmityl alco-hol), no other readily utilizable energy and car-bon sources are known for these spirochetes(98). Saturated or unsaturated fatty acids, fre-quently of chain lengths ranging from 15 to 18carbons, have been used to grow saprophytic orparasitic leptospires. Thus, these aerobic spiro-chetes utilize a rather limited range of oxidiza-ble substrates for growth. Paradoxically, freelong-chain fatty acids, even at very low concen-trations, usually inhibit the growth of lepto-spires, especially the pathogenic strains.Therefore, in culture media it is customary touse fatty acids in bound form, either as serumcomponents or as fatty acid esters, e.g., Tween80 (polyoxyethylene sorbitan monooleate), inorder to minimize the inhibitory or lethal ef-fects. It has been suggested that, in pathogenicleptospires, these effects may be caused byauto-oxidation products of fatty acids (161).Growth of a water leptospire was observed,after a long lag, in a medium containing ace-tate and L-histidine as the only possible sourcesof carbon and/or energy (74).The isolation of aquatic leptospires capable of

growing abundantly in serum-free media (12)has facilitated the study of leptospiral metabo-lism. It was found that growing cells of theaquatic leptospire B16 oxidized oleate to CO2and acetate. The amount of CO2 and acetateproduced accounted for 55.8 and 8.5%, respec-tively, of the oleate utilized. The cell materialformed represented 26.5% of the oleate con-sumed (74). Studies with an aquatic and a path-ogenic strain ofLeptospira indicated that theseorganisms use beta-oxidation for the degrada-tion of long-chain fatty acids (75). Furthermore,enzymatic assays of cell extracts of a waterleptospire and of two pathogenic leptospires in-dicated that enzyme activities of the tricarbox-ylic acid cycle and of the EM and pentose path-ways were present (10). It was concluded thatleptospires oxidize long-chain fatty acids bybeta-oxidation and utilize the resulting two-carbon fragments via the tricarboxylic acid cy-cle (10, 75). Label distribution patterns in cellu-lar amino acids of leptospires grown in the pres-ence of 14CO0 were consistent with the occur-rence of the tricarboxylic acid cycle (40). Enzy-matic activities of the EM and pentose path-ways may be utilized by leptospires mainly forbiosynthesis.

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



Cytochromes of the a, c, cl, and o types werefound in pathogenic and aquatic strains ofLep-tospira (11, 61). Carbon monoxide differencespectra suggested that a cytochrome a oxidase,)ossibly a, or a3, and a pigment with absorptioncharacteristics different from those of previ-ously described cytochromes were present intwo pathogenic strains, but not in an aquaticstrain of Leptospira (11). Another significantdifference betwen aquatic and pathogenicstrains of leptospires is that the former arecatalase negative, whereas the latter possesscatalase activity (11, 55, 141).

Representatives of different pathogenic lep-tospiral serotypes were found to utilize urea asa nitrogen source and to possess urease activity(102). These findings may be useful in inter-preting the localization of certain leptospires inthe kidney during leptospirosis.Other reports on the dissimilatory metabo-

lism of leptospires describe studies on lipaseactivity (16, 17, 41, 53, 94, 104, 108, 133, 134),transamination reactions (122), and various en-zymatic activities (35, 63, 65, 66).

Physiology of T. pallidumCox and Barber (44) reported that suspen-

sions of T. pallidum cells, extracted from in-fected rabbit testicles, consumed 02. Oxygenuptake was inhibited by 10 juM cyanide or 28mM azide. However, uptake of 02 by growingcells was not observed inasmuch as T. palli-dum has not been cultivated in pure culture invitro. It has been widely believed that T. palli-dum is an anaerobe, mainly because of reportsthat the cells retain their motility for longerperiods oftime under anaerobic conditions thanin the presence of air (33, 145). Retention ofmotility and virulence by T. pallidum cellsextracted from rabbit testicles is prolongedwhen the treponemes are suspended in mainte-nance media that have an electro-negative re-dox potential (64, 124).

Suspensions of T. pallidum cells extractedfrom rabbit testicles were reported to incorpo-rate amino acids into protein optimally in at-mospheres containing 10 to 20% 02 (13) and todegrade glucose to C02, acetate, pyruvate, andlactate (13, 131). Under anaerobic conditions,pyruvate and lactate were major products ofglucose degradation, whereas aerobically,greater amounts of CO2 and acetate wereformed (13).

Fitzgerald et al. (58) added T. pallidum tocultures of rabbit testicular tissue cells underaerobic conditions. They reported that some ofthe treponemes attached themselves to andpenetrated into the cultured animal cells. At-

tached and/or intracellular T. pallidum cellsretained virulence for longer periods of timethan cells incubated in the absence of culturedanimal cells. Furthermore, the presence of ani-mal cell monolayers extended the persistence oftreponemal motility. It was suggested that su-peroxide ions may be responsible for the toxiceffects of air on T. pallidum cells extractedfrom rabbit testicles and that within infectedtesticles superoxide dismutase present in themammalian tissue may prevent oxygen toxic-ity, thus allowing growth of the treponemes(44, 58).Interpretation of results obtained with T.

pallidum cells extracted from rabbit testicles iscomplicated by the possibility that such trepo-nemal populations may be heterogeneous. Thiswas suggested by a report of Baseman et al.(14), who found that T. pallidum populationsfrom infected rabbit testicles, when subjected tovelocity sedimentation in discontinuous gra-dients of Hypaque, separated into two distinctbands. The authors suggested that two inter-acting types of treponemes may exist in tissueduring the infectious process. Other authors(82) reported that electron microscopy revealedthe presence of rabbit cells and tissue debris inT. pallidum preparations. These preparationswere obtained from infected rabbit testicles byprocedures similar to those used by previousinvestigators to harvest T. pallidum cells formetabolic studies.A recent article by Jones et al. (99) describes

the cultivation of T. pallidum in a tissue cul-ture system. T. pallidum cells freshly har-vested from infected rabbit testicles, and sus-pended in culture medium, were added to mon-olayers of baby hamster kidney cells. Themixed cultures were incubated in an atmos-phere of 7% CO2 in air. Subculturing was car-ried out by transferring treponemes to freshhamster cell cultures. The authors reportedthat T. pallidum grew through nine subculti-vations (ca. three cell generations per subcul-ture) and that virulent cells were demonstratedin subcultures. The investigators emphasizedthat frequent subculturing (every 24 h) and theuse of a treponeme inoculum from a 43-day-oldtesticular infection were important factors inachieving continued cultivation of virulent T.pallidum cells in the system they used. Trepo-nemes from less prolonged rabbit testicular in-fections caused the cultured tissue cells to lyse,presumably because of the presence of a cyto-toxic factor in the treponemal preparations.

Lipids and Fatty AcidsCells of spirochetes contain a relatively large

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


190 CANALE-PAROLA

amount of lipids. The lipid content of Spiro-chaeta species ranges from 24 to 36% of the celldry weight (29, 100), the lipid content of trepo-nemes from 18 to 20% (96, 116), and that ofleptospires from 18 to 26% (97). The lipid compo-sition of a variety of spirochetes has been inves-tigated (43, 56, 96, 97, 100, 113, 116-118, 126,163, 169).Johnson et al. (97) and Livermore and John-

son (117) compared the lipid composition ofLep-tospira, Treponema, and Spirochaeta species.Phospholipid constituted 60 to 70% of the totallipid of leptospires, the remaining lipid beingfree fatty acids. The major phospholipid in lep-tospires was phosphatidyl ethanolamine. Phos-phatidyl choline, which was absent from lepto-spires, was a major phospholipid in various Tre-ponema species. Furthermore, the glycolipidmonogalactosyl diglyceride, not found in lepto-spires, was a major component of lipids in trepo-nemes. Among the Spirochaeta species, S.stenostrepta contained the monogalactosyl di-glyceride, and other species ofSpirochaeta con-tained the glucosyl or mannosyl analogue ofthis lipid. Phosphatidyl choline was not de-tected in species of Spirochaeta.The cellular fatty acid composition of spiro-

chetes reflects the fatty acid composition of theculture medium (96, 97, 163). For example, itwas reported that the Kazan 5 treponeme con-tained saturated and unsaturated fatty acidsranging from 14 to 18 carbon atoms, dependingupon the fatty acids added to the growth me-dium (96).With some exceptions, host-associated spiro-

chetes that have been cultivated require anexogenous supply of long-chain fatty acids forgrowth. Thus, culture media for host-associatedspirochetes usually are supplemented with se-rum or fatty acids complexed with albumin toprovide these required substances (118). Boththe Reiter and Kazan 5 treponemes require asaturated fatty acid with a chain length of atleast 14 carbon atoms and an unsaturated fattyacid with one, two, or three double bonds and achain length of 15 or more carbon atoms (92,132). The two fatty acids can be replaced by asingle 18-carbon monounsaturated fatty acidwith a trans configuration rather than the nat-urally occurring cis configuration (92). Require-ments for exogenous long-chain fatty acidshave been reported for oral treponemes, such asT. vincentii, T. denticola, and T. scoliodontum(117, 129). Leptospires utilize long-chain fattyacids as their major energy and carbon source,and strains that were studied usually requiredlong-chain fatty acids for growth. These strainscannot elongate the carbon chain of fatty acids

BACTERIOL. REV.

available to them (97) and thus require fattyacids with chain lengths of 15 carbons or more(98). B. hermsii has a requirement for exoge-nous long-chain fatty acids (137).

Short-chain fatty acids are known to be re-quired by a number of spirochetes. Some oraltreponemes require isobutyrate (72, 159), and arumen spirochete requires branched andstraight, short-chain fatty acids (176). The lat-ter organism apparently uses the short-chainfatty acids present in the growth medium forthe synthesis of long-chain fatty acids.Two spirochetes, isolated from the human

mouth and two others isolated from pig fecesare unusual because they grow in media lack-ing long-chain fatty acids but containing isobu-tyrate and valerate (118). Presumably, isobu-tyrate and valerate serve as precursors for thesynthesis of long-chain fatty acids.

