Molecular Biology of Spirochetes

MOLECULAR BIOLOGY OF SPIROCHETES

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Series I. Life and Behavioural Sciences – Vol. 373

ISSN: 1566-7693

Molecular Biology of Spirochetes

Edited by

Felipe C. Cabello

New York Medical College, Valhalla, New York, USA

Dagmar Hulinska

Czech Republic National Institute of Public Health, Prague, Czech Republic

and

Henry P. Godfrey

New York Medical College, Valhalla, New York, USA

Amsterdam • Berlin • Oxford • Tokyo • Washington, DC Published in cooperation with NATO Public Diplomacy Division

Proceedings of the NATO Advanced Research Workshop on Molecular Biology of Spirochetes Prague, Czech Republic 5–8 December 2005 © 2006 IOS Press. All rights reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, without prior written permission from the publisher. ISBN 1-58603-665-3 Library of Congress Control Number: 2006933007 Publisher IOS Press Nieuwe Hemweg 6B 1013 BG Amsterdam Netherlands fax: +31 20 687 0019 e-mail: [email protected] Distributor in the UK and Ireland Gazelle Books Services Ltd. White Cross Mills Hightown Lancaster LA1 4XS United Kingdom fax: +44 1524 63232 e-mail: [email protected]

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LEGAL NOTICE The publisher is not responsible for the use which might be made of the following information. PRINTED IN THE NETHERLANDS

Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Preface This book is a compilation of presentations at a NATO Advanced Research Workshop on the “Molecular Biology of Spirochetes” held at the Czech Republic National Institute of Public Health, Prague, Czech Republic, December 5–8, 2005. This meeting was supported by the NATO Programme for Security Through Science, the United States National Institutes of Health, Office of Rare Diseases, the United States Institute of Allergy and Infectious Diseases, the Czech Republic National Institute of Public Health, Czech Republic and New York Medical College, Valhalla, New York, U.S.A. It was organized to foster the exchange of experience among scientists from NATO countries in North America, Western and Eastern Europe. This type of encounter is valuable because diseases produced by spirochetes, including Lyme borreliosis, syphilis and leptospirosis, are on the rise worldwide, and because the biology of their causative organisms, their epidemiology, and clinical presentation display important variations in different geographical areas. For example B. burgdorferi sensu lato produces approximately 20,000 cases of Lyme borreliosis a year in the United States and 60,000 cases in Europe, but B. burgdorferi sensu stricto, B. afzelii and B. garinii are transmitted by different vectors and have different reservoirs and clinical presentations in these different geographic areas. Awareness and better understanding of these variations by researchers in the field is thus highly relevant to improvements in their prevention and treatment, and critical for improvement of human health. The meeting was organized with oral presentations by major speakers and poster sessions by students and postdoctoral fellows from Eastern Europe. This volume includes not only the presentations of the major speakers but also several additional presentations by investigators who were invited but were unable to attend. For many reasons (including meeting organization and funding limitations), this volume does not intend to represent a comprehensive coverage of all aspects of spirochete biology. It rather focuses on a series of state of the art presentations of the research taking place in the laboratories of the contributors. As such, we hope that it may be useful as an introduction to those individuals entering in the burgeoning field of spirochete research. We would also like to believe that the meeting and this book will serve as a stimulus for researchers in the field to widen collaborations and exchanges between investigators in the different geographical areas where spirochetal diseases are common since these interactions can only be of benefit to the field. Finally, we would like to thank the participants who risked the cold weather to attend the meeting, the authors who despite their inability to attend were willing to submit chapters to this book, the funding institutions mentioned above, and in particular, Drs. Phil Baker, Patti Rosa, Tom Schwan, Frank Gherardini, Ms. Marylin E. Kunzweiler (United States National Institute of Allergy and Infectious Diseases), Dr. Jorge Benach (Stony Brook University), Ms. Mary C. Demory (United States National Institutes of Health), Dr. Marina Cinco (University of Trieste), Dr. Michael Norgard (University of Texas Southwestern Medical Center) and Dr. Ira Schwartz (New York Medical College), whose efforts were critical to our securing some of the

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funds that supported the meeting, and TestLine, Qiagen, BioConsult, Roche, Biowestern, Generi BioTech and BagMed who provided additional support for this meeting. We would also like to thank Ms. Leonor Delgado for editorial assistance and Ms. Harriett V. Harrison for her outstanding and continuing assistance in organizing the meeting and in preparing the manuscripts that compose this book. Felipe C. Cabello New York Medical College Valhalla, New York, USA Dagmar Hulinska Czech Republic National Institute of Public Health Prague, Czech Republic Henry P. Godfrey New York Medical College Valhalla, New York, USA July, 2006

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Contents Preface Felipe C. Cabello, Dagmar Hulinska and Henry P. Godfrey

v

Introductory Overview Dissemination and Persistence are Pathogenic Events Common to All of the Major Human Spirochetal Infections Gary P. Wormser

3

Part 1. Molecular Genetics of Spirochetes Transposon Mutagenesis of Infectious Borrelia burgdorferi B31: A Pilot Study Douglas J. Botkin, April Abbott, Jerrilyn K. Howell, Mary Mosher, Philip E. Stewart, Patricia A. Rosa, Hiroki Kawabata, Haruo Watanabe and Steven J. Norris

The Isolation and Characterization of Isogenic Mutants in Infectious Borrelia burgdorferi J. Seshu, Maria Labandeira-Rey, M. Dolores Esteve-Gassent, Magnus Höök and Jonathan T. Skare Motility Gene Regulation and Chemotaxis in Borrelia burgdorferi Nyles W. Charon, Melanie Sal, Michael R. Miller, Richard G. Bakker, Chunhao Li and Md. Abdul Motaleb

Targeted and Random Mutagenesis in Leptospira biflexa: Application for the Functional Analysis of Iron Transporters Hélène Louvel, Simona Bommezzadri, Nora Zidane, Paula Ristow, Zoé Rouy, Claudine Medigue, Caroline Boursaux-Eude, Isabelle Saint Girons, Christiane Bouchier and Mathieu Picardeau

Antibiotic Resistance in Borrelia burgdorferi: Applications for Genetic Manipulation and Implications for Evolution D. Scott Samuels Development of Treponeme Genetic Systems Howard K. Kuramitsu and Caroline E. Cameron

13

25 42

50

56 71

Part 2. Genomics and Diversity Comparative Genomics of Borrelia burgdorferi Sherwood R. Casjens, Wai Mun Huang, Eddie B. Gilcrease, Weigang Qiu, William D. McCaig, Benjamin J. Luft, Steven E. Schutzer and Claire M. Fraser

79

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Treponema Genomics George M. Weinstock, David Šmajs, Petra Matĕjková, Michal Strouhal, Thomas J. Albert, Steven J. Norris, Timothy Palzkill and Erica J. Sodergren Comparative Analysis of Pathogenic Leptospira Genomes Richard L. Zuerner, Dieter M. Bulach, Torsten Seemann, Ross L. Coppel and Ben Adler Leptospira interrogans: Genomics and “Immunomics” Ana L.T.O. Nascimento

Genotypic Variation and Borrelia burgdorferi Pathogenesis Ira Schwartz, Guiqing Wang, Radha Iyer, Caroline Ojaimi, Darya Terekhova, Sabina Sandigursky, Gary P. Wormser and Dionysios Liveris

Multilocus Sequence Analysis (MLSA) as an Alternative to Whole DNA/DNA Hybridization (WDDH) in Borrelia burgdorferi sensu lato Taxonomy Guy Baranton and Danièle Postic

Diversity and Variability of Protein-Encoding Genes of Borrelia burgdorferi sensu lato and Implications for Pathogenesis and Diagnosis of Lyme Borreliosis in Europe Bettina Wilske, Volker Fingerle and Ulrike Schulte-Spechtel Are Borrelia recurrentis and Borrelia duttonii the Same Spirochaete? Sally J. Cutler, Julie C. Scott and David J.M. Wright Genotyping of Borrelia burgdorferi sensu lato in Russia Edward I. Korenberg, Valentina V. Nefedova, Irina A. Fadeeva and Nataliya B. Gorelova

Ecological and Genetic Diversity Within the Leptospiraceae Family: Implications for Epidemiology Yulia V. Ananyina, Anna P. Samsonova, Evgeny M. Petrov, Igor A. Shaginyan, Marina Yu. Chernukha, Marina S. Zemskaya and Yulia S. Alyapkina

Characterization of Borrelia burgdorferi sensu lato from Czech Patients and Ticks by Culture and PCR-Sequence Analysis Dagmar Hulinska, Martin Bojar and Václav Hulinsky

Infection of Ixodid Ticks, Mosquitoes and Patients with Borrelia, Bartonella, Rickettsia, Anaplasma, Ehrlichia and Babesia in Western Siberia, Russia Olga Morozova, Vera Rar, Yana Igolkina, Andrey Dobrotvorsky, Igor Morozov and Felipe C. Cabello

96 101 115 124

135

146 159 174

200

208 221

Part 3. Gene Expression Genetic Studies of the Borrelia burgdorferi bmp Gene Family Felipe C. Cabello, Lidiya Dubytska, Anton V. Bryksin, Julia V. Bugrysheva and Henry P. Godfrey

235

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Porins of Borrelia Marija Pinne, Yngve Östberg, Roland Benz and Sven Bergström

250

Use of Green Fluorescent Protein Transcriptional Reporters to Study Differential Gene Expression by Borrelia burgdorferi Christian H. Eggers, Melissa J. Caimano and Justin D. Radolf

264

The Telomere Resolvase ResT and Evolution of the Borrelia Genomes George Chaconas

292

Regulation of Expression of the Integrin Ligand P66 in Borrelia burgdorferi Melisa S. Medrano, Paul Policastro, Tom G. Schwan and Jenifer Coburn

Hairpin Telomeres of Linear Bacterial Chromosomes and Plasmids: How to Make Them Wai Mun Huang, Qiurong Ruan and Sherwood R. Casjens

281

299

Part 4. Interactions of Spirochetes and Hosts Blood-Induced Transcriptional Changes in Borrelia burgdorferi Rafal Tokarz and Jorge L. Benach

Roles of Leptospiral Outer Membrane Proteins in Pathogenesis and Immunity David A. Haake

Genetic Analysis of Attachment of Borrelia burgdorferi to Host Cells and Extracellular Matrix Nikhat Parveen and John M. Leong

Borrelia burgdorferi and Ixodes scapularis: Exploring the Pathogen-Vector Interface Utpal Pal, John F. Anderson and Erol Fikrig The Lyme Disease Spirochete Erp Protein Family: Structure, Function and Regulation of Expression Brian Stevenson, Tomasz Bykowski, Anne E. Cooley, Kelly Babb, Jennifer C. Miller, Michael E. Woodman, Kate von Lackum and Sean P. Riley

Lyme Disease Spirochetes Evade Innate Immunity by Acquisition of Complement Regulators, Factor H, and FHL-1 Reinhard Wallich, Peter F. Zipfel, Christine Skerka, Michael Kirschfink, Markus M. Simon, Brian Stevenson, Susan M. Lea and Peter Kraiczy

Outer Surface Lipoproteins of Borrelia burgdorferi: Role in Virulence, Persistence of the Pathogen, and in Protection Against Lyme Disease Markus M. Simon, Nico Birkner, Rinus Lamers and Reinhard Wallich

311 323 333 345 354

373

383

Localization of Lyme Disease and Relapsing Fever Spirochetes in Mammalian Hosts Infected with Different Borrelia Species, Strains, and Serotypes Diego Cadavid

393

Author Index

399

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Introductory Overview

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Dissemination and Persistence are Pathogenic Events Common to All of the Major Human Spirochetal Infections Gary P. WORMSER 1 Division of Infectious Diseases, Department of Medicine of New York Medical College, Valhalla, NY 10595 Abstract. In all of the major human spirochetal infections, the fundamental pathogenetic event underlying the most serious complications of these diseases is documented or presumed hematogenous dissemination of the spirochete from the site of inoculation to distant sites. Lyme borreliosis is attractive for study of spirochetal dissemination for a variety of reasons including: the availability of a large patient base, the ability to identify patients early during the course of infection, and the ability to culture Lyme borrelia readily in vitro. Tick feeding per se does not directly lead to blood stream invasion by Lyme borrelia. The genotype of the strain of Borrelia burgdorferi introduced by the tick, however, does appear to be an important determinant of dissemination in humans and other mammals. Recent evidence suggests that host factors affect both the development of infection and subsequent dissemination of this spirochete as well. In one United States study, independent risk factors for hematogenous dissemination of B. burgdorferi included having a first episode of Lyme borreliosis and being more than 55 years of age. In another United States study, infection due to the least invasive genotype of B. burgdorferi was associated with carriage of the HLA class II allele DRB1*0101. All of the major human spirochetal infections are also characterized by persistence of the spirochete in mammalian hosts, and depending on the specific spirochete and host involved, this phenomenon may be closely linked to the pathogenesis of important clinical manifestations and to communicability to uninfected hosts. In conclusion, spirochetemia and persistence are common and important pathogenetic features of the major human spirochetal infections. Keywords. Borrelia burgdorferi, Lyme disease, spirochetes, dissemination, persistence

Spirochetes are important causes of infection throughout the world. What might be regarded as the major spirochetal infections of humans are listed in Table 1 [1]. An estimated 12 million cases of syphilis occur annually worldwide with the majority of cases occurring in developing countries [2]. In industrialized countries, Lyme borreliosis is also a common spirochetal infection, at least in parts of North America and Europe [3, 4].

1 Corresponding Author: New York Medical College, Division of Infectious Diseases, Munger Pavilion, Room 245, Valhalla, NY 10595, USA; E-mail: [email protected].

G.P. Wormser / Dissemination and Persistence are Pathogenic Events

4

Spirochetal infections have quite diverse modes of transmission. For example, syphilis is spread predominantly through intimate physical contact between humans, whereas Lyme borreliosis can only be acquired through the bite of an infected tick. Clinical manifestations are also diverse with prominent cutaneous abnormalities in syphilis [5], the endemic treponematoses [6], Lyme disease [7] and rat bite fever due to Spirillum minus [8]. In contrast, cutaneous lesions are usually absent in leptospirosis [9] and relapsing fever [10]. All of these infections, however, share certain features central to the pathogenesis of their most serious complications or that play a pivotal role in communicability and spread of infection. Documented or presumed spirochetemia is a fundamental aspect of the pathobiology of these infections (Table 1). In syphilis, for example, if Treponema pallidum were confined to the site of entry into the skin, then even in untreated patients, this infection would be a short-lived, self-limited, almost trivial skin condition rather than a multisystem disease associated with substantial morbidity and even mortality [11, 12].

Table 1. Bacteremia and persistence in the major human spirochetal infections. Infection

Bacteremia

Persistence

Syphilis

Yes

Yes

Yaws

Yes

Yes

Endemic syphilis

Yes

Yes

Pinta

Yes

Yes

Leptospirosis

Yes

Yes (especially kidney)

Relapsing fever

Yes

Yes

Lyme disease

Yes

Yes

Rat bite fever

Yes

Yes

Another important feature that the major spirochetal infections share is the ability to persist in an untreated host (Table 1). Differences do exist among the listed spirochetal infections as to duration and sites of persistence; in some cases, such as leptospirosis, persistence occurs principally in non-human rather than human hosts [9]. One of the great enigmas concerning the persistence of these predominantly extracellular bacteria is that it may occur despite an intense humoral and cellular immune response to the microorganism. Among the major human spirochetal infections, syphilis has been the best documented and may be the only one capable of persisting for as long as decades [5]. Obviously the impact of persistence on the natural history of syphilis and the other major spirochetal infections has been dramatically altered by the timely use of antibiotic therapy to treat these infections. No convincing evidence exists for persistence of any of these infections after appropriate antibiotic therapy [13]. Potential mechanisms for persistence include location in an anatomic sanctuary protected from the sterilizing effects of the immune response, antigenic variation or modulation by the spirochete to avoid the immune response, antigenic concealment

G.P. Wormser / Dissemination and Persistence are Pathogenic Events

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through lack of expression of surface antigens or acquisition of an outer coating of host-derived materials such that key surface antigens are prevented from being recognized by the immune system, or immunomodulation of the host by the spirochete such that an effective immune response is abrogated [12, 14-18]. Antigenic variation seems to be the mechanism for persistence adopted by the relapsing fever spirochetes [19]. Leptospira have found a sanctuary in the urinary tract [9, 20]. The mechanism(s) for persistence in the other major spirochetal infections is less well understood and conceivably could be multifactorial with the spirochetes adopting more than one of the strategies discussed above. The reader is referred elsewhere for reviews of much of the available information on how T. pallidum [12] or Lyme borrelia [14] might persist. Most of the serious consequences of human spirochetal infections arise from spread from the site of entry in the skin or mucous membranes to visceral locations. Despite the obvious importance of clarifying the natural history and mechanisms for hematogenous dissemination, this issue has been difficult to study directly due to a number of logistical issues, such as the inability to cultivate many of the spirochetal agents in vitro. Lyme borreliosis is attractive for study of hematogenous dissemination because the infection can be identified clinically in its early stages based on the characteristic appearance of the cutaneous lesion (called erythema migrans) that occurs at the site of deposition of the spirochete into the skin, or even earlier if the patient were to recognize and remove an attached infected tick. Lyme borrelia can be grown successfully in vitro, and a sufficiently large number of patients with this infection exist to conduct meaningful clinical research studies.