Studies conducted with several host-associ-ated spirochetes have shown that the require-ment for long-chain fatty acids results from theorganisms' inability to manufacture these com-pounds, which are needed for cellular lipid bio-synthesis. For example, the Kazan 5 treponemecannot synthesize, modify the chain length of,reduce, or desaturate long-chain fatty acids(96). Similar findings have been reported forthe Reiter treponeme (126). Leptospires, grow-ing in media including long-chain fatty acidsand 14C-labeled acetate, incorporated only asmall amount of label into cellular fatty acids(163). Desaturation of fatty acids was accom-plished by leptospires in the presence of molec-ular oxygen (97, 163).Among the free-living spirochetes, the ability

to synthesize long-chain fatty acids is not un-common. S. zuelzerae synthesizes its cellularfatty acids via a pathway originally discoveredin C. butyricum (125, 146). Furthermore, sincewater leptospire B16 (74) and species of Spiro-chaeta (27, 78) grow in media devoid of long-chain fatty acids, it can be inferred that thesefree-living spirochetes are able to synthesize allfatty acids required for their cellular lipids. Theability to synthesize long-chain fatty acids isprobably widespread among free-living spiro-chetes other than leptospires.A rumen spirochete was shown to hydrogen-

ate linoleic acid to octadec-trans-11-enoate(144). The hydrogenation occurred in two steps.First, the conjugated fatty acid octadeca-cis-9,trans-11-dienoate was formed from linoleicacid through the action of an isomerase. Subse-quently, the conjugated fatty acid was reducedto octadec-trans-11-enoate (179). The isomerasewas associated with a protein- and lipid-richparticulate fraction possibly derived from a lip-


mbr.asm

.org/D

ownloaded from



oprotein layer of the protoplasmic cylinder(179). Ferredoxin may be involved in this hy-drogenation process (180).

Pickett and Kelly reported a study on themetabolism of lipids in three species of relaps-ing fever borreliae (137). Thin-layer chromatog-raphy of chloroform-methanol extracts of cul-ture supernatant liquid indicated that lysoleci-thin was removed from complex growth mediaby the growing spirochetes, whereas lecithin,sphingomyelin, triglycerides, and cholesterolesters were not affected by the growth of theborreliae. Enzymatic assays of sonic cell ex-tracts showed that lysolecithinase (EC 3.1.1.5),glycerophosphorylcholine diesterase (EC3.1.4.2), and acid phosphatase (EC 3.1.3.2) ac-tivities were present. The authors concludedthat the borreliae sequentially metabolize lyso-lecithin to fatty acids, choline, inorganic phos-phate, and glycerol. Enzymes of lecithin catab-olism were not detected, a finding in agreementwith the observation that borreliae fail to uti-lize lecithin as the sole lipid substrate in cul-ture media.

Substantial amounts of cholesterol were de-tected in cell preparations of the Reiter trepo-neme (126). This sterol was not synthesized bythe organisms but was taken up by the cellsfrom the growth medium, where it was presentas a serum constituent. It has also been re-ported that relapsing fever borreliae selectivelyremove cholesterol from the culture mediumduring growth (137). Although cholesterol ap-parently has a nutritional role for a strain of T.refringens (140), it is not known whether thissterol serves an essential function in the physi-'ology of treponemes and borreliae. Pickett andKelly (137) pointed out that nonspecific adsorp-tion of cholesterol to the cells may be responsi-ble for the removal of this compound from cul-ture media during growth of spirochetes. Var-ious free-living spirochetes have been grown inmedia from which cholesterol was absent andobviously do not have a requirement for it (27,74, 78). Meyer and Meyer (126) reported that S.zuelzerae neither synthesizes nor requires ster-ols.

Carotenoid PigmentsS. aurantia and spirochete RS1, the only

known free-living, facultatively anaerobic spi-rochetes, produce carotenoid pigments (27, 67).Aerobically grown colonies of S. aurantia areyellow-orange, whereas those of spirochete RS1are red. Anaerobically grown colonies arewhite.The molecular structure of the major pig-

ments of spirochete RS1 and S. aurantia strain

J1 was determined by analytical procedures in-volving mass spectrometry, infrared spectros-copy, chromatographic analysis, hydride reduc-tion, and acetylation and silylation experi-ments (67). It was found that the major pigmentof spirochete RS1 was 4-keto-1',2'-dihydro-1'-hydroxytorulene, also called deoxyflexixanthin(Fig. 4). This pigment accounted for at least90% of the total pigment content of spirocheteRS1. The major pigment from S. aurantia was1',2'-dihydro-1'-hydroxytorulene (Fig. 4), dif-fering from deoxyflexixanthin only in a substi-tution in the cyclohexene ring. Chromato-graphic and spectrophotometric evidence indi-cated that 1',2'-dihydro-1'-hydroxytorulenewas also present, as a minor carotenoid compo-nent, in spirochete RS1.The two major carotenoid pigments from S.

aurantia and spirochete RS1 (67) had been pre-viously detected almost exclusively in glidingbacteria, such as species ofFlexibacter (2), Stig-matella (112), and Myxococcus (142). The possi-ble evolutionary significance of the occurrenceof these pigments in both spirochetes and glid-ing bacteria is discussed in a subsequent sec-tion of this review. Saproxanthin, a carotenoidpigment remarkably similar in chemical struc-ture (Fig. 4) to the identified pigments of spiro-chete RS1 and S. aurantia, is the major carote-noid ofSaprospira grandis (1). It is noteworthythat S. grandis is a gliding bacterium previ-ously believed to be a spirochete (25, 33).

Motility and ChemotaxisSpirochetes perform locomotory, rotatory,

and flexing movements. Movements involvingflexing of the cell are quite varied and includelashing, looping, and bending motions, vibra-tions, undulations, formation of helical wavestraveling along the body, and production ofplanar waves (87, 177). Because of these mani-fold and incessant movements, spirochetes areoften described as being "extremely flexible"(e.g., see 150). Movement occurs in liquid me-dia and agar gels and is not dependent uponcontact with a solid surface. However, as previ-

A TxC N_

- ~~~~~~OH0

B

OH

C SHO OH

FIG. 4. Deoxyflexixanthin (A), 1',2'-dihydro-1'-hydroxytorulene (B), and saproxanthin (C).

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


192 CANALE-PAROLA

ously mentioned, creeping or crawling move-ment on solid surfaces has been observed (23,45).

It is widely believed or assumed that axialfibrils play a role in the movements of spiro-chetes. This belief is held mainly because au-thors who studied these organelles found themto be strikingly similar in fine structure andchemical characteristics to bacterial flagella.Individual axial fibrirs consist of three mainmorphological components structurally similarto those of bacterial flagella: (i) an insertionapparatus including a number of basal disks;(ii) a proximal hook near the insertion appara-tus; and (iii) a shaft or filamentous portion (18,19, 23, 81, 84, 86, 90, 101, 115, 128). In nega-tively stained preparations examined by elec-tron microscopy, the filamentous portion ap-pears to consist of a core enveloped by a non-striated sheath or coat (23, 81, 84, 86, 101, 128).Axial fibrils consist of one or more species ofprotein strikingly similar in overall amino acidcomposition to bacterial flagella protein (18, 20,101, 128). Furthermore, the response of purifiedaxial fibrils to chemicals or enzymes was foundto be largely similar to that reported for bacte-rial flagella (18, 20, 101, 128).Even though direct evidence is lacking, it

seems reasonable to infer that axial fibrils andbacterial flagella may serve similar or analo-gous functions because they so closely resembleeach other morphologically and chemically. Itshould be noted, however, that axial fibrils,unlike bacterial flagella, are completely endo-cellular organelles. Thus, even though axialfibrils probably play a role in motility, theirmode of action may be expected to be differentfrom that of bacterial flagella.

Antisera against axial fibrils were used todetermine whether these organelles are in-volved in motility of spirochetes (21). Since an-tisera to spirochetes are capable of immobiliz-ing them, it seemed possible that the immobi-lizing factor in antisera was directed toward theaxial fibrils. It was reasoned that if antisera topurified axial fibrils immobilized the orga-nisms, this would constitute evidence implicat-ing axial fibrils in motility. However, antiseraagainst axial fibrils purified from S. zuelzeraedid not immobilize this spirochete. In addition,it was found that the immobilizing factor pres-ent in antisera to whole cells was directed to-ward a cell component other than axial fibrils.It was suggested that the spirochetes were notimmobilized by anti-axial fibril antibodies be-cause the latter did not have access to the axialfibrils, due to the endocellular location of theseorganelles.

Other workers reported that anticell seraproduced against whole leptospires had immo-bilizing activity, which apparently resultedfrom damage to, and degradation of, the cellstructure (39). Anti-axial fibril serum hadsome, but not strong, immobilizing activityagainst leptospiral cells.Attempts have been made to interpret the

mechanisms of spirochetal movement (15, 23,45, 88, 103, 175). The movements of Cristispiraand of large free-living spirochetes were re-corded by cinematography and time-exposure,single-frame photographs by Jahn and Land-man (88). According to these authors, the spiro-chetes they studied are driven by irrotationalhelical waves which travel along the flexiblebody of the organisms. The organisms movewith almost no slippage; that is, as the spiro-chetal cell advances through a liquid, the rearcoils follow the path of the anterior tip almostperfectly. The spirochetes swim with high hy-drodynamic efficiency and have no means ofresisting the torque generated by the travelinghelical waves. In a subsequent publication,Wang and Jahn (175) presented a theory thatassumes that rotation about a local body axisoccurs and that it cancels the torque producedby the traveling helical waves.Blakemore and Canale-Parola (23) observed

and photographed moving cells of the large,free-living S. plicatilis. The cells translatedboth in contact and with no contact with solidsurfaces. During the latter type of motility, S.plicatilis cells appeared to rotate rapidly abouttheir longitudinal axis, and wide waves movedalong the length of the organisms. Further-more, the cells vigorously flexed, looped, anddarted through the liquid environment. Cellstranslating in contact with glass surfaces"crept" foward through a fixed pattern of coils,virtually without slippage.

Observations of wet-mount preparations re-vealed that cells of S. aurantia Ml usuallyswim in straight lines or nearly straight linesand, as they travel in the liquid environment,appear to spin about their longitudinal axis (E.P. Greenberg and E. Canale-Parola, unpub-lished data). Occasionally a cell stops its loco-moting and spinning, flexes its body, and thenresumes travel in a new direction. Frequently,after flexing, the cell end that was the anteriorend becomes the posterior end.Cox and Twigg (45) reported that cells of L.

interrogans serotype icterohaemorrhagiae,when moving freely in liquids, do not rotateabout their axes. According to these authors,translating leptospires are propelled by helicalwaves, which travel for a short distance from

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



the anterior cell end toward the trailing cellend. Whereas the anterior end is not hooked,the trailing end forms a broad semicircularhook. This hook waves in an approximatelycircular motion in the opposite direction to thehelical waves, thus providing stability and pre-venting rotation ofthe cell body. In nontransla-tional movement, both cell ends are hooked andwave in opposite directions, their rapid move-ment giving the illusion of spinning (45).Drugs such as procaine-hydrochloride, sero-

tonin creatinine sulfate, atropine sulfate, andothers, in a concentration range of 10-3 to 106M, affect the motility patterns of flagellatedbacteria (57, 138). It was found that these drugsalso inhibit or interfere with the movements ofleptospires (138). These observations suggestthat a common mechanism may operate in themotor control of motility in flagellated bacteriaand in spirochetes (138).As discussed in more detail below, Kaiser

and Doetsch (103) reported that translationalmovement of leptospires was strikingly en-hanced in viscous environments. Both the num-ber of leptospiral cells exhibiting translationalmovement and the velocity of the cells weremarkedly increased in viscous solutions.A comprehensive model for propulsion of spi-

rochetes has been proposed recently by Berg(15). The model is based on the assumptionsthat the protoplasmic cylinder is semirigid,that the outer sheath is flexible, and that theaxial fibrils rotate in a manner similar to thatof bacterial flagella. According to the model,the rotation ofthe axial fibrils causes the proto-plasmic cylinder to rotate within the outersheath. When the axial fibrils rotate in thesame direction, the protoplasmic cylinder ro-tates in the opposite direction. A change in therotational direction of the fibrils results in acorresponding change in the rotational direc-tion of the protoplasmic cylinder. In a spiro-chete that has two axial fibrils, when only oneof the fibrils changes direction of rotation, theprotoplasmic cylinder stops rotating. If theouter sheath is free, it would be expected torotate in a direction opposite to that of theprotoplasmic cylinder. Thus, the outer sheathbehaves like the tread of a tank, rolling aboutthe protoplasmic cylinder. When the proto-plasmic cylinder is planar, it rotates in place.When the protoplasmic cylinder is helical, thecell rotates about its longitudinal axis and hastranslational movement. According to Berg's(15) model, thrust may be generated "in twoopposing ways: by the circumferential slip ofthe helix through the medium" and "by theimbalance in the longitudinal viscous forces

due to the roll of the sheath at the outer andinner surfaces of the helix." To interpret thecreeping motility of S. plicatilis (23), Berg pro-posed that the long and irregularly shaped pro-toplasmic cylinder ofthis spirochete, when neara solid surface, may not be free to rotate. Insuch a situation "the roll of the sheath willcause the cell to slide in a direction nearlyparallel to the local helical axis" (15). Berg (15)suggested that evidence indicating whether theaxial fibrils rotate could be yielded by experi-ments similar to those carried out by Silvermanand Simon (149) to determine rotation ofEsche-richia coli flagella. Thus, the axial fibrils couldbe linked to glass or to polystyrene latex beadswith antibodies, after rupturing the spiro-chetes' outer sheath. Similar experiments withlatex beads would be useful in monitoring themotion of the outer sheath (15).As spirochetes grow in agar media, they usu-

ally move through the agar gel, forming so-called growth "veils" (36). Thus, colonies inmedia containing as much as 1% (or even more)agar tend to spread or diffuse. When cells of S.aurantia are inoculated in the center of agar(0.5%) medium plates, they migrate throughthe agar gel toward the periphery of the plates,forming characteristic concentric growth rings(30). The rings of growing cells originate fromthe inoculum in the center of the plates, in-crease in diameter during incubation, and mayreach the edge of the plates. In growth mediacontaining glucose (0.02 to 0.1%, wt/vol), thecells in the outermost ring utilize all glucoseavailable as they migrate toward the peripheryof the plate. Thus, no glucose is left in theportion of the agar plate circumscribed by thisring. Measurements of ring diameter duringincubation showed that the rate of migration ofglucose-utilizing rings is greatest at low glu-cose concentrations (30). This behavior was in-terpreted as follows. As S. aurantia cells inthese rings metabolize glucose, a glucose con-centration gradient is formed in the agar gel,and the population continuously migrates to-ward zones of higher glucose concentrationwithin the concentration gradient. There theycontinue to dissimilate glucose and, as a result,the glucose gradient shifts toward the periph-ery of the plate. At low glucose concentrationsthis shift is more rapid and, consequently, therate of population migration is greater. It wasconcluded that migration of cell populationspresent in these rings was due to a chemotacticresponse toward glucose (30).More recently, a quantitative assay was de-

veloped to investigate the chemotactic behaviorof S. aurantia Ml (Greenberg and Canale-Par-

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


194 CANALE-PAROLA