Table 2. Yield of blood cultures in representative studies in which a low volume of blood was cultured. Author

Year

Material

Volume

Patients

Yield

USA Benach [21]

1983

Citrated blood

4 mL

36

5.6%

Steere [22]

1984

Citrated plasma

<1 mL

65

3.1%

Berger [23]

1994

Whole blood

<1 mL

52

3.8%

Goodman [24]

1995

EDTA blood and components

<1 mL

74

5.3%

Maraspin [25]

2001

Citrated plasma

1-3 mL

28

1.2%

Arnez [26]

2001

Citrated plasma

0.5 mL

134

9.0% 1

Oksi [27]

2001

NA 2

NA

74

8.1%

EUROPE

1 2

Only children studied. NA = Not available.

Until recently the relative importance of the hematogenous route of dissemination of Lyme borrelia was not well established, because the yield of blood cultures was very low, typically around 5% (Table 2) [21-27]. To the contrary, direct spread through the soft tissues was supported by certain observational and experimental data. In one study

G.P. Wormser / Dissemination and Persistence are Pathogenic Events

6

of 12 dogs which were fed upon by infected ticks, it was noticed that the first episode of lameness developed in the joint closest to the tick bite site in all of the dogs [28]. A random distribution of infection of different joints would have been anticipated if the spirochete had traveled to the joint space through the blood stream. In a separate study of humans with neuroborreliosis, the presence of erythema migrans on an extremity was also topographically associated with motor paresis of that particular extremity [29]. In prior attempts to culture borrelia from the blood of patients with Lyme borreliosis only a very small quantity of blood or a blood component was cultured, typically ” 1 mL (Table 2). To improve the yield of blood cultures in this infection, several studies were carried out in the United States of adult Lyme borreliosis patients with erythema migrans. In these studies the blood component (whole blood vs. serum vs. plasma) and/or the volume of material that was cultured, were varied in a systematic manner [30-32]. These studies showed that the yield from plasma exceeded that from serum or whole blood and that increasing the volume of plasma cultured beyond 9 mL only marginally improved the rate of recovery of Borrelia burgdorferi. The recovery rate of B. burgdorferi from • 9 mL of plasma in adult patients with erythema migrans approached 50% [31, 32]. This high yield was recently confirmed by another group of investigators in the United States using the same culture method [33]. The results of culturing plasma semi-quantitatively were consistent with an estimate of 0.1 cultivable spirochete per milliliter of whole blood, which would explain why culturing larger volumes provided a superior yield [32]. A > 40% prevalence of spirochetemia in cutaneous Lyme borrelia infection contrasts markedly with the approximately 2% rate of blood culture positivity in patients with community acquired bacterial cellulitis [34], underscoring the particular significance of hematogenous dissemination in the natural history of Lyme borreliosis.

Table 3. Genomic subgroups (genotypes) of Lyme borrelia recovered from skin or blood at New York Medical College from 1991 to 2002. 1 Material Cultured

Number of Isolates (%) RST1

RST2

RST3

P Value 2

Skin (n=293)

81 (28%)

129 (44%)

83 (28%)

---

Blood (n=127)

53 (42%)

57 (45%)

17 (13%)

<0.001

1

Schwartz I (Unpublished data, March, 2006). 2 Frequency distributions compared by chi square test.

These observations raise a number of important questions about spirochetemia in Lyme borreliosis, such as: what host or spirochetal factors predispose to blood stream invasion, when during the course of early infection does it occur and are there associated clinical or laboratory features that might help to identify it. Several studies in both humans and mice have demonstrated the differential pathogenicity of genomic subgroups (genotypes) of Lyme borrelia [35-40]. Thus, hematogenous dissemination appears to be dependent at least in part on the specific strain of spirochete causing the infection. One way to differentiate strains of B. burgdorferi is through the use of a polymerase chain reaction (PCR)-restriction fragment length polymorphism (RFLP)

G.P. Wormser / Dissemination and Persistence are Pathogenic Events

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typing method targeting the 16S-23S rDNA spacer region [41]. Using this technique B. burgdorferi can be separated into three genotypes arbitrarily called RST1, RST2, and RST3. Based on this classification scheme, it has been demonstrated that cutaneous RST3 infections are less likely to be associated with spirochetemia compared with RST1 or RST2 infections [35]. Our cumulative results as of the end of 2002 on typing of blood or skin isolates of B. burgdorferi, excluding mixed infections with more than one RST type, is found in Table 3. The frequency distribution for the genotypes recovered from skin differs significantly from the distribution of genotypes among the blood isolates, with RST2 strains predominating in both skin and blood (p<0.001) [I. Schwartz, unpublished data]. RST3 strains are the least likely to disseminate hematogenously. In an attempt to understand better the natural history and predisposing factors for spirochetemia, we recently analyzed the clinical and laboratory characteristics of 213 adult New York State patients with erythema migrans for whom a high-volume (• 9 mL) plasma blood culture was performed prior to initiation of antibiotic therapy [42]. Spirochetemic patients were more likely to be symptomatic (89.2% vs. 74.2%, p=0.006) and to have multiple erythema migrans lesions (i.e., 2 or more lesions) (41.9% vs. 15.0%, p <0.001) compared to patients with negative blood cultures. Nevertheless, spirochetemic patients had few of the features typically associated with bacteremia due to more conventional pathogens. The vast majority of Lyme borreliosis patients (94.6%) were afebrile at the time the blood culture was obtained, 59.1% had no history of recent fever or chills, and 98.9% did not have an elevated white blood cell count. Indeed, 22.8% of the spirochetemic patients had only a single erythema migrans lesion and no systemic symptoms at all. Of interest, the frequency of obtaining a positive blood culture did not vary significantly according to either the size (though > 16 cm in largest diameter) or duration (through > 21 days) of the erythema migrans skin lesions for the entire group of patients and separately for those with single or multiple skin lesions [42]. This pattern of blood culture positivity, which was independent of size or duration of the erythema migrans skin lesion, invites the question of whether spirochetemia might even precede the onset of the skin lesion. Although never studied, this is certainly plausible, since it is well established in syphilis that spirochetemia may antedate the primary skin lesion (chancre) in that infection [11,12]. In the study of spirochetemic patients discussed above [42], a multivariate analysis revealed that only 5 clinical or laboratory features were independently associated with spirochetemia (Table 4). The findings suggested that age less than 55 years or having recovered from a prior episode of Lyme disease is protective against blood stream invasion with B. burgdorferi. Thus, in addition to genotypic differences in the infecting strains of Lyme borrelia, host factors also appear to play a critical role with regard to whether dissemination of this spirochete will occur. In a recent study on potential human host factors in the pathogenesis of Lyme borreliosis, the question of whether certain HLA class II alleles might influence susceptibility to infection by the different genotypes of B. burgdorferi was investigated in 89 adult United States patients with erythema migrans [43]. This study demonstrated that carriage of the DRB1*0101 allele was associated with infection due to the RST3 genotype of B. burgdorferi (P = 0.01). The carriage rate of this allele was over five times greater in patients infected with RST3 strains of B. burgdorferi compared with those infected with RST1 strains. Although these results need to be confirmed, they do provide further evidence that host factors play an integral role in Lyme borreliosis,

G.P. Wormser / Dissemination and Persistence are Pathogenic Events

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particularly with regard to the development of infection caused by the least invasive strains of B. burgdorferi.

Table 4. Multivariate analysis of factors associated with spirochetemia in adult patients from the United States with erythema migrans [42]. Variable

Odds ratio

95% CI

P value

5.9

2.8-12.3

<0.001

2.9

1.3-6.6

0.009

2.5

1.1-5.7

0.03

> 55 yrs

2.7

1.1-6.5

0.03

Lymphopenia

5.4

2.3-12.9

<0.001

>1 EM Tick bite

1

First episode

2

1

Recollection of a tick bite at the site of the erythema migrans skin lesion. 2 First episode of Lyme borreliosis.

Prior in vitro studies on the pathogenesis of blood stream invasion in Lyme borreliosis have assessed endothelial cell or subendothelial matrix attachment, and penetration of endothelial cell monolayers by Lyme borrelia [44-51]. These studies have suggested that spirochetes may enter or exit from the blood stream by penetration through intercellular tight junctions or host cell cytoplasm. However, because the investigators did not compare disseminating and non-disseminating clinical isolates of Lyme borrelia, the results of these studies must be regarded as inconclusive. Repeating these experiments using strains of B. burgdorferi well-characterized with respect to their capacity for dissemination may be especially instructive in determining the validity of the endothelial cell monolayer system as a means for assessing the potential for spirochetes to enter and exit from the vascular system. Whatever the mechanism for blood stream invasion in Lyme borreliosis, it does not appear to be due simply to intravascular inoculation of the spirochete by the tick vector. A study in mice in which the site of tick attachment in skin was resected at various time intervals after tick detachment, clearly demonstrated that the spirochete remains localized at the initial skin site for at least 2 days after tick feeding is completed [52]. In summary, spirochetemia and persistence are common and important pathogenetic features of the major human spirochetal infections. Research is needed to identify host and spirochetal factors associated with entry into and exit from the blood stream, termination of spirochetemia (e.g., is this a function of immunologic factors or phenotypic changes in the spirochete) and host and spirochetal factors associated with persistence and with the transition from clinical latency to active disease.

Acknowledgements Lisa Giarratano, Justin Radolf, Ira Schwartz and Robert Nadelman provided invaluable assistance to the completion of this manuscript.

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9

References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21] [22] [23] [24] [25] [26] [27]

Mandell, Douglas and Bennett’s Principles and Practice of Infectious Diseases. Sixth Edition. Edited by: Mandell GL, Bennett JE, Dolin R. Elsevier, Churchill, Livingstone; Philadelphia, PA 2005. Peeling RW, Hook EW III. Syphilis control – a continuing challenge. N Engl J Med 2004;351:122-124. Centers for Disease Control and Prevention. Lyme Disease-United States, 2001-2002. MMWR 2004;53:365-9. O’Connell S, Granstrom M, Gray JS, Stanek G. Epidemiology of European Lyme borreliosis. Zentralbl Bakteriol 1998;287:229-240. Tramont EC. Treponema pallidum (syphilis). In: Mandell, Douglas and Bennett’s Principles and Practice of Infectious Diseases. Sixth Edition. Edited by: Mandell GL, Bennett JE, Dolin R. Elsevier, Churchill, Livingstone; Philadelphia, PA 2005; 2768-2785. Hook EW II. Endemic treponematoses. In: Mandell, Douglas and Bennett’s Principles and Practice of Infectious Diseases. Sixth Edition. Edited by: Mandell GL, Bennett JE, Dolin R. Elsevier, Churchill, Livingstone; Philadelphia, PA 2005; 2785-2788. Stanek G, Strle F. Lyme borreliosis. Lancet 2003;362:1639-1647. Washburn RG. Spirillum minus (rat bite fever). In: Mandell, Douglas and Bennett’s Principles and Practice of Infectious Diseases. Sixth Edition. Edited by: Mandell GL, Bennett JE, Dolin R. Elsevier, Churchill, Livingstone; Philadelphia, PA 2005; 2810. Levett, PN. Leptospirosis. In: Mandell, Douglas and Bennett’s Principles and Practice of Infectious Diseases. Sixth Edition. Edited by: Mandell GL, Bennett JE, Dolin R. Elsevier, Churchill, Livingstone; Philadelphia, PA 2005; 2789-2795. Rhee KY, Johnson WD Jr. Borrelia species (relapsing fever). In: Mandell, Douglas and Bennett’s Principles and Practice of Infectious Diseases. Sixth Edition. Edited by: Mandell GL, Bennett JE, Dolin R. Elsevier, Churchill, Livingstone; Philadelphia, PA 2005; 2795-2798. Stokes JH, Beerman H, Ingrham NR. Modern Clinical Syphilology. Saunders, Philadelphia 1944. Radolf JD, Hazlett KRO, Lukehart SA. Pathogenesis of syphilis. In: Molecular Biology of Treponemes and Pathogenesis of Treponemal Infections. Edited by: Radolf JD, Lukehart SA, Horizon Scientific Press, Norfolk, UK (In press). Shapiro ED, Dattwyler R, Nadelman RB, Wormser GP. Response to meta-analysis of Lyme borreliosis symptoms. Int J Epidemiol 2005;34:1437-1439. Barthold SW. Lyme borreliosis. In: Persistent Bacterial Infections. Edited by: Nataro JP, Blaser MJ, Cunningham-Rundles S. Washington, D.C.: ASM Press, 2000: 281-304. Liang FT, Nelson FK, Fikrig E. Molecular adaptation of Borrelia burgdorferi in the murine host. J Exp Med 2002;196:275-80. Diterich I, Rauter C, Kirschning CJ, Hartung T. Borrelia burgdorferi-induced tolerance as a model of persistence via immunosuppression. Infect Immun 2003;71:3979-3987. Liang FT, Brown EL, Wang T, Iozzo RV, Fikrig E. Protective niche for Borrelia burgdorferi to evade humoral immunity. Am J Pathol 2004;165:977-985. Rhen M, Eriksson S, Clements M, Bergstrom S, Normark SJ. The basis of persistent bacterial infections. Trends Microbiol 2003;11:80-86. Barbour AG. Antigenic variation of a relapsing fever borrelia species. Annu Rev Microbiol 1990;44:155-171. Levett PN. Leptospirosis. Clin Microbiol Rev 2001;14:296-326. Benach JL, Bosler EM, Hanrahan JP, et al. Spirochetes isolated from the blood of two patients with Lyme disease. N Engl J Med 1983;308:740-742. Steere AC, Grodzicki RL, Craft JE, Shrestha M, Kornblatt AN, Malawista SE. Recovery of Lyme disease spirochetes from patients. Yale J Biol Med 1984;57:557-560. Berger BW, Johnson RC, Kodner C, Coleman L. Cultivation of Borrelia burgdorferi from the blood of two patients with erythema migrans lesions lacking extracutaneous signs and symptoms of Lyme disease. J Am Acad Dermatol 1994;30:48-51. Goodman JL, Bradley LF, Ross AE, et al. Blood stream invasion in early Lyme disease: results from a prospective, controlled, blinded study using polymerase chain reaction. Am J Med 1995;99:6-12. Maraspin V, Ruzic-Sabljic E, Cimperman J, et al. Isolation of Borrelia burgdorferi sensu lato from blood of patients with erythema migrans. Infection 2001;29:65-70. Arnez M, Ruzic-Sabljic E, Ahcan J, Radsel-Medvescek A, Pleterski-Rigler D, Strle F. Isolation of Borrelia burgdorferi sensu lato from blood of children with solitary erythema migrans. Pediatr Infect Dis J 2001;20:251-255. Oksi J, Marttila H, Soini H, Aho H, Uksila J, Viljanen MK. Early dissemination of Borrelia burgdorferi without generalized symptoms in patients with erythema migrans. APMIS 2001;109:581-588.