ola, Abstr. Annu. Meeting Am. Soc. Microbiol.,1976, I122, p. 131; unpublished data). The as-say, similar to that used by Adler (3) to studychemotaxis in E. coli, was conducted as follows.The open end of a microcapillary tube contain-ing an attractant in a phosphate buffer-cysteinesolution was immersed into a suspension of S.aurantia cells. After 1 h, the number of spiro-chetes that entered the microcapillary tube wasdetermined by plate counts and was comparedwith the number of spirochetes that entered anidentical microcapillary tube filled with phos-phate buffer-cysteine solution lacking the at-tractant. The presence of L-cysteine in the solu-tion enhanced the chemotactic response, forreasons not yet determined. It was found that avariety of sugars, which served as energysources for the growth of S. aurantia Ml (D-mannose, D-fructose, n-galactose, n-glucose, n-xylose, maltose, and cellobiose), also served asattractants. On the other hand, 1-mannitol,used by S. aurantia Ml as an energy source,did not serve as an attractant. Some sugarsthat were not used as energy sources (n-gluco-samine, n-fucose, 2-deoxy-n-glucose, a-methyl-n-glucoside) attracted S. aurantia Ml. This in-dicated that metabolism, and probably trans-port of the attractant, were not required forchemotaxis. Finally, other sugars not used asenergy sources (L-glucose, 6-deoxy-n-glucose, n-ribose, n-sorbitol) did not elicit a positive chem-otactic response. The threshold concentrationsfor attractants, that is, the lowest concentra-tions at which a response was observed, were aslow as 2 x 10-7. Maximum accumulation ofspirochetes in the capillary occurred at attrac-tant concentrations of 10-4 to 10-1 M. Aminoacids such as L-serine, glycine, L-aspartate, L-methionine, and others did not attract S. au-rantia Ml. These studies also showed that ga-lactose taxis in S. aurantia Ml is inducible,inasmuch as it occurs in cells grown in thepresence of n-galactose or of n-galactose and D-glucose together, but not in cells cultured with1-glucose, as energy source.

EVOLUTION OF SPIROCHETESIn attempting to interpret the evolutionary

history of spirochetes, answers to two funda-mental questions must be sought: (i) what evo-lutionary processes were responsible for theunique cellular architecture present in such aheterogeneous assemblage of bacteria?; (ii)what evolutionary steps led to the physiologicaland ecological diversity we observe in present-day spirochetes?

In this section, I shall set forth two hy-potheses that may be used to answer, in part,

the two fundamental questions. Furthermore,the hypotheses may serve as preliminary andincomplete interpretations of the evolutionarydevelopment of spirochetes.

The Protospirochete HypothesisPresumably the first cells evolved over 3 bil-

lion years ago, and during the Precambrian eraancestral procaryotic cells were selected outover their competitors (109, 120, 121). It is con-sidered probable that these primitive procar-yotes, existing in the O2-free environment ofancient Earth, were anaerobic fermentativeheterotrophs, which eventually developed theability to generate ATP by metabolizing carbo-hydrates via the EM pathway (68, 83, 109, 120,121).An obligately anaerobic, carbohydrate-fer-

menting, free-living protospirochete may haveevolved from these primitive procaryotesthrough mutations leading to morphologicaldifferentiation. Such differentiation conferredupon the developing protospirochete the traitsof "spirocheteness," that is, a helical shape,axial fibrils, and an outer sheath, as well as theunique motility mechanisms associated withthis type of cellular configuration. Spirochete-ness persisted and was retained by the ances-tral spirochetes because it offered them selec-tive advantages. Possible advantages of spiro-cheteness are discussed in the following subsec-tion of this article.

According to the protospirochete hypothesis,all spirochetes that exist today are descendantsof the protospirochete. Among modern spiro-chetes, the closest relatives of the protospiro-chete may be the free-living, obligate anaer-obes that ferment carbohydrates via the EMpathway, e.g., certain species of Spirochaeta(see section, Anaerobic energy-yielding metab-olism).

It is believed that the emergence of orga-nisms capable of using water as an electrondonor for photosynthesis resulted in the appear-ance of molecular oxygen in the Earth's atmos-phere, an event that made possible the develop-ment of aerobic energy-yielding mechanisms inliving cells (109, 120, 121). At this stage ofevolution, some of the free-living, anaerobicspirochetes, as well as many other anaerobicbacteria, may have acquired the ability to res-pire aerobically. Present-day, free-living, facul-tatively anaerobic spirochetes (e.g., S. auran-tia) may be the descendants of these first spiro-chetes capable of generating ATP by electrontransport to molecular oxygen. In time, selec-tive pressures may have led to the developmentof free-living, obligately aerobic spirochetes,

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



e.g., strains of Leptospira.A plethora of new habitats became available

to bacteria after the appearance of animals onEarth. On or within the body of these highlycomplex organisms, spirochetes establishedthemselves in a variety of habitats rangingfrom the outer surface of protozoa (24, 42, 153)to the epithelial layer of the human colon (127,164). In these habitats the spirochetes persistedand thrived, developing complex physiologicalinteractions with the host cells and with otherresident microorganisms. Most likely, fromthese ancient host-associated spirochetesevolved those present-day spirochetes that areindigenous to healthy animals, such as speciesof Treponema and Cristispira. Survival of theancestral host-associated spirochetes dependedon their being transmitted from host to host.Probably transmittal took place by mechanismssimilar to those responsible for propagation ofmodern host-associated spirochetes (26, 33, 46,47, 62, 69, 166, 167). Thus, animals were colo-nized by spirochetes through direct contactwith a host, or through contact with the host'ssecretions or excretions in which the spiro-chetes temporarily survived. Some of the host-associated spirochetes released by their hostinto aquatic environments remained viable forrelatively short periods of time in the free stateand swam to new hosts, which they colonized.Furthermore, arthropod vectors may have beenresponsible for transmittal. It is possible thatspirochetes capable of producing diseaseevolved from ancient symbiotic or commensalspirochetes that gained the ability to overcomethe natural defenses of the host.

Convergent Evolution HypothesisThe convergent evolution hypothesis states

that in the course of evolution, different procar-yotes developed the characteristics of spiro-cheteness. Accordingly, the various kinds ofspirochetes we observe today did not evolvefrom a common protospirochete ancestor butderived from procaryotes that had already be-come diversified physiologically and ecologi-cally. These procaryotes acquired spirochete-ness through independent, but converging,morphological evolutionary processes.

It is likely that selective advantages weregained by procaryotes that attained spirochete-ness. Apparently, an important advantage thatat least some of them acquired was a markedincrease in velocity of translation during move-ment through highly viscous environments.Kaiser and Doetsch have shown that spiro-chetes of the genus Leptospira exhibit maxi-mum velocity (30 gm/s) when swimming in

environments with viscosities exceeding 300centipoises (103). In contrast, certain flagel-lated bacteria, such as Spirillum serpens,which translate at a maximum velocity of 38.5pm/s at 2.5 centipoises, show a rapid decrease invelocity at higher viscosities (147). Thus, lep-tospires and possibly other spirochetes maymove rapidly through viscous natural habitats.Viscous natural habitats of spirochetes includethe crystalline style of molluscs, fluids of thegingival crevice, intracellular and intercellularregions of animal hosts, and mucosal surfaces,as well as microbial slimes and viscid mudpresent in aquatic environments inhabited byfree-living spirochetes.

Possession of an outer sheath may also confera selective advantage to spirochetes. This struc-ture may act as a permeability barrier, whichprotects the axial fibrils from disruptive ordamaging environmental agents such as pHextremes or the action of enzymes secreted byother microorganisms. As previously men-tioned, Bharier and Rittenberg reported thatwhereas antisera against whole cells immobi-lize S. zuelzerae cells, antisera against axialfibrils do not (21). In light of this report, itseems possible that the outer sheath serves as ashield against external agents that may affectthe functioning of the axial fibrils. If, indeed,axial fibrils play a role in motility, as is sus-pected, the benefits of this type of protection tothe spirochetes are evident.The traits of spirocheteness, acquired by cells

in the course of evolution, have persisted, andpresent-day procaryotes that possess such traitsare quite successful in terms of competitionwith other microorganisms. Thus, it seemsprobable that in addition to the selective advan-tages suggested above, spirocheteness impartsto cells advantages that are not obvious to us atpresent.

Presently available experimental evidencedoes not contradict the convergent evolutionhypothesis or the protospirochete hypothesis,but it is too meager to favor either hypothesisover the other. The finding that a wide range ofguanine plus cytosine contents occurs in thedeoxyribonucleic acid of spirochetes (Table 1)does not necessarily support the convergent ev-olution hypothesis. On the contrary, it mayindicate that the protospirochete originatedearly in the evolutionary history of procaryotesand changes in the deoxyribonucleic acid oc-curred subsequently.

Energy-Yielding Pathways of SpirochetesAs pointed out by Hall (68), a useful approach

toward the construction of a phylogenetic tree

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


196 CANALE-PAROLA

for procaryotes is the comparison of physiologi-cal characteristics of existing microorganisms.Thus, analysis of available information on theenergy-yielding pathways of spirochetes mayserve to uncover fragments of the develop-mental history of these organisms and to sug-gest possible evolutionary processes.

It is considered likely that the EM pathwaywas the first, or among the first, of the pres-ently known fermentative pathways to evolve(83). The EM pathway is present in all free-living and host-associated spirochetes thathave been assayed for it. Anaerobic and facul-tatively anaerobic spirochetes ferment carbohy-drates via this pathway. As previously men-tioned, in free-living anaerobes and facultativeanaerobes, such as species of Spirochaeta, theEM pathway is coupled to a clostridial-typepyruvate clastic reaction (Fig. 2). Through thisreaction Spirochaeta species cleave pyruvate toCO2, a two-carbon fragment used for -acetateand ethanol production, and electrons, whichare transferred to the hydrogenase system (Fig.2). Furthermore, at least some anaerobic host-associated.spirochetes, e.g., T. denticola, use aclostridial-type cleavage to metabolize pyru-vate derived from carbohydrates via the EMpathway or from amino acids (Fig. 3). Thus, theevidence indicates that ATP generationthrough the EM pathway coupled to a clos-tridial-type cleavage of pyruvate is a commoncharacteristic among anaerobic and faculta-tively anaerobic spirochetes. In view of itswidespread occurrence, it may be speculatedthat this was the earliest or most primordialtype of energy-yielding metabolism available tospirochetes.

Investigations of the energy-yielding path-ways in the host-associated T. denticola indi-cated that whereas this spirochete has con-served the ancestral properties of fermentingcarbohydrates via the EM pathway and of me-tabolizing pyruvate through a clostridial clasticsystem, it has acquired the ability to catabolizea variety of amino acids. Some of these, e.g.,cysteine or serine, are fermented through path-ways in which pyruvate is a key intermediate(Fig. 3). Other amino acids, such as arginine orcitrulline, are dissimilated via pathways thatdo not involve pyruvate (Fig. 3). Thus, T. denti-cola has a more complex system of energy-yielding pathways than the free-living anaero-bic spirochetes. Competition for substrates inthe gingival crevice must be acute, consideringthat the microbial population in this region ofthe human body averages 1.3 x 1011 organismsper g (wet weight) of gingival debris, as deter-mined by microscopic counts (157). The meta-

bolic versatility of T. denticola offers to thisorganism a distinct selective advantage in thisdensely populated habitat.On the basis of the limited available informa-

tion, a possible developmental pattern may besuggested for the energy-yielding pathways ofanaerobic spirochetes. It appears that the free-living anaerobic spirochetes have not signif'i-cantly modified their ATP-generating mecha-nisms since primordial times. However, at leastsome of the anaerobic spirochetes that havebecome adapted to life in specialized host-asso-ciated habitats have evolved additional ATP-yielding pathways, probably as a result of selec-tive pressures. As mentioned in a precedingsection, the anaerobic dissimilatory pathwaysof facultatively anaerobic spirochetes are essen-tially identical to those of the free-living obli-gate anaerobes (Fig. 2). In air, the facultativeanaerobe S. aurantia performs an incompleteoxidation of carbohydrates mainly to CO2 andacetate. It does not possess a tricarboxylic acidcycle, but it has developed mechanisms of oxi-dative phosphorylation and a rudimentary elec-tron transport system, involving one or twocytochromes. In contrast, the leptospires,which are the only known obligately aerobicspirochetes, do not dissimilate carbohydrates.These spirochetes generate ATP by oxidizinglong-chain fatty acids to two-carbon fragments,which are channeled into the tricarboxylic acidcycle. The leptospires have enzymes of the EMpathway but apparently use this pathway onlyin a biosynthetic direction. Furthermore, theelectron transport of leptospires, which in-cludes cytochromes of the a, c, c,, and o types, ismore complex than that of S. aurantia. Inas-much as the facultatively anaerobic spirocheteshave developed aerobic electron transport sys-tems that enable them to derive energythrough oxidative phosphorylation, it may beinferred that they represent an evolutionaryform more advanced than the obligately anaer-obic free-living forms. On the other hand, thelack of a tricarboxylic acid cycle and the posses-sion of a rudimentary electron transport systemapparently places the facultatively anaerobicspirochetes on a lower evolutionary level thanthe leptospires, which have more efficient aero-bic ATP-generating systems.

It cannot be excluded that at least some of theexisting free-living spirochetes were derivedfrom host-associated spirochetal ancestors,which, after being released from their animalhosts, succeeded in becoming adapted to life inthe free state. It seems unlikely, however, thatthis is or was a major evolutionary route forspirochetes, inasmuch as experimental evi-

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



dence has shown that host-associated spiro-chetes are defective in various biosyntheticabilities. As discussed in the next section,growth and survival of these "defective" spiro-chetes depends largely on metabolites orgrowth factors readily available to them intheir host-associated environment. The defec-tive forms, separated from their host, would notbe expected to compete successfully with micro-organisms well adapted to a free-living exis-tence.

Development of Associations with HostsLoss of the ability to synthesize a compound

required for growth does not prevent microor-ganisms from multiplying, as long as they canobtain that particular compound from the envi-ronment. When such a condition is met, theseauxotrophic or defective mutants, lacking a bio-synthetic function, may have a selective advan-tage over parental strains requiring a greaternumber ofenergy-consuming biosynthetic stepsfor growth (181). Thus, it is not surprising thatspirochetes indigenous to nutrient-rich habi-tats within higher organisms are defective incertain biosynthetic abilities. For example, asmentioned above, many spirochetes associatedwith humans and animals are unable to syn-thesize long-chain fatty acids, which they re-quire to manufacture cellular lipids. In the en-vironments inhabited by these spirochetes,fatty acids are available, being produced by thehost's metabolism and by other resident micro-organisms. In contrast, free-living spirochetes,such as species of Spirochaeta, retain the abil-,ity to synthesize long-chain fatty acids, pre-sumably because these compounds are notreadily available to them in their habitats.Among other required substances not synthe-sized by host-associated spirochetes are thia-mine pyrophosphate and N-acetylglucosamine(106, 129, 130, 162).An enzymatic activity absent from many

host-associated spirochetes, probably becauseof mutations resulting in evolutionary selec-tion, is the production of H2 (31, 60, 79, 114). Onthe other hand, all known free-living anaerobicspirochetes synthesize hydrogenase systemsand produce H2 (see section, Anaerobic energy-yielding metabolism). Unless anaerobic host-associated spirochetes compensate for the lackof hydrogenase by manufacturing other elec-tron-accepting systems, they may require anddepend on electron acceptors present in theirnatural habitats or growth media.

In conclusion, it is. likely that the loss ofcertain biosynthetic abilities is one of the fac-tors responsible for the development of spiro-

chetes restricted to life within specialized, host-associated habitats. This loss is followed byevolutionary selection favoring the auxotrophicmutants over the parental strains.Some spirochetes have developed special cel-

lular structures by which they are attached toeucaryotic cells. Thus, spirochetes are anchoredto the surface ofprotozoa by a "rootlet" or "nose-like specialization" of their proximal cell end(24, 153). The intimate physical association be-tween these spirochetes, or intestinal spiro-chetes (127, 164), and host cells strongly sug-gests the existence of symbiotic interactionsbetween the animals and the attached mi-crobes. Spirochetes attached to eucaryotic cellsmay be dependent on these interactions for sur-vival.

Possible Relationship with Gliding BacteriaSome recent reports suggest that the phylo-

genetic relationship between spirochetes andgliding bacteria may be closer than previouslybelieved. Greenberg and Canale-Parola (67) de-termined the molecular structure of the majorcarotenoid pigments of the facultative anaer-obes S. aurantia and spirochete RS1 (Fig. 4),the only known pigmented spirochetes. Exceptfor their presence in the spirochetes, and in anunidentified microorganism (4), these carote-noids have been detected only in gliding bacte-ria (see section, Carotenoid pigments).Another characteristic common to the facul-

tatively anaerobic spirochetes and gliding bac-teria is that their growth is strongly inhibitedby actinomycin D (49, 67). Since gliders andspirochetes are both gram negative, this is anunusual response, inasmuch as gram-negativebacteria generally are not appreciably in-hibited by actinomycin D (49). Possibly, thesensitivity of gliding bacteria and spirochetesto this antibiotic reflects similarities in cellsurface composition or in specific physiologicalprocesses.

Studies of motility mechanisms may revealadditional similarities between spirochetes andgliding bacteria. As previously mentioned, spi-rochetes not only swim free-floating in liquids,but also "creep" or "crawl" on solid surfaces (23,45). The mechanism responsible for the lattertype of movement may prove to be identical orsimilar to that which propels gliding bacteria.

Spirochetes, Eucaryotic Flagella, and CiliaIt has been suggested that spirochetes, or

spirochete-like organisms, might have been theprecursors of eucaryotic flagella and cilia. Thissuggestion is part of a theory interpreting theevolutionary origin of eucaryotic cells in terms

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


198 CANALE-PAROLA

of a series of symbioses (120). According to thistheory, primitive amoebae were formed as theresult of a symbiotic association between ananaerobic pleomorphic microbe and a smalleraerobic procaryote. The anaerobic organismserved as the host, contributing nuclear mate-rial and cytoplasm to the association, and itharbored the aerobe (the endosymbiont), whichlater evolved into mitochondria. Still accordingto the theory, the next major step in the forma-tion of present-day eucaryotic cells occurredwhen free-living, motile, spirochete-like orga-nisms became attached to the surface of themitochondria-containing amoebae. The ecto-symbiotic spirochete-like organisms, whichwere attached by one of their cell poles to theancestral amoebae, conferred motility to thecomplex, presumably by undulating in a coordi-nated manner. The symbiotic theory states thatthe eucaryotic flagellum and cilium, with its 9+ 2 fibrillar arrangement, evolved from thesespirochete-like organisms.The participation of spirochetes in evolution-