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[28] Straubinger RK. PCR-Based quantification of Borrelia burgdorferi organisms in canine tissues over a 500-day postinfection period. J Clin Microbiol 2000;38:2191-9. [29] Hansen K, Lebech AM. The clinical and epidemiological profile of Lyme neuroborreliosis in Denmark 1985-1990. A prospective study of 187 patients with Borrelia burgdorferi specific intrathecal antibody production. Brain 1992;115 (Pt 2):399-423. [30] Wormser GP, Nowakowski J, Nadelman RB, Bittker S, Cooper D, Pavia C. Improving the yield of blood cultures for patients with early Lyme disease. J Clin Microbiol 1998; 36:296-298. [31] Wormser GP, Bittker S, Cooper D, Nowakowski J, Nadelman RB, Pavia C. Comparison of yields of blood cultures using serum or plasma from patients with early Lyme disease. J Clin Microbiol 2000;38:1648-50. [32] Wormser GP, Bittker S, Cooper D, Nowakowski J, Nadelman RB, Pavia C. Yield of large volume blood cultures in patients with early Lyme disease. J Infect Dis 2001;184:1070-2. [33] Coulter P, Lema C, Flayhart D, et al. Two-year evaluation of Borrelia burgdorferi culture and supplemental tests for definitive diagnosis of Lyme disease. J Clin Microbiol 2005;43:5080-4. [34] Perl B, Gottehrer NP, Raveh D, Schlesinger Y, Rudensky B, Yinnon AM. Cost-effectiveness of blood cultures for adult patients with cellulitis. Clin Infect Dis 1999;29:1483-8. [35] Wormser GP, Liveris D, Nowakowski J, et al. Association of specific subtypes of Borrelia burgdorferi with hematogenous dissemination in early Lyme disease. J Infect Dis 1999;180:720-725. [36] Wang G, Ojaimi C, Iyer R, et al. Impact of genotypic variation of Borrelia burgdorferi sensu stricto on kinetics of dissemination and severity of disease in C3H/HeJ mice. Infect Immun 2001;69:4303-4312. [37] Wang G, Ojaimi C, Wu H, et al. Disease severity in a murine model of Lyme borreliosis is associated with the genotype of the infecting Borrelia burgdorferi sensu stricto strain. J Infect Dis 2002;186:782791. [38] Seinost G, Dykhuizen DE, Dattwyler RJ, et al. Four clones of Borrelia burgdorferi sensu stricto cause invasive infection in humans. Infect Immun 1999;67:3518-24. [39] Liveris D, Wang G, Girao G, et al. Quantitative detection of Borrelia burgdorferi in 2 millimeter skin samples of erythema migrans lesions: correlation of results with clinical and laboratory findings. J Clin Microbiol 2002;40:1249-53. [40] Lagal V, Postec D, Ruzic-Sabljic E, Baranton G. Genetic diversity among Borrelia strains determined by single-strand conformation polymorphism analysis of the OspC gene and its association with invasiveness. J Clin Microbiol 2003;41:5059-65. [41] Liveris D, Gazumyan A, Schwartz I. Molecular typing of Borrelia burgdorferi sensu lato by PCRrestriction fragment length polymorphism analysis. J Clin Microbiol 1996;34:1306-9. [42] Wormser GP, McKenna D, Carlin J, et al. Brief communication: hematogenous dissemination in early Lyme disease. Ann Intern Med 2005;142:751-755. [43] Wormser GP, Kaslow R, Tang J, et al. Association between human leukocyte antigen class II alleles and genotype of Borrelia burgdorferi in patients with early Lyme disease. J Infect Dis 2005;192:20206. [44] Leong JM, Wang H, Magoun L, et al. Different classes of proteoglycans contribute to the attachment of Borrelia burgdorferi to cultured endothelial and brain cells. Infect Immun 1998;66:994-999. [45] Coleman JL, Sellati TJ, Testa JE, Kew RR, Furie MB, Benach JL. Borrelia burgdorferi binds plasminogen, resulting in enhanced penetration of endothelial monolayers. Infect Immun 1995;63:247884. [46] Sadziene A, Barbour AG, Rosa PA, Thomas DD. An OspB mutant of Borrelia burgdorferi has reduced invasiveness in vitro and reduced infectivity in vivo. Infect Immun 1993;61:3590-3596. [47] Ma Y, Sturrock A, Weis JJ. Intracellular localization of Borrelia burgdorferi within human endothelial cells. Infect Immun 1991;59:671-678. [48] Sadziene A, Thomas DD, Bundoc VG, Holt SC, Barbour AG. A flagella-less mutant of Borrelia burgdorferi structural, molecular, and in vitro functional characterization. J Clin Invest 1991;88:82-92. [49] Comstock LE, Thomas DD. Characterization of Borrelia burgdorferi invasion of cultured endothelial cells. Microb Pathog 1991;10:137-148. [50] Szczepanski A, Furie MB, Benach JL, Lane BP, Fleit HB. Interaction between Borrelia burgdorferi and endothelium in vitro. J Clin Invest 1990;85:1637-47. [51] Comstock LE, Thomas DD. Penetration of endothelial cell monolayers by Borrelia burgdorferi. Infect Immun 1989;57:1626-8. [52] Shih C-M, Pollack RJ, Telford SR. III, Spielman A. Delayed dissemination of Lyme disease spirochetes from the site of deposition in the skin of mice. J Infect Dis 1992;166:827-31.

Part 1 Molecular Genetics of Spirochetes

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

13

Transposon Mutagenesis of Infectious Borrelia burgdorferi B31: A Pilot Study Douglas J. BOTKIN a,b,c, April ABBOTT a,c, Jerrilyn K. HOWELL a, Mary MOSHER a, Philip E. STEWART d, Patricia A. ROSA d, Hiroki KAWABATA e, Haruo WATANABE e, and Steven J. NORRIS a,b,c,1 a Department of Pathology and Laboratory Medicine,University of Texas Medical School at Houston, Houston, TX, U.S.A. b Department of Microbiology and Molecular Genetics, University of Texas Medical School at Houston, Houston, TX, U.S.A. c Graduate School of Biomedical Sciences, University of Texas Health Science Center at Houston, Houston, TX, U.S.A. d Laboratory of Human Bacterial Pathogenesis, Rocky Mountain Laboratories, National Institutes of Allergy and Infectious Diseases, N.I.H., Hamilton, MT U.S.A. e National Institute of Infectious Diseases, Tokyo, Japan Abstract. Transposon mutagenesis is a powerful technique for generating random mutations. Previous studies had shown that the mariner transposon Himar1 could be utilized to introduce mutations in high-passage, noninfectious strains of Lyme disease Borrelia; however, transposon mutagenesis has not been reported in low-passage, infectious strains. In this pilot study, Himar1 transposon mutants were generated by transforming the infectious, highly transformable B. burgdorferi B31 clone 5A18 NP1 with the transposon vector pMarGent. The data provide ‘proof in concept’ that random transposon mutagenesis can be utilized for the global analysis of genes involved in the pathogenesis of Lyme disease Borrelia. Keywords. Global mutagenesis, infectious Borrelia, mutagenesis, transposition

Introduction Lyme disease Borrelia possess an unusually complex genome consisting of a linear chromosome and approximately 9 linear plasmids and 11 circular plasmids [1, 2]. The relative consistency of plasmid content in the low-passage strains examined to date 1 Corresponding Author: Steven J. Norris, Ph.D., Department of Pathology and Laboratory Medicine, University of Texas Medical School at Houston, P.O. Box 20708, Houston, TX 77225-0708, U.S.A.; Email: [email protected].

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D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

indicates that these genetic elements are a stable and vital part of the genome of the organism [3]. The plasmids themselves contain large numbers of paralogous genes of unknown function; with a few notable exceptions, the ‘housekeeping’ genes involved in central metabolism and DNA, RNA, and protein synthesis are located on the chromosome. Plasmid loss occurs spontaneously during in vitro culture, and absence of the linear plasmids lp25 and lp28-1 results in reduced infectivity [4í9]. Coupled with this unusual genetic structure is a biphasic life style, in which B. burgdorferi and other members of this group alternate between Ixodes ticks and

Figure 1. Characterization of bacterial pathogens by invasiveness versus toxigenesis. B. burgdorferi, like T. pallidum and M. tuberculosis, is highly invasive and causes persistent infection, but is not known to produce any toxins.

mammalian or avian hosts. Within permissive mammalian hosts, including mice and humans, the spirochete readily disseminates and can infect multiple organ systems. Although the mechanisms of pathogenesis are not well understood, the primary mode of tissue damage appears to be the induction of a host inflammatory response. B. burgdorferi is able to infect mice for at least two years; although the duration of infection in humans is not known, symptoms such as acroderma chronicum atrophicans can occur several years after the initial infection. Thus Lyme disease Borrelia fall into a class of invasive, persistent pathogens, including Treponema pallidum and Mycobacterium tuberculosis, that are able to infect and cause manifestations in human hosts for years to decades (Figure 1). These

D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

15

organisms do not appear to produce toxins, and relatively little is known regarding their mechanisms of pathogenesis.

1. Transformation of B. burgdorferi Genetic studies of Lyme disease Borrelia pathogenesis have been hampered by the low transformation rate of infectious, low-passage B. burgdorferi strains relative to highpassage, noninfectious strains, although progress has been made in recent years [10].

Figure 2. Putative Type IV restriction-modification enzymes of B. burgdorferi. The gene identifiers, plasmid locations, and predicted protein lengths are indicated, and the Escherichia coli type IV enzyme Eco57I is shown for comparison. The endonuclease catalytic-Mg2+ binding domain is shown in black, and the CM sites correspond to methyltransferase signature sequences typical of DNA modification enzymes. The restriction and modification functions are combined in type IV enzymes.

Lawrenz et al. [11] found that transformation of low-passage, infectious B. burgdorferi B31 with the shuttle vector pBSV2 [12] was inefficient; the only clones that were transformed had lost the 25 kb linear plasmid lp25 that is required for pathogenesis [4í8, 13]. A survey of low-passage clones with varied plasmid content indicated that the transformation frequency with pBSV2 was low when lp25 and lp56 were present, intermediate when either one or the other was missing, or high when both were missing [11]. Examination of the open reading frames (ORFs) of these two plasmids revealed that each contained a large ORF with both restriction endonuclease and DNA methyltransferase motifs (Figure 2). These ORFs (BBE02 and BBQ67) are similar to Type IV restriction endonucleases such as Eco57I that possess both activities in one protein; as with restriction endonucleases in general, the homology is limited to the catalytic-Mg++ binding endonuclease domain and the methyltransferase signature sequences [14]. The plasmid lp28-3 also encodes the

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D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

putative restriction-modification protein BBH09; it is 92% similar to BBE02 despite having a markedly different pI (8.07 vs. 6.99), which may reflect differences in recognition sequences or other activities. The possibility that BBH09 may be involved in reducing transformation frequency has not been examined, because low-passage clones lacking lp283 were not included in this study [14]. Our working model is that BBE02 and BBQ67 interfere with transformation by cleaving incoming DNA before it can be methylated at the corresponding restriction sites. Kawabata et al. [15] disrupted BBE02 through the insertion of a kanamycin resistance cassette with a B. burgdorferi flgB promoter (flgBp). They showed that this mutation increased the transformation frequency with the shuttle vector pBSV2C03::gnt kan in 5A4 NP1, a B31 clone that contains all of the B. burgdorferi plasmids. Furthermore, disruption of BBE02 in the clone 5A18 NP1 resulted in a higher transformation frequency, correlating with the lack of lp56 (and hence BBQ67) in this clone. Both 5A4 NP1 and 5A18 NP1 retained full infectivity in mice as measured by median infectious dose (ID50) following needle inoculation [15]. These BBE02 disruptants thus may ease the study of B. burgdorferi pathogenesis by increasing the efficiency of site-directed or random transposon mutagenesis in infectious strains, which has been a stumbling block to date.

2. Transposon Mutagenesis Transposon mutagenesis provides a means for introducing random mutations, producing a library that can be examined for phenotypic changes. Two recent studies have demonstrated the feasibility of using the Himar1 version of the mariner transposon in B. burgdorferi. mariner-based transposons have the advantage of a short target sequence, requiring only a TA dinucleotide sequence for insertion. Stewart et al. [16] utilized the suicide vector pMarGent (Figure 3) to deliver a Himar1 transposon that contains a flgBp::aacC1 cassette, thus conferring gentamicin resistance. An important feature of pMarGent is the presence of the hyperactive C9 transposase expressed from the Borrelia promoter flgBp, thus increasing expression of the transposase following transformation. Using the non-infectious clones A3-89 and B31-AchbC72, a transformation frequency of ~3.0 u 10-5 to 5.5 u 10-5 was obtained, yielding between 5,000 and 40,000 transposon mutants per transformation. Thus each transformation could yield a library with mutations spaced by an average of ~300 bp, assuming a random distribution of mutations. Both of these B. burgdorferi B31 clones lack lp25 and lp56, which may contribute to the high transposition frequency observed. Mutants with slow growth rates were obtained, including those with mutations in BB0414 (encoding cheR-2, a putative chemotaxis methylase), BB0257 (encoding an FtsK/SpoIIIE homolog), and BB0323 (encoding a protein with a LysM motif, which may be involved in peptidoglycan biosynthesis). In contrast, no mutants were obtained in the infectious B. burgdorferi strains B31-A3 and N40, most likely because these strains contain both lp25 and lp56. In another study, Morozova et al. [17] utilized the broad host range vector pED7, which replicates at low copy number in B. burgdorferi, to express the C9 Himar1

D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

17

Figure 3. The Himar1 transposon delivery vector pMarGent, as described by Stewart et al. [14]. flgBp::Himar1is the Himar1 C9 transposase gene fused with the flgB promoter from B. burgdorferi. IR corresponds to the Himar1 inverted repeat sequence that demarcates the transposable element. ColE1 is a plasmid origin of replication that permits propagation of the vector in E. coli, and flgBp::aacC1 confers gentamicin resistance in B. burgdorferi and E. coli. Col and Flg are the locations of primers used for sequencing the insertion site in plasmids rescued from B. burgdorferi transposon mutants. Reprinted with permission from [16].

transposase under control of the Borrelia flaB promoter (flaBp). Non-infectious, highpassage B. burgdorferi B31 containing pED7 were transformed with three different suicide plasmids containing Himar1 inverted repeat sequences, a kanamycin resistance cassette, and a colE1 origin of replication. Transposition frequencies of ~100 clones per Pg DNA were obtained, yielding hundreds of colonies per transposition. Mutants were obtained in the 23S rRNA gene rrlA and in the ftsJ/rrmJ homolog BB0313. Surprisingly, inactivation of one of the two 23S rRNA genes had no significant effect on the growth of B. burgdorferi, which may reflect the relatively low requirement of this slow-growing organism for ribosome production. The ftsJ/rrmJ mutant had elongated organisms, decreased motility, and reduced growth rates. In this case, it is unclear whether the phenotype was due to the interruption of ftsJ/rrmJ or polar effects on downstream genes implicated in cell division [17]. Transposition of infectious strains of B. burgdorferi was not reported in this study.

3. Transposition in Infectious B. burgdorferi Creation of random transposon mutant libraries in infectious B. burgdorferi strains would permit a global examination of the genes important in the Lyme disease infection cycle. In a pilot study, we determined whether Himar1 transposon mutants could be obtained by transforming the BBE02 mutant clones 5A4 NP1 and 5A18 NP1 with pMarGent. Transformation with pMarGent was carried out as described by Stewart et al. [16] in two independent experiments. Following incubation for 24 hours in BSK II medium [18] without antibiotics, the transformed cultures were inoculated into plates containing BSK II

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D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

medium with 0.65% low melting point agarose (SeaKem LE, Cambrex), 40 Pg/ml gentamicin, 200 Pg/ml kanamycin, and an antibiotic mixture to suppress growth of other microorganisms (Sigma-Aldrich, Product No. A1956). Colonies were visible after 9í14 days of incubation, and well-isolated colonies were selected for additional analysis. All clones isolated contained the gentamicin resistant cassette as assessed by PCR [16]. The insertion sites of the Himar1 transposon were determined by the ligation of HindIII fragments from genomic B. burgdorferi DNA, rescue of gentamicin resistance-conferring plasmids in Escherichia. coli, and sequencing of the Borrelia DNA flanking the transposon [16]. Screening of clones for infectivity was accomplished by needle inoculation of individual C3H/HeN mice with 104 organisms of each clone, and culturing multiple tissue sites 2 weeks post-infection, as described previously [19]. The infectivity phenotype of tenuous potentially attenuated clones was confirmed by inoculating groups of 2 mice with 106, 104, or 102 organisms and culturing tissue sites 2 weeks thereafter. No gentamicin-resistant colonies were obtained when 5A4 NP1 was transformed with either pMarGent or the shuttle vector pBSV2-G, which also carries the gentamicin resistance cassette flgBp::aacC1. This result indicates that transposition with pMarGent (or pBSV2-G) is still an inefficient process if BBE02 is disrupted but lp56 (and hence BBQ67) is still present. In contrast, transformation of 5A18 NP1 with pMarGent resulted in 4 gentamicin-resistant colonies in the first transformation, and 29 such colonies in the second experiment. Presence of the pMarGent transposon in each of these clones was confirmed by PCR of the aacC1 cassette region and/or sequencing of the flanking DNA (data not shown). The insertion sites of 30 of the 33 transposon mutants have been determined (Table 1). The groups of circular plasmids and linear plasmids each contained 12/30 (40%) of the mutations, whereas only 6 (20%) were localized in the chromosome. A relatively low number of mutations in the B. burgdorferi chromosome was also observed by Stewart et al. [16]. This finding most likely reflects a high proportion of essential genes in the chromosome, precluding survival of clones in which the mutation has occurred in a required gene. Otherwise, the distribution of mutations appeared to be random. All insertions were at TA sequences, consistent with the specificity of Himar1 tranposons. To screen for infectivity phenotype, each of the 33 mutants was inoculated (104 organisms) into one mouse per clone (Table 1). Eighteen of the clones were consistently recovered from all four sites assayed and were considered fully infectious, although some could have infectivity defects not detected by this relatively insensitive assay. Eleven of the clones were culture negative at one or more sites and thus were potentially attenuated. Of these, 6 clones were missing lp28-1, which is known to be required for full infectivity [4,68]. Four clones were not recovered from any of the sites and were preliminarily considered non-infectious. Clone A3 was found to lack lp25 (which is required for infectivity [4í9]), so this clone was not pursued further. Based on this screening, four of the lp28-1+ attenuated clones and three apparently non-infectious clones were selected for further analysis of mouse infectivity. (Studies involving clone B29 had not been completed at the time of this writing.) Among these clones, only the guaB mutant A2 exhibited reduced growth rates in BSK II medium (data not shown). Plasmid content was determined by PCR [4] to assess the potential effects of plasmid loss on infectivity. For each clone, groups of two mice were inoculated with 106,

Table 1. Insertion sites of pMarGent mutants (grouped by infectivity phenotype). Clone

+

A4

Replicon/Position2

Insertion site3

Predicted function or description

Chrom./108542

BB0110 (108307-109671)

Hypothetical protein

+

B1

cp32-6/24624

BBM36 (24619-25044)

Borrelia plasmid protein B (bppB) (putative)

+

B2

lp36/17582

BBK25 (17565-16966) promoter

Transposase-like protein (putative), authentic frameshift

+

B3

Chrom./534145

Intergenic - BB0524 (534301-535115) and BB0523(533980-534072)

BB0524 possible Myo-inositol-1 (or 4)- mono-phosphatase (putative); BB0523 is a hypothetical protein

+

B6

cp32-3/27222

BBS41 (26708-27298)

Outer surface protein G (erpG)

+

B7

lp28-3/4210

BBH09 (7728-3892)

Hypothetical protein

+

B8

lp28-1/4515

BBF10 (4488-4985)

Hypothetical protein

+

B12

cp32-8/25599

BBL38 (25951-25175)

Borrelia plasmid protein C (bppC) (putative)

+

B13

lp38/33790

Intergenic - BBJ45 (32498-33220) and BBJ46 (34668-34372)

BBJ45 and BBJ46 are hypothetical proteins

+

B14

cp32-4/24618

BBR37 (24210-24635)

Borrelia plasmid protein B (bppB) (putative)

+

B16

cp26/23390

BBB28 (23255-24499)

Hypothetical protein

+

B21

lp21/7829

Intergenic - BBU05 (2868-3656) and BBU06 (14633-15235)

Within central 63 bp repeat region of lp21

+

B22

lp36/23615

BBK37 (23953-22910)

Homolog of immunogenic protein P37

+

B23

cp32-4/7055

BBR11 (6711-7823)

Hypothetical protein

+

B24

cp32-4/17887

BBR29 (18664-17573)

Conserved hypothetical protein

+

B25

lp28-1/5088

Intergenic - BBF10 (4488-4985) and BBF11 (5435-5542)

BBF10 and BBF11 are hypothetical proteins

+

B26

cp26/25676

BBB29 (24825-26453)

Phosphotransferase system enzyme, maltose and glucosespecific IIABC component (malX)

+

B28

lp17/9413

Intergenic - BBD14 (8269-9381) and BBD15 (10015-9593)

BBD14 is a conserved hypothetical protein and BBD15 is a hypothetical protein

D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

Infectivity 1

19

20

Table 1, cont’d. +/- (lp28-1-)

B4

Chrom./660583

BB0629 (661524-659644)

Phosphotransferase system enzyme II (fruA-2)

B11

lp38/25996

BBJ34 (26407-25337)

Hypothetical protein

B17

lp28-2/20360

BBG24 (22310-19620)

Hypothetical protein

+/-

B18

lp25/21344

Intergenic - BBE29 (19697-20886) and BBE30 (21558-21704)

BBE29 - putative adenine specific methyltransferase, authentic frameshift; BBE30 - hypothetical protein

+/-

B19

lp21/6922

Intergenic - BBU05 (2868-3656) and BBU06 (14633-15235)

Within central 63 bp repeat region of lp21

+/-

B27

cp26/12701

BBB16 (12014-13606)

oligopeptide ABC transporter, periplasmic oligopeptidebinding protein (oppAIV)

BB0743 (785057-786754)

Hypothetical protein

+/- (lp28-1+)

B5

Chrom./786112

+/-

B9

Not determined

+/-

B10

Chrom./841814

BB0797 (839244-841832)

DNA mismatch repair protein (mutS)

+/-

B15

cp9/5623

BBC08 (5534-5980)

Conserved hypothetical protein

+/-

B29

cp32-7/22694

BBO35 (22755-22627)

Hypothetical protein

-

A1

Not determined BBB17 (15107-13893)

IMP dehydrogenase (guaB)

BB0221 (227836-226610)

flagellar motor switch protein (fliG-1)

1

-

A2

cp26/14424

-

A3

Not determined

-

B20

Chrom./226987

+, culture positive at all sites; +/-, culture positive at some sites; -, culture negative at all sites.

2

Chrom. = Chromosome.

3

Position of gene given in parentheses. Reverse coordinates indicate that the gene is encoded on the negative strand.

D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

+/+/-

D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

21

Table 2. Multiple dose inoculations with attenuated and noninfectious pMarGent mutants. 1 Mutant

Culture Positive Sites/Total

Mutant

Culture Positive Sites/Total -

A2 (guaB; cp9 , cp32-3-)

A1 (insertion site ND; no plasmids lost) Inoculum

Ear

Bladder

Heart

Joint

Inoculum

Ear

Bladder

Heart

Joint

102

0/2

0/2

0/2

0/2

All

0/8

102

0/2

0/2

0/2

0/2

0/8

104

0/3

0/3

0/3

0/3

0/12

104

0/3

0/3

0/3

0/3

0/12

106

0/2

0/2

0/2

0/2

0/8

106

0/2

0/2

0/2

0/2

0/8

B5 (hypothetical protein; cp9 -)

All

B15 (hypothetical protein; lp38-)

Inoculum

Ear

Bladder

Heart

Joint

Inoculum

Ear

Bladder

Heart

Joint

102

0/2

½

1/2

2/2

All

3/8

102

0/2

0/2

0/2

0/2

0/8

104

1/3

3/3

3/3

3/3

10/12

104

0/3

1/3

1/3

2/3

4/12

106

0/2

2/2

2/2

2/2

6/8

106

0/2

0/2

0/2

0/2

0/8

B9 (insertion site ND; cp9-)

All

B20 (fliG-1; no plasmids lost)

Inoculum

Ear

Bladder

Heart

Joint

Inoculum

Ear

Bladder

Heart

Joint

102

0/2

0/2

0/2

0/2

All

0/8

102

0/2

0/2

0/2

0/2

0/8

104

0/3

2/3

0/3

3/3

5/12

104

0/3

0/3

0/3

0/3

0/12

106

2/2

2/2

0/2

2/2

6/8

106

0/2

0/2

0/2

0/2

0/8

B10 (mutS; cp9-, cp32-6-)

All

5A18 NP1 (parental strain)

Inoculum

Ear

Bladder

Heart

Joint

All

Inoculum

Ear

Bladder

Heart

Joint

102

0/2

0/2

0/2

0/2

0/8

102

0/2

0/2

0/2

0/2

All

0/8

104

1/3

2/3

1/3

2/3

6/12

104

3/3

3/3

3/3

3/3

12/12

106 1/2 2/2 1/2 2/2 6/8 106 2/2 2/2 2/2 2/2 8/8 Two or three mice were inoculated with each clone and dose. The mice were sacrificed and tissue sites cultured 2 weeks post-inoculation. The site of transposon insertion and plasmids missing (in addition to lp56 and lp28-4 which are absent in the parental strain 5A18 NP1) are indicated for each clone. ND = not determined.

1

104, and 102 bacteria. Table 2 shows the combined results of these experiments and the initial single mouse inoculations. For unknown reasons, the parental clone 5A18 NP1 was culture negative at an inoculum of 102 B. burgdorferi, contrary to the ID50 of 83 organisms obtained in an earlier study [15]. It was confirmed that clones A1, A2, and B20 were noninfectious in the mouse model. The guaB gene, encoding an inosine monophosphate dehydrogenase involved in the interconversion between inosine and guanine, is disrupted in mutant A2. Previous attempts to disrupt guaB in a non-infectious B. burgdorferi strain had yielded a single clone in which one wild-type and one disrupted copy of guaB were present in a cp26 dimer [20]. Colonies with a monomeric form of cp26 with the guaB mutation were isolated upon replating of this clone, indicating that B. burgdorferi with only a disrupted copy of guaB were capable of survival and growth in vitro [20]. The pMarGent

22

D.J. Botkin et al. / Transposon Mutagenesis of Infectious Borrelia burgdorferi B31

mutant A2 is missing plasmids cp9 and cp32-3 (Table 2), which were previously shown to not be required for mouse infection [4, 21]. The mutation in clone B20 is located in fliG-1, one of two fliG switch protein homologs thought to be involved in the reversal of flagellar rotation. B20 is still motile in BSK II medium, yet is noninfectious. Thus this mutation may provide information regarding subtle aspects of motility and its relationship to pathogenesis. The site of the transposon mutation for clone A1 has not been determined as yet, despite intensive efforts. Neither B20 nor A1 were lacking plasmids, other than the lp56 and lp28-4 plasmids absent in the parental 5A18 NP1 strain. Thus the reduced infectivity of these clones is likely to be directly related to the transposon mutation. Clones B9 and B15 were attenuated, with recovery from 11/28 and 4/28 sites, respectively. Interestingly, all of the heart cultures were negative for clone B9, suggesting that tissue tropism is affected in this clone. The mutation site for clone B9 has not been determined. The mutation in clone B15 is located in cp9, which is not required for mouse infection [4, 21]. However, it is missing lp38; no clone lacking this plasmid has been characterized previously, and it is possible that loss of lp38 results in reduced infectivity. Mutant B10, which harbors a transposition event in the gene encoding the DNA mismatch repair protein MutS, was somewhat reduced in culture positivity (12/28 sites positive). The product of this gene may play a role in DNA repair in response to oxidative stress or other assaults in the mammalian host. Further analysis is needed to determine if this clone is indeed attenuated and if its phenotype is related to the loss of cp32-6 (Table 2). Clone B5 was culture positive at some tissues from mice inoculated with 102 Borrelia and from most sites from mice inoculated with higher doses and thus may represent a “false positive” result from the initial screen for attenuation. The B5 mutation is located within the chromosomal hypothetical protein gene BB0743. This pilot study provides ‘proof in concept’ that transposon mutagenesis in infectious B. burgdorferi is possible and will yield valuable information regarding the genes important in mammalian infection. Of 33 mutant clones examined, 8 were found to have reduced infectivity that was not due to loss of lp28-1 or lp25. Gene complementation and transmission through the tick-mouse cycle will be used to determine the roles of the mutated genes in the natural infection process. Expansion of this approach to a larger transposon mutation library, including the planned use of signature tagged mutagenesis, will provide a more global view of the genes required for the complex, biphasic lifestyle of Lyme disease Borrelia.

Acknowledgments We thank Kit Tilly for invaluable advice regarding this project. This work was supported by NIH grant R01 AI59048.

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23

References [1]

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Fraser, C. M., S. Casjens, W. M. Huang, G. G. Sutton, R. Clayton, R. Lathigra, O. White, K. A. Ketchum, R. Dodson, E. K. Hickey, M. Gwinn, B. Dougherty, J. F. Tomb, R. D. Fleischmann, D. Richardson, J. Peterson, A. R. Kerlavage, J. Quackenbush, S. Salzberg, M. Hanson, R. van Vugt, N. Palmer, M. D. Adams, J. Gocayne, J. Weidman, T. Utterback, L. Watthey, L. McDonald, P. Artiach, C. Bowman, S. Garland, C. Fujii, M. D. Cotton, K. Horst, K. Roberts, B. Hatch, H. O. Smith, J. C. Venter and et al. 1997. Genomic sequence of a Lyme disease spirochaete, Borrelia burgdorferi. Nature 390: 580í586. Casjens, S., N. Palmer, R. van Vugt, W. M. Huang, B. Stevenson, P. Rosa, R. Lathigra, G. Sutton, J. Peterson, R. J. Dodson, D. Haft, E. Hickey, M. Gwinn, O. White and C. M. Fraser. 2000. A bacterial genome in flux: the twelve linear and nine circular extrachromosomal DNAs in an infectious isolate of the Lyme disease spirochete Borrelia burgdorferi. Mol. Microbiol. 35: 490í516. Palmer, N., C. Fraser and S. Casjens. 2000. Distribution of twelve linear extrachromosomal DNAs in natural isolates of Lyme disease spirochetes. J. Bacteriol. 182: 2476í2480. Purser, J. E. and S. J. Norris. 2000. Correlation between plasmid content and infectivity in Borrelia burgdorferi. Proc. Natl. Acad. Sci. USA 97: 13865í13870. Purser, J. E., M. B. Lawrenz, M. J. Caimano, J. K. Howell, J. D. Radolf and S. J. Norris. 2003. A plasmidencoded nicotinamidase (PncA) is essential for infectivity of Borrelia burgdorferi in a mammalian host. Mol Microbiol 48: 753í764. Labandeira-Rey, M., E. Baker and J. Skare. 2001. Decreased infectivity in Borrelia burgdorferi strain B31 is associated with loss of linear plasmid 25 or 28-1. Infect. Immun. 69: 446í455. Labandeira-Rey, M., J. Seshu and J. T. Skare. 2003. The absence of linear plasmid 25 or 28-1 of Borrelia burgdorferi dramatically alters the kinetics of experimental infection via distinct mechanisms. Infect Immun 71: 4608í4613. Grimm, D., C. H. Eggers, M. J. Caimano, K. Tilly, P. E. Stewart, A. F. Elias, J. D. Radolf and P. A. Rosa. 2004. Experimental assessment of the roles of linear plasmids lp25 and lp28-1 of Borrelia burgdorferi throughout the infectious cycle. Infect Immun 72: 5938-5946. Xu, Y., C. Kodner, L. Coleman and R. C. Johnson. 1996. Correlation of plasmids with infectivity of Borrelia burgdorferi sensu stricto type strain B31. Infect. Immun. 64: 3870-3876. Rosa, P. A., K. Tilly and P. E. Stewart. 2005. The burgeoning molecular genetics of the Lyme disease spirochaete. Nat Rev Microbiol 3: 129-143. Lawrenz, M. B., H. Kawabata, J. E. Purser and S. J. Norris. 2002. Decreased electroporation efficiency in Borrelia burgdorferi containing linear plasmids lp25 and lp56: impact on transformation of infectious Borrelia. Infect. Immun. 70: 4798-4804. Stewart, P. E., R. Thalken, J. L. Bono and P. Rosa. 2001. Isolation of a circular plasmid region sufficient for autonomous replication and transformation of infectious Borrelia burgdorferi. Mol. Microbiol. 39: 714-721. Revel, A. T., J. S. Blevins, C. Almazan, L. Neil, K. M. Kocan, J. de la Fuente, K. E. Hagman and M. V. Norgard. 2005. bptA (bbe16) is essential for the persistence of the Lyme disease spirochete, Borrelia burgdorferi, in its natural tick vector. Proc Natl Acad Sci U S A 102: 6972-6977. Lawrenz, M. B., H. Kawabata, J. E. Purser and S. J. Norris. 2002. Decreased electroporation efficiency in Borrelia burgdorferi containing linear plasmids lp25 and lp56: impact on transformation of infectious B. burgdorferi. Infect Immun 70: 4798-4804. Kawabata, H., S. J. Norris and H. Watanabe. 2004. BBE02 disruption mutants of Borrelia burgdorferi B31 have a highly transformable, infectious phenotype. Infect. Immun. 72: 7147-7154. Stewart, P. E., J. Hoff, E. Fischer, J. G. Krum and P. A. Rosa. 2004. Genome-wide transposon mutagenesis of Borrelia burgdorferi for identification of phenotypic mutants. Appl Environ Microbiol 70: 5973-5979. Morozova, O. V., L. P. Dubytska, L. B. Ivanova, C. X. Moreno, A. V. Bryksin, M. L. Sartakova, E. Y. Dobrikova, H. P. Godfrey and F. C. Cabello. 2005. Genetic and physiological characterization of 23S rRNA and ftsJ mutants of Borrelia burgdorferi isolated by mariner transposition. Gene 357: 63-72. Barbour, A. G. 1984. Isolation and cultivation of Lyme disease spirochetes. Yale J. Biol. Med. 57: 521-525. Norris, S. J., J. K. Howell, S. A. Garza, M. S. Ferdows and A. G. Barbour. 1995. High- and low-infectivity phenotypes of clonal populations of in vitro-cultured Borrelia burgdorferi. Infect. Immun. 63: 2206-2212. Tilly, K., L. Lubke and P. Rosa. 1998. Characterization of circular plasmid dimers in Borrelia burgdorferi. J. Bacteriol. 180: 5676-5681.

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[21] Elias, A. F., P. E. Stewart, D. Grimm, M. J. Caimano, C. H. Eggers, K. Tilly, J. L. Bono, D. R. Akins, J. D. Radolf, T. G. Schwan and P. Rosa. 2002. Clonal polymorphism of Borrelia burgdorferi strain B31 MI: implications for mutagenesis in an infectious strain background. Infect. Immun. 70: 2139-2150.

Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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The Isolation and Characterization of Isogenic Mutants in Infectious Borrelia burgdorferi J. SESHU a,b, Maria LABANDEIRA-REY a,c, M. Dolores ESTEVE-GASSENT a,b, Magnus HÖÖK c, and Jonathan T. SKARE a,1 a Department of Microbial and Molecular Pathogenesis, Texas A&M University Health Science Center, College of Medicine, College Station, TX, USA b Department of Biology and South Texas Center for Emerging Infectious Diseases, University of Texas at San Antonio, San Antonio, TX, USA c Center for Extracellular Matrix Biology, Texas A&M University HSC, Institute of Biosciences and Technology, Houston, TX, USA Abstract. Borrelia burgdorferi, the causative agent of Lyme disease, is the most prevalent arthropod-borne infectious agent in the United States and, along with infections attributed to other related Borrelia spp. worldwide, contributes to a significant amount of morbidity. Much of what is known regarding pathogenic mechanisms of B. burgdorferi has been limited to biochemical analyses due to the genetic intractability of this spirochetal pathogen. Recently, advances by a number of investigators have made it possible to isolate and genetically complement isogenic mutants in infectious B. burgdorferi as a first step in determining the role of specific genes in Lyme borreliosis. The approach described here involves a customized transposon that contains an antibiotic resistance marker expressed from a strong borrelial promoter, which is used to mutagenize a borrelial gene target in vitro. The resulting transposon mutagenized candidates are then mapped and sequenced. The desired constructs, containing the inactivated gene, are transformed into a low passage but non-infectious B. burgdorferi strain lacking the 25 kb linear plasmid (lp25) and putative transformants selected with the appropriate antibiotic. The lp25 deficient strain used here is more amenable to transformation due to the absence of an lp25-specific restriction/modification system. Using this genetic background, several borrelial genes purported to be involved in B. burgdorferi pathogenesis have been genetically inactivated. The loss of infectivity associated with the lp25 deficient background can be restored back to wild-type levels by transformation with the borrelial shuttle vector pBBE22, which encodes the lp25 bbe22 locus. The bbe22 (pncA) gene of this strain encodes a nicotinamidase enzyme required for de novo biosynthesis of NAD only during infection. Thus, transformation with pBBE22 serves two purposes, as it: (i) restores infectivity in the parent background, and (ii) asks whether the resulting mutant exhibits an infectivity deficient phenotype relative to the parent background using the well-established mouse model of infection. Subsequent genetic complementation with an intact version of the targeted gene in pBBE22 is 1 Corresponding Author: Jonathan T. Skare, Department of Microbial and Molecular Pathogenesis, Texas A&M University Health Science Center, 407 Reynolds Medical Bldg., College Station, TX 77843-1114, USA; Email: [email protected].

J. Seshu et al. / The Isolation and Characterization of Isogenic Mutants

26

then carried out to ensure that the mutant phenotype observed can be attributed to the inactivated gene. The strategy presented here could be applied to the genetic analysis of other borrelial genes to determine their significance in the pathogenesis of Lyme borreliosis. Keywords. Lyme disease, mutagenesis, genetics, transformation, selection

Introduction The Borrelia burgdorferi sensu lato complex defines the borrelial isolates that are associated with Lyme borreliosis. They are specifically, B. burgdorferi sensu stricto, B. afzelii, and B. garinii, and each has an uneven geographic distribution [1]. B. burgdorferi is the dominant cause of Lyme disease in the United States, whereas B. afzelii and B. garinii are common causes of borrelioses in Europe and Asia [2]. Furthermore, the late clinical manifestations of borreliosis caused by the three species differ. B. burgdorferi is primarily associated with arthritis; B. garinii often causes neuroborreliosis and B. afzelii, acrodermatitis [2], although it is important to emphasize that these manifestations are not absolute and there is significant overlap between them within the aforementioned geographic regions. Nevertheless, in the United States, B. burgdorferi infection, if not treated early, can lead to a debilitating arthritis that is refractory to antibiotic therapy [2í4]. Therefore, in endemic areas, Lyme disease contributes to a significant amount of morbidity. The B. burgdorferi genome was sequenced and published in 1997 and updated in 2000 [5, 6]. Despite the wealth of knowledge accrued from the annotation of the B. burgdorferi genome, we understand alarmingly little regarding the molecular pathogenesis of Lyme disease. This is due predominantly to the inability to isolate mutations in putative virulence determinants; thus, until recently, it was not possible to assess molecular Koch's postulates. As such, previous studies on putative borrelial virulence determinants were restricted either to biochemical analyses or the expression of selected genes within an infected host system. However, the development of genetic tools to obtain site-specific knock-out mutations in infectious B. burgdorferi has provided important insight into the roles of different gene products in the physiology and pathogenesis of this organism [7í21]. Despite these advances, the ability to generate and complement isogenic mutants in infectious B. burgdorferi remains cumbersome. The difficulty in obtaining isogenic mutants in B. burgdorferi has resulted in many different methods for isolating mutants in infectious isolates. In this report, an additional approach for isolating site directed mutants is described. The alternative method described here uses in vitro transposition of a customized transposon and initial screening in E. coli to isolate mutations in the gene of interest, followed by mapping and sequencing of the transposition events. The resulting inactivated gene is then used to transform a borrelial isolate that is more amenable to genetic analysis but noninfectious due to the absence of the 25 kilobase linear plasmid lp25 [22í24]. The resulting mutant, obtained by a double crossover recombination event, is transformed independently with (i) a shuttle vector containing the “infectivity restoration marker” (IRM; the bbe22 locus) that restores infectivity to a borrelial isolate that lacks lp25 [25] and (ii) IRM and the inactivated gene to restore infectivity and genetically complement the mutant simultaneously. In theory, this approach can be exploited to isolate mutations in any non-essential borrelial gene.

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27

1. Materials and Methods 1.1. Bacterial Strains and Shuttle Vectors ML23 is a clonal isolate of Borrelia burgdorferi strain B31 that lacks the 25 kilobase linear plasmid lp25 [19, 22, 23] and was the parental strain used for all borrelial transformations described here. All cloning in Escherichia coli was done using TOP10 cells (Invitrogen Corp., Carlsbad, California, USA). The shuttle vectors used were (i) pKFSS1 (kindly provided by Scott Samuels, [26]), which served as the source for the aadA gene and confers resistance to spectinomycin in E. coli and streptomycin in B. burgdorferi, and (ii) pBBE22 (generously provided by Steve Norris, [25]), which confers resistance to kanamycin and contains the bbe22 gene that restores infectivity to ML23 (and to other strains missing lp25 [25]) upon transformation. 1.2. PCR PCR was conducted as previously described [19, 23, 27]. 1.3. Plasmid Encoded Customized Transposon Constructs Plasmid pJS104 has been described previously [27]. Briefly, pJS104 contains a kanamycin resistance determinant expressed from the borrelial flgB promoter (PflgBkanR) flanked by both Tn5 and Tn7 ends (Figure 1). Plasmid pML102 was constructed by first PCR amplifying the PflgB-aadA cassette from pKFSS1 [26], which confers spectinomycin and streptomycin resistance in E. coli and B. burgdorferi, respectively, with BamHI and NotI engineered ends using the following oligonucleotide primers 5' ACGCGGATCCGAAAGATTTCCTATTAAGG 3' and 5' ACGCGCGGCCGCTTATTTGCCGACTACCTTGGT 3', where the BamHI and NotI sites are underlined and italicized respectively (since aadA confers resistance to streptomycin in B. burgdorferi [26], it will be referred to as strR for the remainder of this report). The PCR fragment was then cloned into pCR2.1-TOPO (Invitrogen Corp., Carlsbad, California, USA) to generate pCR2.1-PflgB-aadA, which was subsequently digested with BamHI and NotI and ligated into the Tn7 containing vector pGPS3 (New England Biolabs, Beverly, MA, USA) cut with the same restriction enzymes. The final construct, pML102, contains the PflgB-strR cassette flanked by Tn7 ends (Figure 1). Plasmid pML103 was constructed as follows. First, the PflgB-strR cassette from pKFSS1 was amplified by PCR by using the following oligonucleotides: 5' GAAAGATTTCCTATTAAGG 3' and 5' TTATTTGCCGACTACCTTGGT 3'. The resulting PCR fragment was cloned into pCR2.1-TOPO to generate pTA102. pTA102 was then digested with EcoRI (which flank the cloned PCR fragment) and ligated into pMOD<MCS> (Epicentre Technologies, Madison, WI, USA) also cut with EcoRI to generate pJS234. pJS234 was then digested with PvuII and cloned into EcoRV-digested pGPS3. The final plasmid construct, pML103, contains the PflgB-strR cassette flanked by Tn5 and Tn7 ends (Figure 1). To generate a customized transposon that delivers both an antibiotic resistance determinant and the infectivity restoration marker bbe22 (pncA), the same region of lp25 that was used by Purser et al. (i.e., containing bbe22 and bbe23; [25]) was PCR amplified and cloned into the pCR2.1-TOPO vector. This construct was digested with

28

J. Seshu et al. / The Isolation and Characterization of Isogenic Mutants

Figure 1. Customized transposon constructs used to mutagenize B. burgdorferi genes in vitro. The customized transposons encoded by pJS104 (A), pML102 (B), and pML103 (C) are shown. All constructs shown use the pGPS3 vector backbone. Each customized transposon encodes antibiotic resistance markers (ARM) that employ the strong B. burgdorferi flgB promoter (PflgB) to express either a kanamycin (kanR) or streptomycin (strR) resistance determinant. The location of the Tn5 and/or Tn7 ends as well as restriction enzyme sites are indicated.

BamHI and PstI (which flank the PCR cloning site) and ligated into the Tn5 minitransposon vector pMOD<MCS> (Epicentre Tech., Madison, WI, USA) digested with BamHI and PstI to generate pMOD-pncA. The PflgB-strR cassette from pTA102 (see above) was then digested with EcoRI and the PflgB-strR cassette was ligated into the sole EcoRI site of pMOD-pncA. The resulting construct (pMOD-pncA-PflgB-strR) was

Figure 2. Customized transposon encoded by pJS269. This customized transposon is derived from pML103 (Figure 1) and contains (i) the antibiotic resistance marker (ARM) to streptomycin (strR) excpressed from the B. burgdorferi flgB promoter and (ii) the region from lp25 that functions as an infectivity restoration marker (IRM) in lp25-deficient strains. The restriction sites and transposon (Tn) ends are also indicated.

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29

subsequently digested with PvuII (which flank the pMOD<MCS> Tn5 ends) and cloned into EcoRV cut pGPS3 (New England Biolabs, Beverly, MA, USA). The resulting construct, pJS269, contains PflgB-strR with bbe22 (pncA) and bbe23 flanked by both Tn5 and Tn7 ends (Figure 2). 1.4. Gene Targets for Transposition For all targets of in vitro transposition, an approximate 3 to 4 kb fragment was amplified by PCR and ligated into either pGEM-T (Promega Inc., Madison, WI, USA) or the pCR2.1-TOPO vector (Invitrogen Corp., Carlsbad, CA) with the gene of interest in the middle of the amplified fragment. The genes targeted include: bosR, ospA, oms28, and bbk32. The oligonucleotide primers used for the amplication of bosR and bbk32 fragments have been published [19, 27]. The oligonucleotide primers used for ospA and oms28 were for ospA, 5' ATGAACAATAAAATGAATA 3' and 5' TTGGGCTTGGGTGAAGGA 3' (final construct pJS262); and for oms28, 5' GAATTTCTTGGGGAACTTC 3' and 5' ATCAATTCCTAGCTTATC 3' (final construct pJS263). Both the ospA and oms28 containing PCR fragments were cloned separately into the pCR2.1-TOPO vector. 1.5. In vitro Transposition and Characterization In vitro transposition was conducted essentially as previously described [19, 27]. Briefly, in vitro mutagenesis was carried out by using the GPS-Mutagenesis System (New England Biolabs) according to the manufacturer's instructions with the appropriate borrelial gene as the target and either pML102 or pJS269 as the donors. Donor plasmids were selectively eliminated by digestion with PI-SceI that recognizes a unique 28 bp sequence present in the pGPS3 vector backbone present in both pML102 and pJS269. Transposed constructs were then electroporated into E. coli strain TOP10 and selected for ampR (at 100 Pg/ml) and spectinomycin resistance (at 50 Pg/ml). Transformants were subjected to restriction digestion analysis to crudely map the location of the transposon. In some instances, constructs were screened by PCR to confirm a transposition event using oligonucleotide primers that hybridize to the transposon and the inactivated borrelial gene. Appropriate candidates were dideoxy sequenced to determine the site of transposition. Exact constructs used for transformation and allelic exchange of bosR and bbk32 have been previously reported [19, 27]. The final knockout constructs for ospA and oms28 were designated pJS270 and pJS281, respectively. 1.6. Genetic Complementation The genetic complementation of the bbk32::strR mutant was conducted as previously described [19]. Briefly, a fragment containing 1 kb upstream of the initiation codon and 142 bp downstream from the stop codon of bbk32 was amplified by PCR with BamHI sites engineered on their ends and cloned into the sole BamHI site of pBBE22 [19, 25]. Transformants were selected in E. coli in the presence of kanamycin at 50 ȝg/ml and, following confirmation, transformed into B. burgdorferi bbk32::strR and selected in BSK-II agar with kanamycin at 300 ȝg/ml.

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1.7. Transformation of B. burgdorferi B. burgdorferi cells were made competent as previously described [19, 27, 28]. For allelic exchange, 20í30 ȝg of linearized DNA, containing the inactivated gene of interest with approximately 1 to 1.5 kb of flanking DNA, was electroporated into 109 competent B. burgdorferi as previously described [19, 27, 28]. For shuttle vector constructs, 5í10 ȝg of supercoiled DNA was used. Following electroporation, the samples were diluted in BSK-II media without antibiotic and incubated overnight at 32˚C. After the overnight incubation, B. burgdorferi were plated in BSK-II agarose overlays containing the appropriate antibiotic, either streptomycin at 50 ȝg/ml or kanamycin at 300 ȝg/ml. 1.8. Southern Blot Analysis Isolation of total genomic DNA and subsequent Southern blot analysis was conducted as previously described [19, 27]. 1.9. SDS-PAGE and Immunoblot Analysis SDS-PAGE and Western immunoblotting were conducted as described previously [19, 27]. To characterize the ospA and oms28 mutants using immunoblot analysis, the OspA monoclonal antibody H3TS (kindly provided by Tom Schwan, Rocky Mountain Laboratory, Hamilton, MT, USA) and a rabbit polyclonal antibody to Oms28 [29] were used, respectively. To detect immobilized immunes complexes, either anti-mouse immunoglobulin (Ig) conjugated to horseradish peroxidase (+HRP) or anti-rabbit Ig+HRP (both from GE Healthcare Bio-Sciences Corp., Piscataway, NJ, USA) were employed and developed using the ECL system as previously described [19, 27].