ary processes leading to the formation of eucar-yotic cells has been suggested by the fact thatassociations analogous to that postulated be-tween spirochete-like organisms and the ances-tral amoebae occur among modern microorga-nisms. Spirochetes are present on the surface ofcertain protozoa, to which they adhere by anextremity of their cells (24, 42, 111, 153). Forexample, spirochetes occur on the surface ofMixotricha paradoxa, a large protozoon foundin the gut of the termite Mastotermes darwin-iensis. The coordinated undulations of thou-sands of spirochetes attached to each individualMixotricha cell propel the protozoon uninter-ruptedly and at constant speed (42).

Concluding RemarksIt is apparent that the available information

on the evolutionary history of spirochetes ismeager. Some of the reports mentioned in thisarticle suggest various evolutionary possibili-ties, but conclusions cannot be reached at pres-ent. This is not surprising, inasmuch as weknow little about the biology of spirochetes.Although these bacteria are widespread, only asmall fraction of the many kinds of spirochetesobserved in natural habitats has been culti-vated. Even among those spirochetes that havebeen cultivated, relatively few have been stud-ied extensively.

Interpretable fossil records of spirocheteshave not been found. Thus, investigations onthe evolution of spirochetes must proceed inother directions. One possible approach is thestudy of the biochemical and physiological

characteristics of different kinds of spirochetes.For example, phylogenetic relationships maybe clarified through comparative studies onmetabolic pathways, on motility and chemo-taxis mechanisms, and on amino acid se-quences of spirochetal proteins such as rubre-doxin and ferredoxin (91).

SummarySpirochetes are bacteria with unique mor-

phology and motility mechanisms. However,they do not constitute a homogeneous bacterialgroup but exhibit extreme physiological andecological diversity. All existing spirochetesmay have evolved from an ancestral protospiro-chete whose descendants underwent extensivephysiological differentiation. Possibly, amongpresent-day spirochetes, the free-living, carbo-hydrate-fermenting, obligate anaerobes are themost direct descendants of the protospirochete,whereas free-living aerobic and facultativelyanaerobic forms, and spirochetes indigenous toanimals and humans, developed through fur-ther evolutionary processes.

According to another evolutionary hypothe-sis, the multifarious types of modern spiro-chetes did not evolve from a common protospi-rochete ancestor but from a number of procar-yotes different physiologically and ecologicallyfrom one another. These diverse procaryotesacquired the characteristics of spirocheteness,which conferred to them selective advantages.

Analysis of published information on thephysiology of spirochetes suggests that ATPgeneration from carbohydrate dissimilation viathe EM pathway, coupled to a clostridial-typecleavage of pyruvate, was the earliest kind ofenergy-yielding metabolism available to all ormany of these bacteria. Subsequently, in re-sponse to selective pressures, some spirochetesacquired additional ATP-yielding mechanisms,such as amino acid catabolism or oxidativephosphorylation. Other spirochetes either di-verged radically from the ancestral type ofATP-yielding metabolism or never possessed it,their only energy source being respiration cou-pled to oxidation of a few fatty acids and alco-hols.

Spirochetes restricted to life within special-ized host-associated habitats may have devel-oped from the free-living forms as a result ofevolutionary selection after losing certain en-ergy-consuming biosynthetic functions, e.g.,the ability to synthesize long-chain fatty acids,other cellular constituents, and metabolites.Recent reports suggest that at least some spiro-chetes and gliding bacteria may be phylogeneti-cally closer than previously believed.

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



ACKNOWLEDGMENTSInvestigations by the author on the physiology

and morphology of spirochetes were supported byPublic Health Service grants AI-08248 and AI-12482from the National Institute of Allergy and In-fectious Diseases.

LITERATURE CITED1. Aasen, A. J., and S. Liaaen-Jensen. 1966. The

carotenoids of Flexibacteria. fl. A new xanth-ophyll from Saprospira grandis. Acta Chem.Scand. 20:811-819.

2. Aasen, A. J., and S. Liaaen-Jensen. 1966. Ca-rotenoids of Flexibacteria. III. The structureof flexixanthin and deoxy-flexixanthin. ActaChem. Scand. 20:1970-1988.

3. Adler, J. 1973. A method for measuring chemo-taxis and use of the method to determineoptimum conditions for chemotaxis byEsche-richia coli. J. Gen. Microbiol. 74:77-91.

4. Aguilar-Martinez, M., and S. Liaaen-Jensen.1972. Bacterial carotenoids. XL. 2'-Hydroxy-flexixanthin. Acta Chem. Scand. 26:2528-2530.

5. Ajello, F. 1969. Activation of acetate in thetreponemes (pathogenic pallidum and non-pathogenic Reiter and Kazan treponemes).G. Microbiol. 17:107-114.

6. Alexander, A. D. 1970. Leptospira, p. 244-250.In J. E. Blair, E. H. Lennette, and J. P.Truant (ed.), Manual of clinical microbiol-ogy, 1st ed. American Society for Microbiol-ogy, Bethesda, Md.

7. Allen, S. L., R. C. Johnson, and D. Peterson.1971. Metabolism of common substrates bythe Reiter strain of Treponema palladium.Infect. Immun. 3:727-734.

8. Barban, S. 1954. Studies on the metabolism ofthe Treponemata. I. Amino acid metabolism.J. Bacteriol. 68:493-497.

9. Barban, S. 1956. Studies on the metabolism ofthe Treponemata. II. Transamination in theReiter treponeme. J. Bacteriol. 71:274-277.

10. Baseman, J. B., and C. D. Cox. 1969. Interme-diate energy metabolism of Leptospira. J.Bacteriol. 97:992-1000.

11. Baseman, J. B., and C. D. Cox. 1969. Terminalelectron transport in Leptospira. J. Bacte-riol. 97:1001-1004.

12. Baseman, J. B., R. C. Henneberry, and C. D.Cox. 1966. Isolation and growth ofLeptospiraon artificial media. J. Bacteriol. 91:1374-1375.

13. Baseman, J. B., J.C. Nichols, and N. S. Hayes.1976. Virulent Treponema pallidum: aerobeor anaerobe. Infect. Immun. 13:704-711.

14. Baseman, J. B., J. C. Nichols, J. W. Rumpp,and N. S. Hayes. 1974. Purification of Trepo-nema pallidum from infected rabbit tissue:resolution into two treponemal populations.Infect. Immun. 10:1062-1067.

15. Berg, H. C. 1976. How spirochetes may swim.J. Theor. Biol. 56:269-273.

16. Berg, R. N., S. S. Green, H. S. Goldberg, andD. C. Blenden. 1969. Density-gradient and

chromatographic fraction of leptospiral li-pase. Appl. Microbiol. 17:467-472.

17. Bert6k, L., and F. Kemenes. 1960. Studies onthe lipase system of leptospirae. I. Tributyri-nase activity. Acta Microbiol. Acad. Sci.Hung. 7:251-259.

18. Bharier, M., and D. Allis. 1974. Purificationand characterization of axial filaments fromTreponema phagedenis biotype reiterii (theReiter treponeme). J. Bacteriol. 120:1434-1442.

19. Bharier, M. A., F. A. Eierling, and S. C.Rittenberg. 1971. Electron microscopic ob-servations on the structure of Treponemazuelzerae and its axial filaments. J. Bacte-riol. 105:413-421.

20. Bharier, M. A., and S. C. Rittenberg. 1971.Chemistry of axial filaments of Treponemazuelzerae. J. Bacteriol. 105:422-429.

21. Bharier, M. A., and S. C. Rittenberg. 1971.Immobilization effects of anticell and antiax-ial filament sera on Treponema zuelzerae. J.Bacteriol. 105:430-437.

22. Biggins, J., and W. E. Dietrich, Jr. 1968. Res-piratory mechanisms in the Flexibacteri-aceae. I. Studies on the terminal oxidase sys-tem of Leucothrix mucor. Arch. Biochem.Biophys. 128:40-50.

23. Blakemore, R. P., and E. Canale-Parola. 1973.Morphological and ecological characteristicsof Spirochaeta plicatilis. Arch. Mikrobiol.89:273-289.

24. Bloodgood, R. A., K. R. Miller, T. P. Fitzhar-ris, and J. R. McIntosh. 1974. The ultra-structure of Pyrsonympha and its associatedmicroorganisms. J. Morphol. 143:77-105.

25. Breed, R. S., E. G. D. Murray, and N. R.Smith (ed.). 1957. Bergey's manual of deter-minative bacteriology, 7th ed. The Williams& Wilkins Co., Baltimore.

26. Breznak, J. A. 1973. Biology of nonpathogenic,host-associated spirochetes. Crit. Rev. Mi-crobiol. 2:457-489.

27. Breznak, J. A., and E. Canale-Parola. 1969.Spirochaeta aurantia, a pigmented, faculta-tively anaerobic spirochete. J. Bacteriol.97:386-395.

28. Breznak, J. A., and E. Canale-Parola. 1972.Metabolism of Spirochaeta aurantia. I. An-aerobic energy-yielding pathways. Arch.Mikrobiol. 83:261-277.

29. Breznak, J. A., and E. Canale-Parola. 1972.Metabolism of Spirochaeta aurantia. I.Aerobic oxidation of carbohydrates. Arch.Mikrobiol. 83:278-292.

30. Breznak, J. A., and E. Canale-Parola. 1975.Morphology and physiology of Spirochaetaaurantia strains isolated from aquatic habi-tats. Arch. Microbiol. 105:1-12.

31. Bryant, M. P. 1952. The isolation and charac-teristics of a spirochete from the bovine ru-men. J. Bacteriol. 64:325-335.

32. Buchanan, B. B., and D. I. Arnon. 1970. Ferre-doxins: chemistry and function in photosyn-thesis, nitrogen fixation, and fermentative

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


200 CANALE-PAROLA

metabolism. Adv. Enzymol. 33:119-176.33. Buchanan, R. E., and N. E. Gibbons (ed.).

1974. Bergey's manual of determinative bac-teriology, 8th ed. The Williams & WilkinsCo., Baltimore.

34. Burnett, G. W., and H. W. Scherp. 1968. Oralmicrobiology and infectious disease, 3rd ed.The Williams & Wilkins Co., Baltimore.

35. Burton, G., D. C. Blenden, and H. S. Gold-berg. 1970. Naphthylamidase activity ofLep-tospira. Appl. Microbiol. 19:586-588.

36. Canale-Parola, E. 1973. Isolation, growth andmaintenance of anaerobic free-living spiro-chetes, p. 61-73. In J. R. Norris and D. W.Ribbons (ed.), Methods in microbiology, vol.8. Academic Press Inc., New York.

37. Canale-Parola, E., S. C. Holt, and Z. Udris.1967. Isolation of free-living, anaerobic spiro-chetes. Arch. Mikrobiol. 59:41-48.

38. Canale-Parola, E., Z. Udris, and M. Mandel.1968. The classification of free-living spiro-chetes. Arch. Mikrobiol. 63:385-397.

39. Chang, A., and S. Faine. 1973. Effect of anti-cell and anti-axial filament sera on Leptos-pira. Aust. J. Exp. Biol. Med. Sci. 51:847-856.

40. Charon, N. W., R. C. Johnson, and D. Peter-son. 1974. Amino acid biosynthesis in thespirochete Leptospira: evidence for a novelpathway of isoleucine biosynthesis. J. Bacte-riol. 117:203-211.

41. Chorvath, B., and P. Bakoss. 1972. Studies onleptospiral lipase. II. Lipase activity of viru-lent and avirulent leptospirae. J. Hyg. Epi-demiol. Microbiol. Immunol. 16:352-357.

42. Cleveland, L. R., and A. V. Grimstone. 1964.The fine structure of the flagellate Mixotri-cha paradoxa and its associated microorga-nisms. Proc. R. Soc. London Ser. B 159:668-686.

43. Cohen, P. G., C. W. Moss, and D. Farshtch.1970. Cellular lipids of treponemes. Br. J.Vener. Dis. 46:10-12.

44. Cox, C. D., and M. K. Barber. 1974. Oxygenuptake by Treponema pallidum. Infect. Im-mun. 10:123-127.

45. Cox, P. J., and G. I. Twigg. 1974. Leptospiralmotility. Nature (London) 250:260-261.