2. Results 2.1. Construction of Customized Transposons to Inactivate Borrelial Gene Targets To facilitate a random in vitro mutagenesis strategy to inactivate specific Borrelia burgdorferi “target” genes, several plasmid-based transposons were constructed to serve as “donors” containing selectable antibiotic resistance markers (ARM) expressed from the strong borrelial flgB promoter (Figures 1 and 2). It is important to note that even though the ARM constructs are transcriptional fusions to a strong borrelial promoter, these antibiotic resistance determinants are expressed and functional in E. coli [26, 30]. As such, the transposon knock-out mutants obtained can be screened easily in E. coli. This approach allows for the isolation of numerous transpositional “hits” within the appropriate “target” sequences, in this case borrelial genes. In general, the “target” B. burgdorferi gene was placed in the middle of a 3 to 4 kb fragment such that, following mutagenesis and characterization of the desired genetic knockout, the approximate 1í1.5 kb flanking sequences would aid in the subsequent double crossover event required for allelic exchange [19, 27]. The advantage of this methodology is that numerous distinct insertions can be isolated quickly within the gene of interest (in both

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coding and non-coding regions), as well as in adjacent genes to either address or exclude polarity issues. Several constructs with either Tn7 ends alone or with both Tn5 and Tn7 ends together were designed that encode antibiotic resistant determinants. These customized transposon constructs conferred resistance to either kanamycin or streptomycin in B. burgdorferi upon allelic exchange depending on the initial plasmid utilized to inactivate the borrelial gene (either pJS104 [kanR], pML102 or pML103 [both strR]; Figure 1). Tn5 and Tn7 transposon ends were chosen because the transposase enzymes are commercially available (i.e., the Tn5-specific enzyme EZ-Tn5 from Epicentre Technologies and the Tn7-specific TnsABC from New England Biolabs). The advantage of the commercial Tn5 and Tn7 transposases is that both are functional in vitro and thus require no additional host factors for transposition to occur. All that is needed is the customized transposon construct, the genetic target, and the transposase and appropriate buffer conditions (provided by the manufacturer). An additional plasmid encoded customized transposon, designated pJS269, was designed to deliver both the streptomycin resistance determinant and the bbe22 locus together, which simultaneously facilitated the inactivation of the targeted gene by (i) providing a selectable marker following transformation and allelic exchange (i.e., ARM) and (ii) restoring infectivity in the lp25- parental strain used in these studies (i.e., IRM) (Figure 2). Obviously, mutant strains generated using the pJS269 construct would not require the introduction of pBBE22 encoded IRM since the bbe22 locus would be integrated into the chromosome. Thus, subsequent genetic complementation would be restricted to providing back an intact and expressed form of the inactivated gene through the use of a shuttle vector such as pBSV2 [31]. 2.2. Inactivation of Borrelial Genes by In Vitro Transposition The B. burgdorferi genes analyzed to date using the in vitro transposition method described herein include bosR, bbk32, ospA, and oms28. The details regarding the in vitro transpositional inactivation of bosR (specifically the bosRR39K allele) using pJS104 as the source of the customized transposon was previously published [27]. Since the resulting bosRR39K::kanR construct was recombined into a non-infectious B. burgdorferi strain B31 clonal isolate [27], these constructs and their strain derivatives will not be discussed further in this report. The bbk32 gene encodes a surface-exposed fibronectin-binding lipoprotein in B. burgdorferi that has been purported to be important in Lyme pathogenesis. Previous studies involving BBK32 have been limited to either biochemical analysis or bbk32 expression in different host environments [32í37]. Although these analyses have provided important information pertaining to functional activities and in vivo expression as they relate to BBK32 and bbk32, respectively, the absence of genetic evidence has made it difficult to establish a clear link for BBK32 in B. burgdorferi pathogenesis. As a first step toward isolating a B. burgdorferi mutant in bbk32, a 2954 bp fragment, containing the 1062 bp bbk32 gene in the center of the fragment, was PCR amplified and cloned into pCR2.1-TOPO to generate pMLK32 [19]. Subsequently, bbk32 was inactivated in vitro using the customized Tn7 containing transposon that confers spectinomycin resistance in E. coli (i.e., pML102). Although a large number of transposon-containing clones were obtained in bbk32 following transformation into E. coli, transposon insertions that mapped to the 5’ end of bbk32 (based on restriction digestion analysis) were selectively analyzed by sequencing to determine the exact site

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of transposition. Five transposition mutants that mapped within the first 39 codons of bbk32 were identified (data not shown). These 5’ specific transposon insertions were preferable since they would not synthesize much of the processed (mature) form of BBK32. Ultimately one construct, containing an insertion between codon 10 and 11 of the bbk32 coding sequence (designated pMLK32-192) was used for allelic exchange [19]. Residues 10 and 11 of BBK32 reside within the leader peptide of this lipoprotein antigen, thus this construct, upon recombination via a double crossover into the B. burgdorferi genome, would be incapable of synthesizing an exported or surface exposed form of this protein. A similar construct was made to inactive the ospA (bba15) gene. OspA is the most abundantly produced lipoprotein during in vitro cultivation of B. burgdorferi but is not synthesized early in mammalian infection [18, 38í41]. Recently, OspA has been shown to be involved in the arthropod phase of the B. burgdorferi infectious cycle where it functions as an adhesin within the midgut of an infected Ixodes spp. tick [42, 43]. To inactivate ospA, a 3012 bp fragment with a centrally placed 822 bp ospA gene was amplified by PCR and cloned into pCR2.1-TOPO, generating pJS262. The customized transposon construct, pJS269, which delivered the streptomycin resistance determinant and the infectivity restoration marker bbe22 (IRM) simultaneously, was used to mutagenize pJS262. Several transposon hits were crudely mapped, and one with an insertion at bp 404 of the ospA gene was used for allelic exchange. The final construct containing the ospA::strR-IRM locus was designated pJS270. The B. burgdorferi oms28 gene (bba74) was also inactivated as described above for ospA. Oms28 has pore-forming activity in a synthetic bilayer assay [29] and may be secreted from B. burgdorferi cells [44], suggesting that it may function as a

Figure 3. Putative transposon mutants in oms28 generated in vitro using NEB’s GPS-MTM mutagenesis system. The plasmid pJS269 (Figure 2) was used as the transposon donor, and pJS263, containing the oms28 locus, served as the target in the mutagenesis reactions. Following selective reduction of pJS269 donor construct, the pJS263 mutagenized plasmids were transformed into E. coli under selection. The resulting transformants were analyzed by restriction analysis with ScaI and PstI, and a random pattern indicative of the different transpositional events was observed. The arrows indicate the location of the fragments of the nonmutagenized target plasmid pJS263 digested with ScaI and PstI. The numbers on the left refer to the size of markers in kb.

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pore-forming toxin. To begin mutagenesis of oms28, a 4884 bp fragment containing oms28 was PCR amplified and cloned into pCR2.1-TOPO to generate pJS263. As with ospA, the oms28 construct was mutagenized in vitro using pJS269 as the source for the customized transposon. A typical restriction pattern indicative of random transpositional events with the pJS263 target plasmid is shown in Figure 3. One particular transposon insertion was mapped to the first 50 bp of the oms28 gene, and the resulting oms28::strR-IRM construct used for allelic exchange was designated pJS281. 2.3. Allelic Exchange of Low Passage B. burgdorferi Clonal Isolate ML23 The genetic background used for all of the allelic exchange studies discussed in this report is the strain B31-derived clonal isolate ML23 that lacks only the 25 kb linear plasmid lp25 [22, 23]. Earlier work indicated that lp25 is essential for survival in the murine model of infectivity and encodes a restriction/modification (R/M) system [2225, 45]. Specifically, strains that are missing lp25 are significantly more readily transformable relative to parent strains containing all known plasmids [45], but are noninfectious [22í24]. Purser et al. demonstrated that the region on lp25 including bbe22, which encodes a nicotinamidase (and which is termed IRM for infectivity restoration marker in this report), is sufficient to complement the infectivity deficit associated with the strain lacking lp25, presumably because B. burgdorferi requires de novo synthesis of NAD during mammalian infection to regenerate this cofactor for important metabolic pathways including glycolysis [25]. To test the hypothesis that the lp25 mutant was more amenable to allelic exchange, all of the aforementioned constructs (i.e., bbk32::strR from pMLK32-192, ospA::strRIRM from pJS270 and oms28::strR-IRM from pJS281) were linearized, transformed by electroporation into strain ML23, and plated in BSK-II agar overlays containing appropriate antibiotics. For most transformations, approximately 20í40 transformants were obtained from 1 x 109 competent B. burgdorferi cells that were electroporated with 20 μg of DNA, yielding a transformation frequency of between 1í2 x 10-9 per μg of DNA. Colonies appeared between 14 to 17 days following plating and individual colonies were inoculated into liquid BSK-II media under appropriate antibiotic selection. Once the cultures reached an early logarithmic growth phase, total genomic DNA was purified from individual candidates and analyzed by both PCR and Southern blot analyses. The bbk32::strR strain JS315 obtained using this approach has been described [19]. Following transformation with the ospA::strR-IRM construct, putative ospA mutants were selected for streptomycin resistance and screened for their ability to synthesize OspA. Total protein profiles indicated that the putative mutants did not synthesize an abundant 31 kDa protein consistent with the inactivation of ospA (Figure 4). The putative mutants do not make a 34 kDa protein, presumed to be OspB, consistent with the co-transcription of ospA and ospB and the polar effect of ospB expression due to ospA inactivation [46]. Subsequent Western immunoblot and Southern blot analysis confirmed that the putative mutants did not synthesize OspA and had a transposon insertion within ospA, respectively (Figure 4). Figure 5 shows a PCR of 2 putative insertions in oms28 using oligonucleotide primers that amplify the oms28 open reading frame. Note the approximate 3.6 kb increase in the PCR amplified product, relative to the parental

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Figure 4. Transposon mutation in ospA. (A) Schematic representation of the transposon insertion in ospA (bba15). The position of transposon insertion is depicted by an open circle. The ARM + IRM functions as a single transposable unit as shown in Figure 2. (B) Southern blot of wild-type B. burgdorferi strain MSK5 (lane 1), the lp25 deficient parent ML23 (lane 2), and the ospA mutant JS270 (lane 3) probed with the strR marker. Numbers on the left represent the sizes of markers in kb. (C) Total protein profile of parent ML23 (lane 1) and ospA mutant JS270 (lane 2). Arrows indicate the location of OspA and OspB. (D) Immunoblot of samples shown in panel B, probed with antibodies specific for endoflagella (EF) and OspA.

strain, indicating the presence of the transposon within the oms28 coding sequence (Figure 5). Immunoblot analysis with Oms28-specific antiserum indicated that the putative oms28::strR-IRM did not synthesize Oms28, whereas the parental strain did (data not shown). Although lp25 is essential for the survival of B. burgdorferi in the mouse animal model of infection [22-24] and most likely for other mammalian species, one of the 28 kb linear plasmids, designated lp28-1, is also required for wild-type infectivity in animal models of infection [22-24], presumably because lp28-1 encodes the vls locus that is involved in antigenic variation and immune evasion and/or persistence [47, 48]. To assess whether lp28-1 was present in the putative mutants, PCR using lp28-1 specific primers were utilized (Figure 5). In all mutants tested, lp28-1 was retained based on PCR analysis (Figure 5). Subsequent immunoblot analysis indicated that the mutants synthesize the same amount of VlsE as the parental strain (data not shown). 2.4. Genetic Complementation of Low Passage B. burgdorferi Mutants The genetic background used for allelic exchange is missing the lp25 plasmid and is thus non-infectious independent of the mutation introduced. Therefore, to generate a genetic background comparable to the infectious parent strain ML23/pBBE22, the mutants obtained were first transformed with the infectivity restoration marker containing plasmid pBBE22 and then, separately, the same plasmid with an intact version of the inactivated gene. The overall scheme used is shown in Figure 6. The genetic complementation of the bbk32::strR mutant has been described previously [19].

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Figure 5. PCR analysis of oms28 mutants JS281-2 and JS281-3. Total genomic DNA was extracted from oms28 mutants JS281-2, (lane 2), JS281-3, (lane 3), ML23 (lane 4), and MSK5 (lane 5), and processed for PCR using oligonucleotide primers specific for oms28, bbe22, PflgB-strR, and lp28-1 as indicated above each set of samples. No template negative controls are found in lane 6 for the oms28, bbe22, and lp28-1 panels, and in lane 7 for the PflgB-strR panel. The plasmid pJS281 (the oms28::strR-IRM construct that served as a source for the allelic exchange) was included as a positive control for the PflgB-strR panel in lane 6. Numbers on the left indicated the molecular size of DNA markers in kb.

Constructs were made with sequence 400 bp to 1 kb upstream from the translational start of bbk32, bbk32, and 142 bp downstream from the bbk32 stop codonby cloning these fragments into the unique BamHI site of pBBE22 [25]. The large upstream region was included since it is not known which cis acting sequences are required for bbk32 expression. The final constructs were then transformed into the bbk32::strR strain JS315 and transformants were selected with kanamycin. Putative transformants were screened for BBK32 production using specific antiserum [19]. Six of the 15 putative transformants tested synthesized BBK32. The final genetic complement putative transformants tested synthesized BBK32. The final genetic complement obtained was then analyzed in vitro and in vivo relative to its parent and the bbk32::strR strain [19]. Complementation of the ospA::strR-IRM strain JS270 was not conducted since a comparable strain was reported [18]. The genetic complement to the oms28:strR-IRM strain JS281 is currently being constructed. 2.5. Characterization of Mutants The phenotype of the bbk32::strR strain JS315/pBBE22 and its genetic complement relative to the isogenic parent has been described previously [19]. One aspect of the bbk32 knockout analysis that merits mentioning is the restoration of infectivity in the genetic complement to levels that are in agreement with the isogenic parent strain as well as infectious B. burgdorferi that contain all plasmids, including lp25 [22í24], thus providing further corroboration for the utility of this approach.

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Figure 6. The mutagenesis strategy employed in this study to generate isogenic mutants in B. burgdorferi. As detailed in the text, borrelial genes (indicated by the plasmid encoded white boxes) are inactivated by transposon-based in vitro mutagenesis (indicated by the gray box), screened in E. coli for the desired transpositional mutation, and the final construct used to transform the lp25 deficient strain ML23. ML23 is the genetic parent for all transformations conducted and is more amenable to allelic exchange due to the absence of an lp25-encoded restriction/modification system. Putative mutants in B. burgdorferi are obtained following selection for the antibiotic resistance marker introduced by the transposon. Previous studies indicated that pBBE22, containing the lp25 bbe22 (pncA) locus on a 2 kb fragment (indicated by the plasmid encoded dark boxes), is sufficient to restore infectivity in strains lacking lp25 (referred to as IRM). To determine whether the mutation introduced in the gene of interest has an in vivo phenotype, the resulting mutant is transformed with pBBE22 alone (to restore infectivity via IRM) and with an intact and expressed version of the gene of cloned into pBBE22 to genetically complement the mutated form of the gene and provide IRM simultaneously. The resulting strains are then used to infect mice, and the 50% infectious dose determined.

The ospA::strR-IRM strain JS270 exhibits a serum sensitive phenotype such that cultures grown using non-heat inactivated rabbit serum did not replicate and were killed over time (data not shown). In contrast, the parent strain ML23/pBBE22 grew unabated, regardless of whether the rabbit serum used for cultivation was inactivated or not. This phenotype is consistent with the serum sensitivity reported for the strains B313 and B314 that do not synthesize OspA or OspB [49]. However, these strains are missing numerous linear plasmids, so this phenotype as it relates to OspA and OspB relative to other plasmids encoded proteins, was not clearly established [49]. The selective inactivation of the ospA locus and the serum sensitive phenotype observed for this mutation indicates that OspA and/or OspB are required for rabbit serum resistance in B. burgdorferi.

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The Oms28 protein has pore-forming activity [29] and can be secreted from B. burgdorferi [44], suggesting that Oms28 may be a membrane active toxin. To date, no in vitro phenotype was determined, and additional in vivo infectivity studies are pending the isolation of the genetically complemented clone of the oms28::strR-IRM strain JS281.

3. Discussion The genetic manipulation of Borrelia burgdorferi has improved considerably in the past few years due to the development of several antibiotic resistance markers, under the control of either borrelial or heterologous promoters, as well as the construction of shuttle vector replicons [20, 26, 30, 31, 50]. Despite these advances, isolating isogenic mutants is still difficult due to potent borrelial restriction/modification systems and the concurrent loss of plasmids (or mini-chromosomes) required for downstream infectivity analyses with B. burgdorferi [22í24]. Previous studies indicated that two linear plasmids, 25 kb and 28 kb in length and designated lp25 and lp28-1, are required for infectivity in needle-inoculated mice [22í24]. One of the restriction/modification systems is encoded on lp25 and thus cells that lack this plasmid are more easily transformed, but non-infectious [22í24, 45]. Subsequently, Purser et al. demonstrated that a fragment from lp25 that contains a gene encoding a nicotinamidase enzyme can be cloned into a shuttle vector (creating pBBE22) and effectively complement the infectivity defect observed in strains missing lp25 [25]. The activity of the nicotinamidase is apparently essential for de novo NAD biosynthesis [25], and cells lacking this activity are cleared between 24 to 48 hours following infection [22]. Thus, an lp25 deficient genetic background would be predicted to be more amenable to genetic transformation and, following subsequent transformation with plasmid borne bbe22 (i.e., pBBE22; [25]), would result in an comparable background to evaluate the role of the given inactivated gene in Lyme pathogenesis relative to its isogenic infectious parent. To test this hypothesis, a clonal isolate lacking lp25 (strain ML23) was electroporated with individually inactivated borrelial genes using linear DNA fragments and the desired recombination event selected with the appropriate antibiotic. Although the frequency of transformation was low (i.e., 1-2 x 10-9 per μg of DNA), this approach allowed for the isolation of putative mutants, whereas transformations of fully virulent B. burgdorferi containing all known plasmids yielded no transformants. Following the isolation of the putative mutants and their subsequent confirmation by molecular methodologies, the mutants were genetically complemented for two distinct activities. First, the mutant was transformed with pBBE22 alone to genetically complement the aforementioned lp25 infectivity associated defect such that the effect of the genetic knockout could be assessed relative to the non-mutagenized parental background. Second, the mutant is transformed with pBBE22 containing an intact form of the inactivated gene to attempt to restore the lost activity due to the mutation and complement the lp25-specific infectivity defect simultaneously. Recently, this approach has been used to evaluate the role of bbk32 in B. burgdorferi pathogenesis [19]. It should be emphasized that the methodology described here represents an alternative method for isolating mutants in B. burgdorferi. Several other investigators have used different strain backgrounds to isolate mutations in different borrelial genes [7í21]. In general, the genetic backgrounds used are infectious clonal isolates from strains 297 or B31 that contain all or most of the linear and circular plasmids. One

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exception is the B31-A3 strain that is missing the 9 kb circular plasmid cp9; however, since cp9 is not required for experimental infection in mice, this genetic background is well suited for infectivity analyses [8, 10, 20]. More recently an additional strain was isolated with an insertion in the borrelial bbe02 gene that encodes the restriction/modification system on lp25 [13]. As expected, the resulting bbe02::kanR strain was more competent for transformation and retained infectivity since the remainder of lp25, specifically the bbe22 locus or infectivity restoration marker (IRM), was present within the final strain construct. The advantage of this strain over the system reported herein using the lp25 deficient strain ML23 is the presence of the additional lp25 genes that may be important in other phases of the infectious cycle. Along these lines, a recent publication demonstrated that the lp25 encoded bbe16 locus was required for optimal colonization of Ixodes ticks [15]. Thus, the methodology described here is specific for the mammalian side of the B. burgdorferi infectious cycle. One advantageous aspect of the mutagenic strategy outlined here is the in vivo selection of the shuttle vector due to the presence of IRM on the pBBE22 vector backbone. During in vitro cultivation, shuttle vectors, including those that encode the intact form of the mutagenized gene in question (i.e., the genetic complement), are easily maintained by simple antibiotic selection. However, if these strains are then used to infect animals, there is no guarantee that the shuttle vector (with or without the complemented gene) would be passed to daughter cells without access to antibiotics ad libitum, either in feed or water. Since the vector backbone used here, pBBE22, contains the infectivity restoration marker bbe22 locus (IRM; see Figure 6 for details), there is a positive selection imposed for this shuttle vector during animal infection independent of antibiotic selection. This built-in selection ensures that all required genetic elements are maintained and thus the subsequent in vivo phenotype observed (i.e., the 50% infectious dose) can be appropriately compared between the parent, mutant, and complemented samples. Although all the strains used in the infectivity analysis contained the bbe22 (or pncA) locus required for infectivity, it is also well established that lp28-1 is also required for optimal B. burgdorferi infection, presumably due to the antigenic variant locus vlsE that it encodes ([22]. In all instances to date, the mutants screened retained lp28-1 and vlsE (based on PCR) and produced VlsE both prior to and following mutagenesis and genetic complementation (data not shown). This is an important consideration, since any subsequent analyses, particularly following infection in mice, could then be evaluated in the context of the desired mutation and resulting mutant strain in a manner that is independent of lp28-1. The use of random transposition to mutagenize a given gene in vitro has some important advantages. First, this type of mutagenesis results in a large collection of genetic knock-outs in a short period of time in both the coding and non-coding regions of the gene of interest as well as in flanking genes. Second, insertions in a gene with multiple functional domains could facilitate in the analysis of truncated forms of the encoded protein lacking one (or several) of the aforementioned domains, allowing one to determine the importance of these motifs in the context of functional assays and/or borrelial pathogenesis. Finally, since the transposition event is recombination mediated, the multiple cloning steps/ligation reactions required to generate an engineered restriction site within the gene to be inactivated, and the subsequent inactivation of the gene with an antibiotic resistance determinant, are eliminated.

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In summary, an alternate strategy has been developed using a non-infectious strain B31 derivative with enhanced transformation efficiency (i.e., strain ML23 missing lp25) to isolate genetic knock-outs in B. burgdorferi. The premise of this approach is that mutants can be isolated in the restriction/modification deplete background and, following characterization to confirm the isolation of the desired mutation, the mutants can be tested for an infectivity defect by providing the bbe22-containing IRM locus in trans. Concurrent genetic complementation with an intact form of the inactivated gene provides all of the appropriate comparators to test the effect of a given gene in B. burgdorferi pathogenesis. As such, this strategy for analyzing mutations independently in specific B. burgdorferi genes could be applied to any genetic locus assuming that the gene(s) in question is(are) not required for in vitro cultivation.

Acknowledgements The authors thank Steve Norris for generously providing pBBE22. The authors are also grateful to Scott Samuels for the kind gift of pKFSS1, which served as the source of the streptomycin resistance determinant in these studies. This work was supported by United States Public Health Service grants AI65953 (to J.S.), AI20624 (to M.H.), AI42345 and AI58086 (both to J.T.S.) from the National Institute of Allergy and Infectious Diseases.

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[11] Hubner, A., Revel, A.T., Nolen, D.M., Hagman, K.E. and Norgard, M.V. (2003) Expression of a luxS gene is not required for Borrelia burgdorferi infection of mice via needle inoculation. Infect Immun 71, 2892í2896. [12] Hubner, A., Yang, X., Nolen, D.M., Popova, T.G., Cabello, F.C. and Norgard, M.V. (2001) Expressionof Borrelia burgdorferi OspC and DbpA is controlled by a RpoN-RpoS regulatory pathway. Proc Natl Acad Sci USA 98, 12724í12729. [13] Kawabata, H., Norris, S.J. and Watanabe, H. (2004) BBE02 disruption mutants of Borrelia burgdorferi B31 have a highly transformable, infectious phenotype. Infect Immun 72, 7147-7154. [14] Pal, U., Yang, X., Chen, M., Bockenstedt, L.K., Anderson, J.F., Flavell, R.A., Norgard, M.V. and Fikrig, E. (2004) OspC facilitates Borrelia burgdorferi invasion of Ixodes scapularis salivary glands. J Clin Invest 113, 220-230. [15] Revel, A.T., Blevins, J.S., Almazan, C., Neil, L., Kocan, K.M., de la Fuente, J., Hagman, K.E. and Norgard, M.V. (2005) bptA (bbe16) is essential for the persistence of the Lyme disease spirochete, Borrelia burgdorferi, in its natural tick vector. Proc Natl Acad Sci USA 102, 6972í6977. [16] Tilly, K., Grimm, D., Bueschel, D.M., Krum, J.G. and Rosa, P. (2004) Infectious cycle analysis of a Borrelia burgdorferi mutant defective in transport of chitobiose, a tick cuticle component. Vector Borne Zoonotic Dis 4, 159í168. [17] Yang, X.F., Alani, S.M. and Norgard, M.V. (2003) The response regulator Rrp2 is essential for the expression of major membrane lipoproteins in Borrelia burgdorferi. Proc Natl Acad Sci USA 100, 11001í11006. [18] Yang, X.F., Pal, U., Alani, S.M., Fikrig, E. and Norgard, M.V. (2004) Essential Role for OspA/B in the Life Cycle of the Lyme Disease Spirochete. J Exp Med 199, 641-648. [19] Seshu, J., Esteve-Gassent, M.D., Labandeira-Rey, M., Kim, J.H., Trzeciakowski, J.P., Hook, M. and Skare, J.T. (2006) Inactivation of the fibronectin-binding adhesin gene bbk32 significantly attenuates the infectivity potential of Borrelia burgdorferi. Mol Microbiol 59, 1591í1601. [20] Elias, A.F., Stewart, P.E., Grimm, D., Caimano, M.J., Eggers, C.H., Tilly, K., Bono, J.L., Akins, D.R., Radolf, J.D., Schwan, T.G. and Rosa, P. (2002) Clonal polymorphism of Borrelia burgdorferi strain B31 MI: implications for mutagenesis in an infectious strain background. Infect Immun 70, 2139í2150. [21] Caimano, M.J., Eggers, C.H., Hazlett, K.R. and Radolf, J.D. (2004) RpoS is not central to the general stress response in Borrelia burgdorferi but does control expression of one or more essential virulence determinants. Infect Immun 72, 6433í6445. [22] Labandeira-Rey, M., Seshu, J. and Skare, J.T. (2003) The absence of linear plasmid 25 or 28-1 of Borrelia burgdorferi dramatically alters the kinetics of experimental infection via distinct mechanisms. Infect Immun 71, 4608í4613. [23] Labandeira-Rey, M. and Skare, J.T. (2001) Decreased infectivity in Borrelia burgdorferi strain B31 is associated with loss of linear plasmid 25 or 28-1. Infect Immun 69, 446í455. [24] Purser, J.E. and Norris, S.J. (2000) Correlation between plasmid content and infectivity in Borrelia burgdorferi. Proc Natl Acad Sci USA 97, 13865í13870. [25] Purser, J.E., Lawrenz, M.B., Caimano, M.J., Howell, J.K., Radolf, J.D. and Norris, S.J. (2003) A plasmid-encoded nicotinamidase (PncA) is essential for infectivity of Borrelia burgdorferi in a mammalian host. Mol Microbiol 48, 753í764. [26] Frank, K.L., Bundle, S.F., Kresge, M.E., Eggers, C.H. and Samuels, D.S. (2003) aadA confers streptomycin resistance in Borrelia burgdorferi. J Bacteriol 185, 6723í6727. [27] Seshu, J., Boylan, J.A., Hyde, J.A., Swingle, K.L., Gherardini, F.C. and Skare, J.T. (2004) A conservative amino acid change alters the function of BosR, the redox regulator of Borrelia burgdorferi. Mol Microbiol 54, 1352í1363. [28] Samuels, D.S. (1995) Electrotransformation of the spirochete Borrelia burgdorferi, in Methods in Molecular Biology 47: 253í259. Humana Press, Totowa, New Jersey, USA. [29] Skare, J.T., Champion, C.I., Mirzabekov, T.A., Shang, E.S., Blanco, D.R., Erdjument-Bromage, H., Tempst, P., Kagan, B.L., Miller, J.N. and Lovett, M.A. (1996) Porin activity of the native and recombinant outer membrane protein Oms28 of Borrelia burgdorferi. J. Bacteriol. 178, 4909í4918. [30] Bono, J.L., Elias, A.F., Kupko, J.J., 3rd, Stevenson, B., Tilly, K. and Rosa, P. (2000) Efficient Targeted Mutagenesis in Borrelia burgdorferi. J Bacteriol 182, 2445í2452. [31] Stewart, P.E., Thalken, R., Bono, J.L. and Rosa, P. (2001) Isolation of a circular plasmid region sufficient for autonomous replication and transformation of infectious Borrelia burgdorferi. Mol Microbiol 39, 714í721. [32] Fikrig, E., Feng, W., Barthold, S.W., Telford, S.R., 3rd and Flavell, R.A. (2000) Arthropod- and hostspecific Borrelia burgdorferi bbk32 expression and the inhibition of spirochete transmission. J Immunol 164, 5344í5351.

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[33] Kim, J.H., Singvall, J., Schwarz-Linek, U., Johnson, B.J., Potts, J.R. and Hook, M. (2004) BBK32, a fibronectin binding MSCRAMM from Borrelia burgdorferi, contains a disordered region that undergoes a conformational change on ligand binding. J Biol Chem 279, 41706í41714. [34] Liang, F.T., Nelson, F.K. and Fikrig, E. (2002) Molecular adaptation of Borrelia burgdorferi in the murine host. J Exp Med 196, 275í280. [35] Probert, W.S. and Johnson, B.J. (1998) Identification of a 47 kDa fibronectin-binding protein expressed by Borrelia burgdorferi isolate B31. Mol Microbiol 30, 1003í1015. [36] Probert, W.S., Kim, J.H., Hook, M. and Johnson, B.J. (2001) Mapping the ligand-binding region of Borrelia burgdorferi fibronectin- binding protein BBK32. Infect Immun 69, 4129í4133. [37] Raibaud, S., Schwarz-Linek, U., Kim, J.H., Jenkins, H.T., Baines, E.R., Gurusiddappa, S., Hook, M. and Potts, J.R. (2005) Borrelia burgdorferi binds fibronectin through a tandem beta-zipper, a common mechanism of fibronectin binding in staphylococci, streptococci, and spirochetes. J Biol Chem 280, 18803í18809. [38] Montgomery, R.R., Malawista, S.E., Feen, K.J. and Bockenstedt, L.K. (1996) Direct demonstration of antigenic substitution of Borrelia burgdorferi ex vivo: exploration of the paradox of the early immune response to outer surface proteins A and C in Lyme disease. J. Exp. Med. 183, 261í269. [39] Philipp, M.T. (1998) Studies on OspA: a source of new paradigms in Lyme disease research. Trends Microbiol 6, 44í47. [40] Schwan, T.G. and Piesman, J. (2000) Temporal changes in outer surface proteins A and C of the lyme disease-associated spirochete, Borrelia burgdorferi, during the chain of infection in ticks and mice. J Clin Microbiol 38, 382í388. [41] Schwan, T.G., Piesman, J., Golde, W.T., Dolan, M.C. and Rosa, P.A. (1995) Induction of an outer surface protein on Borrelia burgdorferi during tick feeding. Proc Natl Acad Sci USA 92, 2909í2913. [42] Pal, U., de Silva, A.M., Montgomery, R.R., Fish, D., Anguita, J., Anderson, J.F., Lobet, Y. and Fikrig, E. (2000) Attachment of Borrelia burgdorferi within Ixodes scapularis mediated by outer surface protein A. J Clin Invest 106, 561í569. [43] Pal, U., Li, X., Wang, T., Montgomery, R.R., Ramamoorthi, N., Desilva, A.M., Bao, F., Yang, X., Pypaert, M., Pradhan, D., Kantor, F.S., Telford, S., Anderson, J.F. and Fikrig, E. (2004) TROSPA, an Ixodes scapularis receptor for Borrelia burgdorferi. Cell 119, 457í468. [44] Cluss, R.G., Silverman, D.A. and Stafford, T.R. (2004) Extracellular secretion of the Borrelia burgdorferi Oms28 porin and Bgp, a glycosaminoglycan binding protein. Infect Immun 72, 62796286. [45] Lawrenz, M.B., Kawabata, H., Purser, J.E. and Norris, S.J. (2002) Decreased electroporation efficiency in Borrelia burgdorferi containing linear plasmids lp25 and lp56: impact on transformation of infectious B. burgdorferi. Infect Immun 70, 4798í4804. [46] Howe, T.R., LaQuier, F.W. and Barbour, A.G. (1986) Organization of genes encoding two outer membrane proteins of the Lyme disease agent Borrelia burgdorferi within a single transcriptional unit. Infect Immun 54, 207í212. [47] McDowell, J.V., Sung, S.Y., Hu, L.T. and Marconi, R.T. (2002) Evidence that the variable regions of the central domain of VlsE are antigenic during infection with Lyme disease spirochetes. Infect Immun 70, 4196í4203. [48] Zhang, J.R., Hardham, J.M., Barbour, A.G. and Norris, S.J. (1997) Antigenic variation in Lyme disease borreliae by promiscuous recombination of VMP-like sequence cassettes. Cell 89, 275í285. [49] Sadziene, A., Thomas, D.D. and Barbour, A.G. (1995) Borrelia burgdorferi mutant lacking Osp: biological and immunological characterization. Infect Immun 63, 1573í1580. [50] Sartakova, M., Dobrikova, E. and Cabello, F.C. (2000) Development of an extrachromosomal cloning vector system for use in Borrelia burgdorferi. Proc Natl Acad Sci USA 97, 4850í4855.

Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Motility Gene Regulation and Chemotaxis in Borrelia burgdorferi Nyles W. CHARON a,1 , Melanie SALa, Michael R. MILLER b, Richard G. BAKKER a, Chunhao LI a, and Md. Abdul MOTALEBa a Departments of Microbiology, Immunology, and Cell Biology b Biochemistry & Molecular Pharmacology, Robert C. Byrd Health Sciences Center West Virginia University, Morgantown, West Virginia 26506-9177 Abstract. Motility and chemotaxis are important in the disease process for many species of bacteria. When infected ticks feed, B. burgdorferi moves from the midgut to the salivary gland, and when uninfected ticks feed on infected animals, the spirochete migrates from the animal into the tick. Motility and chemotaxis are likely to be involved in this traveling of B. burgdorferi between the two hosts. Compared with the well-studied Escherichia coli, motility and chemotaxis in B. burgdorferi is quite complex: it has 2 cheA, 3 cheY, 2 cheB, 2 cheR and 2 fliG homologues. Instead of the E. coli phosphatase CheZ, B. burgdorferi has a CheY-P phosphatase called CheX. Furthermore, B. burgdorferi has to coordinate two bundles of periplasmic flagella, extending from opposite ends of the cell, to move toward attractants and away from repellents. B. burgdorferi exhibits different motility modes, consisting of runs, flexes, and reverses, which depend on movement of the bundles of periplasmic flagella. Analysis of the B. burgdorferi genome sequence revealed that it has no homologues of the sigma-28 or anti-sigma factor genes responsible for cascade control of motility gene regulation found in E. coli or Salmonella enterica serovar Typhimurium. Rather, studies indicate that motility genes in B. burgdorferi are regulated by sigma-70 promoters. To determine the role(s) of individual motility genes, we deployed targeted mutagenesis technology and analyzed the resulting cell phenotypes based on the flagellar structure, cell swimming behavior, and effect of specific gene inactivation on other genes and on chemotaxis. Novel functions of many of these genes of the Lyme disease organism are summarized. Keywords. Borrelia burgdorferi, periplasmic flagella, translational control, chemotaxis, CheA

Introduction Lyme disease is caused by the motile spirochete Borrelia burgdorferi [1]. This disease is the most prevalent arthropod borne infection in the United States, with approximately 23,763 cases reported in 2002 [2]. The life cycle of B. burgdorferi involves 1

Corresponding Author: Nyles W. Charon, Department of Microbiology, Immunology and Cell Biology, West Virginia University, Box 9177, Robert C. Byrd Health Sciences Center, Morgantown, West Virginia 26506-9177. Phone: (304) 293-4170. Fax (304) 293-7823. E-mail: [email protected].