46. Crawford, R. P., J. M. Heinemann, W. F.McCulloch, and S. L. Diesch. 1971. Humaninfections associated with waterborne leptos-pires, and survival studies on serotype po-mona. J. Am. Vet. Med. Assoc. 159:1477-1484.

47. Davis, B. D., R. Dulbecco, H. N. Eisen, H. S.Ginsberg, and W. B. Wood. 1973. Microbiol-ogy. Harper and Row, Hagerstown, Md.

48. Doelle, H. W. 1975. Bacterial metabolism, 2nded., p. 465, 649. Academic Press Inc., NewYork.

49. Dworkin, M. 1969. Sensitivity of gliding bacte-ria to actinomycin D. J. Bacteriol. 98:851-852.

50. Ehrenberg, C. G. 1835. Dritter Beitrag zur Er-kenntnis grosser Organisation in der Rich-tung des kleinsten Raumes. Abh. Dtsch.

Akad. Wiss. Berlin Kl. Chem., Geol. Biol., p.313.

51. Ellinghausen, H. C., and W. G. McCullough.1965. Nutrition of Leptospira pomona andgrowth of thirteen other serotypes: a serum-free medium employing oleic albumin com-plex. Am. J. Vet. Res. 26:39-44.

52. Ellinghausen, H. C., and W. G. McCullough.1965. Nutrition of Leptospira pomona andgrowth of thirteen other serotypes: fractiona-tion of oleic albumin complex and a mediumof bovine albumin and polysorbate 80. Am. J.Vet. Res. 26:45-51.

53. Ellinghausen, H. C., and 0. Sandvik. 1965.Tributyrinase activity of leptospires: fixedand soluble tributyrinase demonstrated bymeans of an agar diffusion test. Acta Pathol.Microbiol. Scand. 65:259-270.

54. Eskow, R. N., and W. J. Loesche. 1971. Oxy-gen tensions in the human oral cavity. Arch.Oral Biol. 16:1127-1128.

55. Faine, S. 1960. Catalase activity in pathogenicLeptospira. J. Gen. Microbiol. 22:1-9.

56. Faure, M., and J. Pillot. 1960. Compositionantigenique des treponemes. III. L'antigenelipidique de Wassermann. Ann. Inst. Pas-teur Paris 99:729-742.

57. Faust, M. A., and R. N. Doetsch. 1971. Effect ofdrugs that alter excitable membranes on themotility of Rhodospirillum rubrum andThiospirillum jenense. Can. J. Microbiol.17: 191-196.

58. Fitzgerald, T. J., J. N. Miller, and J. A. Sykes.1975. Treponema pallidum (Nichols strain)in tissue cultures: cellular attachment, en-try, and survival. Infect. Immun. 11:1133-1140.

59. Fulghum, R. S., and W. E. C. Moore. 1963.Isolation, enumeration, and characteristicsof proteolytic ruminal bacteria. J. Bacteriol.85:808-815.

60. Fulton, J. D., and P. J. C. Smith. 1960. Carbo-hydrate metabolism in Spirochaeta recurren-tis. I. The metabolism of spirochaetes in vivoand in vitro. Biochem. J. 76:491-499.

61. Fulton, J. D., and D. F. Spooner. 1956. Themetabolism of Leptospira icterohaemorrhag-iae in vitro. Exp. Parasitol. 5:154-177.

62. Gillespie, R. W. H., and J. Ryno. 1963. Epide-miology of leptospirosis. Am. J. PublicHealth 53:950-955.

63. Goldberg, H. S., and J. C. Armstrong. 1959.Oxidase reaction with leptospiral coloniesand its adaptation to antibiotic sensitivitytesting. J. Bacteriol. 77:512-513.

64. Graves, S. R., P. L. Sandok, H. M. Jenkin, andR. C. Johnson. 1975. Retention of motilityand virulence of Treponema pallidum (Ni-chols strain) in vitro. Infect. Immun.12:1116-1120.

65. Green, S. S., and H. S. Goldberg. 1967. Electro-phoretic determination of leptospiral en-zymes. J. Bacteriol. 93:1739-1740.

66. Green, S. S., H. S. Goldberg, and D. C. Blen-den. 1967. Enzyme patterns in the study ofLeptospira. Appl. Microbiol. 15:1104-1113.

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



67. Greenberg, E. P., and E. Canale-Parola. 1975.Carotenoid pigments of facultatively anaero-bic spirochetes. J. Bacteriol. 123:1006-1012.

68. Hall, J. B. 1971. Evolution of the prokaryotes.J. Theor. Biol. 30:429-454.

69. Hanson, A. W. 1970. Isolation of spirochaetesfrom primates and other mammalian spe-cies. Br. J. Vener. Dis. 46:303-306.

70. Hardy, P. H., Jr. 1970. Borrelia, p. 236-239. InJ. E. Blair, E. H. Lennette, and J. P. Truant(ed.), Manual ofclinical microbiology, 1st ed.American Society for Microbiology, Be-thesda, Md.

71. Hardy, P. H., Jr., W. R. Fredericks, and E. E.Nell. 1975. Isolation and antigenic character-istics of axial filaments from the Reiter tre-poneme. Infect. Immun. 11:380-386.

72. Hardy, P. H., Jr., and C. 0. Munro. 1966.Nutritional requirements of anaerobic spiro-chetes. I. Demonstration of isobutyrate andbicarbonate as growth factors for a strain ofTreponema microdentium. J. Bacteriol.91:27-32.

73. Helprin, J. J., and C. W. Hiatt. 1957. The effectof fatty acids on the respiration ofLeptospiraicterohemorrhagiae. J. Infect. Dis. 100:136-140.

74. Henneberry, R. C., J. B. Baseman, and C. D.Cox. 1970. Growth of a water strain of Lep-tospira in synthetic media. Antonie van

Leeuwenhoek J. Microbiol. Serol. 36:489-501.

75. Henneberry, R. C., and C. D. Cox. 1970. (-

Oxidation of fatty acids by Leptospira. Can.J. Microbiol. 16:41-45.

76. Henry, R. A., R. C. Johnson, B. B. Bohlool,and E. L. Schmidt. 1971. Detection ofLeptos-pira in soil and water by immunofluoresencestaining. Appl. Microbiol. 21:953-956.

77. Hespell, R. B., and E. Canale-Parola. 1970.Carbohydrate metabolism in Spirochaetastenostrepta. J. Bacteriol. 103:216-226.

78. Hespell, R. B., and E. Canale-Parola. 1970.Spirochaeta litoralis sp.n., a strictly anaero-bic marine spirochete. Arch. Mikrobiol.74:1-18.

79. Hespell, R. B., and E. Canale-Parola. 1971.Amino acid and glucose fermentation by Tre-ponema denticola. Arch. Mikrobiol. 78:234-251.

80. Hespell, R. B., and E. Canale-Parola. 1973.Glucose and pyruvate metabolism of Spiro-chaeta litoralis, an anaerobic marine spiro-chete. J. Bacteriol. 116:931-937.

81. Holt, S. C., and E. Canale-Parola. 1968. Finestructure of Spirochaeta stenostrepta, a free-livig, anaerobic spirochete. J. Bacteriol.96:822-835.

82. Horvath, I., R. J. Arko, and J. C. Bullard.1975. Effects of oxygen and nitrogen on thecharacter of T. pallidum in subcutaneouschambers in mice. Br. J. Vener. Dis. 51:301-304.

83. Horvath, R. S. 1974. Evolution of anaerobicenergy-yielding metabolic pathways of theprocaryotes. J. Theor. Biol. 47:361-371.

84. Hovind-Hougen, K., and A. Birch-Andersen.1971. Electron microscopy of endoflagellaand microtubules in Treponema Reiter. ActaPathol. Microbiol. Scand. Sect. B 79:37-50.

85. Hughes, R., H. J. Olander, and C. B. Williams.1975. Swine dysentery: pathogenicity of Tre-ponema hyodysenteriae. Am. J. Vet. Res.36:971-977.

86. Jackson, S., and S. H. Black. 1971. Ultrastruc-ture of Treponema pallidum Nichols follow-ing lysis by physical and chemical methods.II. Axial filaments. Arch. Mikrobiol. 76:325-340.

87. Jahn, T. L., and E. C. Bovee. 1965. Movementand locomotion of microorganisms. Annu.Rev. Microbiol. 19:21-58.

88. Jahn, T. L., and M. D. Landman. 1965. Loco-motion of spirochetes. Trans. Am. Microsc.Soc. 84:395-406.

89. Jenkin, H. M., L. E. Anderson, R. T. Holman,I. A. Ismail, and F. D. Gunstone. 1969. Ef-fect of isomeric cis-octadecenoic acids on thegrowth of Leptospira interrogans serotypepatoc. J. Bacteriol. 98:1026-1029.

90. Jepsen, 0. B., K. Hovind-Hougen, and A.Birch-Andersen. 1968. Electron microscopyof Treponema pallidum Nichols. Acta Pa-thol. Microbiol. Scand. 74:241-258.

91. Johnson, P. W., and E. Canale-Parola. 1973.Properties of rubredoxin and ferredoxin iso-lated from spirochetes. Arch. Mikrobiol.89:341-353.

92. Johnson, R. C., and L. M. Eggebraten. 1971.Fatty acid requirements of the Kazan 5 andReiter strains of Treponema pallidum. In-fect. Immun. 3:723-726.

93. Johnson, R. C., and N. D. Gary. 1963. Nutri-tion of Leptospira pomona. H. Fatty acid re-quirements. J. Bacteriol. 85:976-982.

94. Johnson, R. C., and V. G. Harris. 1968. Purineanalogue sensitivity and lipase activity ofleptospires. Appl. Microbiol. 16:1584-1590.

95. Johnson, R. C., V. G. Harris, and J. K. Walby.1969. Characterization of leptospires accord-ing to fatty acid requirements. J. Gen. Mi-crobiol. 55:399-407.

96. Johnson, R. C., B. P. Livermore, H. M. Jen-kin, and L. Eggebraten. 1970. Lipids of Tre-ponema pallidum Kazan 5. Infect. Immun.2:606-609.

97. Johnson, R. C., B. P. Livermore, J. K. Walby,and H. M. Jenkin. 1970. Lipids of parasiticand saprophytic leptospires. Infect. Immun.2:286-291.

98. Johnson, R. C., and J. K. Walby. 1972. Culti-vation of leptospires: fatty acid require-ments. Appl. Microbiol. 23:1027-1028.

99. Jones, R. H., M. A. Finn, J. J. Thomas, and C.Folger. 1976. Growth and subculture ofpath-ogenic T. pallidum (Nichols strain) in BHK-21 cultured tissue cells. Br. J. Vener. Dis.52:18-23.

100. Joseph, R. 1972. Fatty acid composition ofSpi-rochaeta stenostrepta. J. Bacteriol. 112:629-631.

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


202 CANALE-PAROLA

101. Joseph, R., and E. Canale-Parola. 1972. Axialfibrils of anaerobic spirochetes: Ultrastruc-ture and chemical characteristics. Arch.Mikrobiol. 81:146-168.

102. Kadis, S., and W. L. Pugh. 1974. Urea utiliza-tion by Leptospira. Infect. Immun. 10:793-801.

103. Kaiser, G. E., and R. N. Doetsch. 1975. En-hanced translational motion ofLeptospira inviscous environments. Nature (London)255:656-657.

104. Kasarov, L. B., and L. Addamiano. 1969. Me-tabolism of the lipoproteins of serum by lep-tospires: degradation of the triglycerides. J.Med. Microbiol. 2:165-168.

105. Kawata, T. 1967. Electron transport system inthe cell fraction of the Reiter treponeme withspecial reference to the presence of cyto-chromes. Jpn. J. Bacteriol. (Nippon Saikin-gaku Zasshi) 22:590-594.

106. Kelly, R. 1971. Cultivation of Borrelia hermsi.Science 173:443-444.

107. Kelly, R. T. 1974. Borrelia, p. 355-357. In E. H.Lennette, E. H. Spaulding, and J. P. Truant(ed.), Manual of clinical microbiology, 2nded. American Society for Microbiology,Washington, D.C.

108. Kemenes, F., and L. Lovrekovich. 1959. Uberden Feltabbau durch Pathogene Leptospi-ren. Acta Vet. Acad. Sci. Hung. 9:235-241.

109. Kenyon, H., and G. Steinman. 1969. Biochemi-cal predestination. McGraw-Hill, New York.

110. Kinyon, J. M., and D. L. Harris. 1974. Growthof Treponema hyodysenteriae in liquid me-dium. Vet. Rec., Sept. 7, p. 219-220.

111. Kirby, H., Jr. 1941. Organisms living on and inprotozoa, p. 1009-1113. In G. N. Calkins andF. M. Summers (ed.), Protozoa in biologicalresearch. Columbia University Press, NewYork.

112. Kleinig, H., and H. Reichenbach. 1969. Carot-enoid pigments of Stigmatella aurantiaca(Myxobacterales). I. The minor carotenoids.Arch. Mikrobiol. 68:210-217.

113. Kondo, E., and N. Ueta. 1972. Composition offatty acids and carbohydrates in Leptospira.J. Bacteriol. 110:459-467.

114. Krichevsky, M. I., and E. G. Hampp. 1966.Metabolic gas production by a variety of spi-rochetes. J. Dent. Res. 45:165-168.

115. Listgarten, M. A., and S. S. Socransky. 1964.Electron microscopy of axial fibrils, outerenvelope, and cell division of certain oralspirochetes. J. Bacteriol. 88:1087-1103.

116. Livermore, B. P., and R. C. Johnson. 1970.Isolation and characterization of a glyco-lipid from Treponema pallidum, Kazan 5.Biochim. Biophys. Acta 210:315-318.

117. Livermore, B. P., and R. C. Johnson. 1974.Lipids of the Spirochaetales: comparison ofthe lipids of several members of the generaSpirochaeta, Treponema, and Leptospira. J.Bacteriol. 120:1268-1273.

118. Livermore, B. P., and R. C. Johnson. 1975.The lipids of four unusual non-pathogenic

host-associated spirochetes. Can. J. Micro-biol. 21:1877-1880.

119. Lovenberg, W., and B. E. Sobel. 1965. Rubre-doxin: a new electron transfer protein fromClostridium pasteurianum. Proc. Natl.Acad. Sci. U. S. A. 54:193-199.

120. Margulis, L. 1970. Origin of eukaryotic cells.Yale University Press, New Haven, Conn.

121. Margulis, L. 1971. Microbial evolution on theearly earth, p. 480-484. In R. Buvet and C.Ponnamperuma (ed.), Chemical evolutionand the origin of life. North-Holland Pub-lishing Co., Amsterdam.

122. Markovetz, A. J., and A. D. Larson. 1959.Transamination in Leptospira biflexa. Proc.Soc. Exp. Biol. Med. 101:638-640.

123. Marshall, P. B. 1949. Measurement of aerobicrespiration in Leptospira icterohaemorrhag-iae. J. Infect. Dis. 84:150-152.

124. Metzger, M., and W. Smog6r. 1966. Study ofthe effect of pH and Eh values of the Nelson-Diesendruck medium on the survival of viru-lent Treponema pallidum. Arch. Immunol.Ther. Exp. 14:445-453.

125. Meyer, H., and F. Meyer. 1969. Biosynthesis ofunsaturated fatty acids by a free-living spi-rochete. Biochim. Biophys. Acta 176:202-204.

126. Meyer, H., and F. Meyer. 1971. Lipid metabo-lism in the parasitic and free-living spiro-chetes Treponema pallidum (Reiter) andTreponema zuelzerae. Biochim. Biophys.Acta 231:93-106.

127. Minio, F., G. Tonietti, and A. Torsoli. 1973.Spontaneous spirochete infestation in the co-lonic mucosa of healthy men. Rend. Gas-troenterol. 5:183-195.

128. Nauman, R. K., S. C. Holt, and C. D. Cox.1969. Purification, ultrastructure, and com-position of axial filaments from Leptospira.J. Bacteriol. 98:264-280.

129. Nevin, T. A., and E. G. Hampp. 1959. Partiallydefined medium for the cultivation ofBorre-lia vincentii. J. Bacteriol. 78:263-266.

130. Nevin, T. A., E. G. Hampp, and B. V. Duey.1960. Interaction between Borrelia vincentiiand an oral diphtheroid. J. Bacteriol. 80:783-786.

131. Nichols, J. C., and J. B. Baseman. 1975. Car-bon sources utilized by virulent Treponemapallidum. Infect. Immun. 12:1044-1050.

132. Oyama, V. I., H. G. Steinman, and H. Eagle.1953. The nutritional requirements of Trepo-nemata. V. A detoxified lipide as the essen-tial growth factor supplied by crystallized se-rum albumin. J. Bacteriol. 65:609-616.

133. Parnas, J., A. Koslak, and M. Krukowska.1960. Untersuchungen der Leptospirenli-pase. Zentralbl. Bakteriol. Parasitenkd. In-fektionskr. Hyg. Abt. 1 Orig. 180:386-389.

134. Patel, V., H. S. Goldberg, and D. Blenden.1964. Characterization of leptospiral lipase.J. Bacteriol. 88:877-884.

135. Peterson, J. A., D. Basu, and M. J. Coon. 1966.Enzymatic w-oxidation. I. Electron carriers

BACTERIOL. REV.


mbr.asm

.org/D

ownloaded from



in fatty acid and hydrocarbon hydroxylation.J. Biol. Chem. 241:5162-5164.

136. Peterson, J. A., M. Kusunose, E. Kusunose,and M. J. Coon. 1967. Enzymatic w-oxida-tion. II. Function of rubredoxin as the elec-tron carrier in w-hydroxylation. J. Biol.Chem. 242:4334-4340.

137. Pickett, J., and R. Kelly. 1974. Lipid catabo-lism of relapsing fever borreliae. Infect. Im-mun. 9:279-285.

138. Pitney, S. N., and R. N. Doetsch. 1971. Effectof pharmacologically active drugs on the mo-tility ofLeptospira interrogans (biflexa). LifeSci. 10:1237-1246.

139. Pittman, K. A., S. Lakshmanan, and M. P.Bryant. 1967. Oligopeptide uptake by Bacte-roides ruminicola. J. Bacteriol. 93:1499-1508.

140. Rajkovic, A. D. 1967. Cultivation of the Nogu-chi strain of Treponema pallidum in a lipidmedium in the absence of serum. Z. Med.Mikrobiol. Immunol. 153:297-312.

141. Rao, P. J., A. D. Larson, and C. D. Cox. 1964.Catalase activity in Leptospira. J. Bacteriol.88:1045-1048.

142. Reichenbach, H., and H. Kleinig. 1971. Thecarotenoids of Myxococcus fulvus (Myxobac-terales). Arch. Mikrobiol. 76:364-380.

143. Sachan, D. S. 1974. Sodium requiring and avi-din sensitive oxaloacetate decarboxylase inglucose grown Treponema (Borrelia) strainB25. Biochem. Biophys. Res. Commun.60:70-75.

144. Sachan, D. S., and C. L. Davis. 1969. Hydro-genation of linoleic acid by a rumen spiro-chete. J. Bacteriol. 98:300-301.

145. Sandok, P. L., H. M. Jenkin, S. R. Graves, andS. T. Knight. 1976. Retention of motility ofTreponema pallidum (Nichols virulentstrain) in an anaerobic cell culture systemand in a cell-free system. J. Clin. Microbiol.3:72-74.

146. Scheuerbrandt, G., H. Goldfine, P. E. Baron-owsky, and K. Bloch. 1961. A novel mecha-nism for the biosynthesis of unsaturatedfatty acids. J. Biol. Chem. 236:PC70-PC71.

147. Schneider, W. R., and R. N. Doetsch. 1974.Effect of viscosity on bacterial motility. J.Bacteriol. 117:696-701.

148. Shenberg, E. 1967. Growth of pathogenic Lep-tospira in chemically defined media. J. Bac-teriol. 93:1598-1606.

149. Silverman, M., and M. Simon. 1974. Flagellarrotation and the mechanism of bacterial mo-tility. Nature (London) 249:73-74.

150. Skerman, V. B. D. 1967. A guide to the identi-fication of the genera ofbacteria, 2nd ed. TheWilliams & Wilkins Co., Baltimore.

151. Smibert, R. M. 1973. Spirochaetales, a review.Crit. Rev. Microbiol. 2:491-552.

152. Smibert, R. M., and R. L. Claterbaugh. 1972.A chemically defined medium for Treponemastrain PR-7 isolated from the intestine of apig with swine dysentery. Can. J. Microbiol.18:1073-1078.

153. Smith, H. E., and H. J. Arnott. 1974. Epi- andendobiotic bacteria associated with Pyrson-ympha vertens, a symbiotic protozoon of thetermite Reticulitermes flavipes. Trans. Am.Microsc. Soc. 93:180-194.

154. Smith, P. J. C. 1960. Carbohydrate metabolismin Spirochaeta recurrentis. 2. Enzymes asso-ciated with disintegrated cells and extractsof spirochaetes. Biochem. J. 76:500-508.

155. Smith, P. J. C. 1960. Carbohydrate metabolismin Spirochaeta recurrentis. 3. Properties ofaldolase in spirochaetes. Biochem. J. 76:508-514.

156. Smith, P. J. C. 1960. Carbohydrate metabolismin Spirochaeta recurrentis. 4. Some proper-ties of hexokinase and lactic dehydrogenasein spirochaetes. Biochem. J. 76:514-520.

157. Socransky, S. S. 1970. Relationship of bacteriato the etiology of periodontal disease. J.Dent. Res. 49:203-222.

158. Socransky, S. S., M. Listgarten, C. Hubersak,J. Cotmore, and A. Clark. 1969. Morphologi-cal and biochemical differentiation of threetypes of small oral spirochetes. J. Bacteriol.98:878-882.

159. Socransky, S. S., W. J. Loesche, C. Hubersak,and J. B. MacDonald. 1964. Dependency ofTreponema microdentium on other oral orga-nisms for isobutyrate, polyamines, and acontrolled oxidation-reduction potential. J.Bacteriol. 88:200-209.

160. Stalheim, 0. H. V., and J. B. Wilson. 1964.Cultivation of leptospirae. I. Nutrition ofLeptospira canicola. J. Bacteriol. 88:48-54.

161. Staneck, J. L., R. C. Henneberry, and C. D.Cox. 1973. Growth requirements of patho-genic leptospira. Infect. Immun. 7:886-897.

162. Steinman, H. G., V. I. Oyama, and H. 0.Schulze. 1954. Carbon dioxide, cocarboxyl-ase, citrovorum factor, and coenzyme A asessential growth factors for a saprophytictreponeme (S-69). J. Biol. Chem. 211:327-335.

163. Stern, N., E. Shenberg, and A. Tietz. 1969.Studies on the metabolism of fatty acids inLeptospira: the biosynthesis of A9- and All-monounsaturated acids. Eur. J. Biochem.8:101-108.

164. Takeuchi, A., and J. A. Zeller. 1972. Ultra-structural identification of spirochetes andflagellated microbes at the brush border ofthe large intestinal epithelium of the rhesusmonkey. Infect. Immun. 6:1008-1018.

165. Tauber, H., G. R. Cannefax, A. W. Hanson,and H. Russell. 1962. Enzymes of trepo-nemes. A comparative study. Exp. Med.Surg. 20:324-328.

166. Turner, L. H. 1967. Leptospirosis I. Trans. R.Soc. Trop. Med. Hyg. 61:842-855.

167. Turner, L. H. 1970. Leptospirosis III. Trans. R.Soc. Trop. Med. Hyg. 64:623-646.

168. Ueda, T., E. T. Lode, and M. J. Coon. 1972.Enzymatic w-oxidation. VI. Isolation of ho-mogeneous reduced diphosphopyridine nu-cleotide-rubredoxin reductase. J. Biol.

VOL. 41, 1977


mbr.asm

.org/D

ownloaded from


204 CANALE-PAROLA

Chem. 247:2109-2116.169. Vaczi, L., K. Kiraly, and A. Rethy. 1966. Lipid

composition of treponemal strains. Acta Mi-crobiol. Acad. Sci. Hung. 13:79-84.

170. VanEseltine, W. P., and S. A. Staples. 1961.Nutritional requirements of leptospirae. I.Studies on oleic acid as a growth factor for astrain of Leptospira pomona. J. Infect. Dis.108:262-269.

171. Veldkamp, H. 1960. Isolation and characteris-tics of Treponema zuelzerae nov. spec., ananaerobic, free-living spirochete. Antonievan Leeuwenhoek J. Microbiol. Serol.26:103-125.

172. Vogel, H., and S. H. Hutner. 1961. Growth ofLeptospira in defined media. J. Gen. Micro-biol. 26:223-230.

173. Wahren, A., and T. Holme. 1973. Amino acidand peptide requirement ofFusiformis necro-phorus. J. Bacteriol. 116:279-284.

174. Wallace, A. L., and A. Harris. 1967. Reiter-treponeme.'Bull. W.H.O. 36(Suppl. 2):1-103.

175. Wang, C.-Y., and T. L. Jahn. 1972. A theory forthe locomotion of spirochetes. J. Theor. Biol.36:53-60.

176. Wegner, G. H., and E. M. Foster. 1963. Incor-poration of isobutyrate and valerate into cel-lular plasmalogen by Bacteroides succino-

genes. J. Bacteriol. 85:53-61.177. Weibull, C. 1960. Movement, p. 153-205. In I.

C. Gunsalus and R. Y. Stanier (ed.), Thebacteria, vol. 1. Academic Press Inc., NewYork.

178. Woratz, H. 1957. Wachstumsversuche mit Fett-sauren an Leptospira canicola. Zentralbl.Bakteriol. Parasitenkd. Infektionskr. Hyg.Abt. 1 Orig. 169:269-274.

179. Yokoyama, M. T., and C. L. Davis. 1971. Hy-drogenation of unsaturated fatty acids byTreponema (Borrelia) strain B25, a rumenspirochete. J. Bacteriol. 107:519-527.

180. Yokoyama, M. T., and C. L. Davis. 1971. Stim-ulation of hydrogenation of linoleate in Tre-ponema (Borrelia) sp. strain B25 by reducedmethyl viologen and by reduced benzyl violo-gen. Biochem. J. 125:913-915.

181. Zamenhof, S., and H. H. Eichhorn. 1967.Study of microbial evolution through loss ofbiosynthetic functions: establishment of "de-fective" mutants. Nature (London) 216:456-458.

182. Ziolecki, A., H. Tomerska, and M. Wojcie-chowicz. 1975. Characteristics of pure cul-tures ofBorrelia sp. isolated from the bovinerumen. Acta Microbiol. Pol. Ser. B 7:45-50.

BACTERIOL. Rzv.


mbr.asm

.org/D

ownloaded from


Date post:	24-Sep-2020
Category:	Documents
Upload:	others
View:	1 times
Download:	0 times

Physiology andEvolution ofSpirochetes · spirochetes indigenous to healthy humans or animals...

Documents