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transmission from a tick vector to mammals and back to the tick over the course of several seasons [1]. The clinical course of B. burgdorferi infections include symmetrical spread of the spirochetes through the dermis resulting in a rash referred to as erythema migrans, and movement across facial planes to deep organs. Disease manifestations also include arthritis, cardiac abnormalities, and neuropathies [1, 3-5].

1. B. burgdorferi Motility Gene Regulation 1.1. B. burgdorferi Structure B. burgdorferi is a spirochete, and as such, has a characteristic cell morphology [6-8]. Its diameter is approximately 0.33 ȝm, and it is 10 to 20 ȝm long [6]. Thus, these bacteria are considerably larger than those of the well studied species Escherichia coli. B. burgdorferi has a characteristic flat-wave morphology, which is different than many other spirochete species which are helical [6-8]. Its wave-length is approximately 2.83 μm, and there are approximately 5-7 wave-lengths per undividing cell [6, 7]. Outermost is a membrane sheath, and within this sheath are the periplasmic flagella (PFs) and protoplasmic cell cylinder. A given PF is attached subterminally at only one end of the cell cylinder, and it overlaps in the center of the cell with those from the opposite end [9]. There are approximately 7 to 11 PFs attached near each cell end. Thus, there are two bundles of PFs with each bundle originating at a given cell end [9]. High voltage electron microscopy indicates that these PFs form a continuous bundle of 67 nm that extends from one end to the other. This bundle winds around the cell axis in a left handed sense with a helix pitch 2.83 μm, which is identical to the wave length of the cell as noted above [6]. 1.2. Periplasmic Flagellum Structure The PFs have been studied in some detail. In common with the flagella of other bacteria, a given PF is composed of a basal body, hook, and filament [8-11]. However, in contrast to E. coli and Salmonella entrica serovar Typhimurium flagella, which are composed of a polymer of a single protein component FliC [11], those from B. burgdorferi have a major flagellin protein designated as FlaB, and a minor flagellin protein referred to as FlaA [12-14]. The location of FlaA in B. burgdorferi is unknown, but in other spirochetes it forms a sheath around the FlaB core [8, 14-16]. The PF basal structure contains the characteristic MS and P rings found in those of other bacteria, but it lacks an L ring typical of Gram negative bacteria [8, 17]. In surveying the B. burgdorferi genome and the putative genes that encode the PF apparatus, one finds that there is extensive sequence identity to their counterparts found in other bacteria [8, 14, 18]. Thus, for example, FlgE, which comprises the hook protein of B. burgdorferi, has a 62% sequence similarity to its counterpart in E. coli. The purified PFs of B. burgdorferi are helical in shape, left-handed, and have a helix pitch of 1.48 Pm and a helix diameter of 0.28 Pm [19].

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1.3. Cytoskeletal and Motility Function of Periplasmic Flagella We have begun to understand the causal relationship between PFs, cell morphology, and motility. We know that the PFs are directly involved in motility [20, 21], and they propel the organisms by rotation between the outer sheath and cell cylinder [6, 7]. In addition, the PFs are clearly more rigid than the cell cylinder, as mutant cells that lack the PFs are rod-shaped [6, 20]. This flat-wave like morphology of the cell is independent of cell metabolism. Thus, proton ionophores and prolonged cold incubation result in cells that are non-motile but have the flat-wave appearance [7, 20]. Clearly, the PFs have a skeletal function. A model has been formulated which specifies how B. burgdorferi swims. Rotation of the bundles of PFs in opposite directions (see below), causes backward moving waves along the cell body that propel the bacteria forward [6-8, 14]. This coordinated rotation of the two PF bundles results in backward moving flat waves. These waves propel the spirochete in a given direction: the cell runs. During the run, the two bundles rotate in opposite directions, i.e., asymmetrically. The posterior end rotating CW [we use the frame of reference of viewing the PF from its distal end to where it inserts in the cell], and the anterior end going CCW. To balance the rotation of the PFs, the cell body rolls around the body axis in the opposite direction. Cells reverse directions by stopping and/or flexing; a flex occurs when both bundles rotate in the same direction (both in CW or CCW) [7, 8, 22]. 1.4. Periplasmic Flagellum Gene Regulation Flagellum synthesis, as described in E. coli and S. enterica serovar Typhimurium, is a finely orchestrated succession of motility gene expression and protein assembly that requires tight regulation [23, 24]. In most bacteria, flagella synthesis is regulated by a cascade of transcriptional events involving the ordered expression of Class I, Class II, and Class III motility genes [23-25]. Expression of class I genes, which comprise the master operon, directs the transcription of class II genes. Class II genes are those that encode structural proteins involved with the early stages of flagella synthesis and regulatory proteins controlling the expression of the late stage class III flagella and chemotaxis genes. Among the Class II regulatory genes are fliA and flgM, encoding the motility specific transcription factor sigma-28 and the anti-sigma-28 factor, respectively. FlgM and sigma-28 remain bound as a complex in the cytoplasm during basal body and hook synthesis. Upon completion of the hook, this structure provides an export route for the anti-sigma factor FlgM. As FlgM exits the cell, sigma-28 is free to initiate the transcription of late genes, including FliC. When an intact hook is not formed or is not functional, FlgM fails to escape into the medium and remains associated with sigma-28 to prevent the transcription of flagellin genes. Thus, completion of the basal body-hook structure acts as a general assembly checkpoint in the regulation of flagella synthesis whereby the flagellin genes are transcriptionally controlled in response to the state of hook completion [23, 24]. The regulation of PF synthesis in B. burgdorferi differs markedly from that in other species of bacteria. Homologs for the class II transcriptional regulatory genes fliA and flgM have not been identified, and no sigma-28 promoters are evident in the B. burgdorferi genome [13, 14, 18]. Furthermore, when analyzing promoter sequences in B. burgdorferi, no sigma-54 recognition sequences were detected in motility operons. Instead, this bacterium has sigma-70 promoters controlling its motility genes [8, 14,

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45

26]. Other than the apparent lack of flagella-specific transcriptional regulatory proteins utilized by other bacteria, little is known about the regulation of flagella synthesis in B. burgdorferi. Recently, efforts have been made to elucidate the events involved in PF regulation and synthesis. Motaleb et al. described an insertion mutant in the major flagellin gene flaB that does not make flagella, is non-motile, and has a rod-shaped morphology [20] (Table 1). Complementation of this mutant in cis and more recently in trans restored its motility and wave-like morphology [21] (M. Motaleb and N. Charonunpublished). The flaB mutant was characterized in detail by Western blot to assess the effects of this mutation on other motility and chemotaxis genes [13]. Interestingly, the only putative motility and chemotaxis protein altered with respect to accumulation in the flaB mutant was FlaA, the minor PF filament protein. The reason for this inhibition of FlaA is poorly understood, but results revealed that the decrease of FlaA in a flaB mutant occured post-transcriptionally. Specifically, evaluation of the flaB mutant by real-time RT-PCR revealed that the flaA message was transcribed at wild-type levels, indicating that the reduction of FlaA synthesis does not occur by transcriptional regulation. FlaA was turned over with a half-time of 2 hours in flaB mutant [13], so it is conceivable that the decrease in FlaA accumulation is related to either its turnover, by translational control, or both.

Table 1. Properties of flaB and flgE mutants and complements of B. burgdorferi. Periplasmic flagella

Motility

Morphology

FlaA

FlaB

flaA transcript

flaB transcript

Wild-type (B31A)

+

+

Wave-like

+

+

+

+

flab

-

-

Rod-shaped

Trace

-

+

-

flaB

+

+

Wave-like

+

+

+

+

flgE

-

-

Rod-shaped

Trace

Trace

+

+

+

+

Wave-like

+

+

+

+

Strain

+

+

flgE

1.5. Analysis of a flgE Mutation on Periplasmic Flagella Gene Regulation As described above in other bacteria, the hook is necessary for flagella structure and also provides a regulatory checkpoint during flagella synthesis [23, 24, 27]. We tested these two aspects in regard to the importance of the PF hook protein in B. burgdorferi. First, to test the hypothesis that the hook is critical for flagella based motility and the flat-wave morphology, we used targeted mutagenesis to inactivate the gene encoding the PF hook protein by insertion of a kanamycin resistance cassette [28, 29]. We found that the flgE mutant had similar morphological characteristics as the flaB mutant [20]: It lacked periplasmic flagella, was non-motile, and rod-shaped [29] (Table 1). We complemented the flgE mutant in trans, and this complemented strain restored morphology, motility, and protein synthesis to the wild-type level (see later) [29]. These results yield further evidence that the PFs have a skeletal function and are involved in motility.

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We investigated the effects of a flgE mutation on PF protein accumulation [29]. Western blot analysis indicated that both FlaA and FlaB were markedly decreased (greater than 70% compared to the wild-type) in the flgE mutant (Table 1). Conceivably, these decreases in FlaA and FlaB could be the result of a decrease in transcription of these genes. However, quantitative RT-PCR analysis indicated that mRNA levels of these genes were equivalent to those found in the wild type (Table 1); thus, transcription is not inhibited in the major flagellar genes as is found in flgE mutants in other bacteria. Using the protein synthesis inhibitor spectinomycin coupled to Western blot analysis, we found that the FlaB protein was stable in the mutant, whereas FlaA was turned over with a half-life of 2 hours. These results suggest that the decrease in FlaB accumulation is likely to be under translational control, whereas the decrease in FlaA is due to either protein turnover, translational control, or both [29].

2. B. burgdorferi Chemotaxis A robust motility and chemotaxis system is likely to be vital for B. burgdorferi in its overall life cycle. Approximately six per cent of its genome encodes putative chemotaxis and motility genes [8, 18], and between 10-14 % of the cellular total protein is comprised of flagellar filament proteins [13]. Many of the motility and chemotaxis genes are expressed both in the tick and the mammalian host [30-32]. Thus, this system is evidently important but energetically expensive to the spirochete. What are the possible roles of chemotaxis and motility in the B. burgdorferi life cycle? Motility may be important for the spirochetes to migrate from the tick gut to the salivary glands for deposition into the new host upon infection [31]. In the mammal, motility and chemotaxis may be necessary for the spirochetes to penetrate into the blood stream after being deposited in the skin by the tick bite, and also for specific tissue and organ localization [4, 33]. For the cycle to be completed, tick salivary proteins and other compounds could conceivably serve as chemoattractants during tick feeding [34, 35]. This chemotactic signaling would result in the spirochetes concentrating at the site of the tick bite for cycle continuation. Bacterial chemotaxis involves a sensory transduction system that enables cells to swim toward a favorable environment, or away from one that is toxic [36, 37]. Bacteria swim up an attractant gradient in a manner best described as a biased random walk. This walk is composed of runs (CCW rotation of the flagella), and tumbles (when one or more of the flagella rotates CW). In E. coli and S. enterica, which serve as model systems [36-39], a two-component regulatory system involving a histidine kinase (CheA) and response regulator (CheY) plays an essential role in chemotaxis. The phosphorylated form of the response regulator CheY (CheY-P) interacts with the switch complex at the flagellar motor to change direction of flagellar rotation from the default CCW state to a clockwise CW state. The level of CheY-P is regulated in part by chemical attractants and repellents. Attractants bind directly or indirectly to receptors, referred to as methyl accepting chemotaxis proteins, which are embedded in the cell membrane. This binding results in a decrease in CheA autophosphorylation and a concomitant reduction in the level of CheY-P. The reduction of CheY-P sustains CCW flagellar rotation, resulting in relatively long runs up the gradient during chemotaxis [36, 37, 39]. Spirochete motility is more complicated than that of E. coli. For these organisms to run, the two bundles need to rotate asymmetrically: one CCW, the other CW [8, 22]. In

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47

contrast, in E. coli and S. enterica, running occurs when all the flagella rotate CCW. When cheA of E. coli is inactivated, the concentration of CheY-P decreases, and such cells fail to tumble; their flagella constantly rotate CCW [40]. If this also occurs in B. burgdorferi, i.e., all the periplasmic flagella rotate CCW, we hypothesized that such cells should constantly flex. To test this hypothesis, we inactivated the cheA genes (cheA1 and cheA2) of B. burgdorferi [22]. Swarm plate and capillary tube chemotaxis assays indicated that only cheA2 was involved in chemotaxis [22] (Table 2). Complementation of cheA2 resulted in the regaining of chemotaxis (Li et al., unpublished) [41]. In contrast to our expectations, cheA2 mutants constantly ran [22]. Thus, in the default state (i.e., low CheY-P concentrations), cells rotated their bundles of PFs asymmetrically [22].

Table 2. Selected properties of B. burgdorferi cheA1, cheA2, and cheX mutants and complements. Strain

Chemotaxis

Runs

Stops/Flexes

Reverses

Wild-type (B31A)

+

+

+

+

cheA1

+

+

+

+

cheA2

-

+

-

-

+

+

+

+

+

cheA1-cheA2

-

+

-

cheA2

cheX

-

-

+

cheX+

+

+

+

1

1

+

cheX constantly flexes.

We developed a new hypothesis relative to the results obtained [8, 22]. We hypothesized that there were differences in the motors at both ends of the cells. We proposed that certain factors associated with the motors at the old cell ends but not the new cell ends. This association resulted in these motors rotating in the CW rather than the CCW direction in the default state. This hypothesis is consistent with the results that certain proteins in other bacterial species localize at specific ends of the cells [4244]. As a corollary to this hypothesis, we predicted that cells with a high CheY-P concentration should constantly run in the opposite direction, i.,e. those motors that rotated CCW in the low CheY-P environment should now rotate CW when the CheY-P concentration is high, and those that rotated CW in low CheY-P concentration should rotate in CCW when the environment is high in CheY-P. To test this hypothesis, we inactivated a gene that was expected to produce high CheY-P in the B. burgdorferi cells. CheX is a phosphatase that specifically dephosphorylates CheY-P [45]. In E. coli, inactivation of the gene that encodes the phosphatase that acts on Che-Y-P (cheZ) results in cells that that constantly tumble [46]; their CheY-P concentration is high, and their flagella rotate in CW. In contrast to our expectation, we found that a mutant in cheX constantly flexed [45] (Table 2). Thus, the simple hypothesis that motors at both ends of the cell are different cannot simply explain the phenotype of the cheA2 mutant. Clearly, we are far from obtaining a molecular understanding of spirochete chemotaxis.

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Acknowledgements We thank R. Bourret, S. Goldstein, and R. Silversmith for their interest and support. This research was supported by U.S. Public Health Service grants AI29743 to N.W.C., AR050656-01 to C.L. and RR16440 to the West Virginia University Flow Cytometric Core Facility; West Virginia University Health Science Center Internal Grants, Office of Research and Graduate Education to M.R.M and WVU School of Medicine MD/PhD program to RB; American Heart Association grant 0365225B to M.A.M.

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Li,C., Bakker,R.G., Motaleb,M.A., Sartakova,M.L., Cabello,F.C., Charon,N.W., Asymmetrical flagellar rotation in Borrelia burgdorferi nonchemotactic mutants., Proc. Natl. Acad. Sci. U. S. A., 99 (2002) 6169-6174. Aldridge,P., Hughes,K.T., Regulation of flagellar assembly, Curr. Opin. Microbiol., 5 (2002) 160-165. Chilcott,G.S., Hughes,K.T., Coupling of flagellar gene expression to flagellar assembly in Salmonella enterica serovar Typhimurium and Escherichia coli, Microbiol. Mol. Biol. Rev., 64 (2000) 694-708. Anderson,D.K., Newton,A., Posttranscriptional regulation of Caulobacter flagellin genes by a late flagellum assembly checkpoint., J. Bacteriol., 179 (1997) 2281-2288. Ge,Y., Old,I.G., Saint Girons,I., Charon,N.W., Molecular characterization of a large Borrelia burgdorferi motility operon which is initiated by a consensus V70 promoter, J. Bacteriol., 179 (1997) 2289-2299. Bonifield,H.R., Yamaguchi,S., Hughes,K.T., The flagellar hook protein, FlgE, of Salmonella enterica serovar typhimurium is posttranscriptionally regulated in response to the stage of flagellar assembly, J. Bacteriol., 182 (2000) 4044-4050. Bono,J.L., Elias,A.F., Kupko III,J.J., Stevenson,B., Tilly,K., Rosa,P., Efficient targeted mutagenesis in Borrelia burgdorferi., J. Bacteriol., 182 (2000) 2445-2452. Sal, M.S., Periplasmic flagella of the spirochetes Borrelia burgdorferi and Brachyspira hyodysenteriae. Ph.D. Dissertation, West Virginia Univ. 2005. Revel,A.T., Talaat,A.M., Norgard,M.V., DNA microarray analysis of differential gene expression in Borrelia burgdorferi, the Lyme disease spirochete, Proc. Natl. Acad. Sci. U. S. A, 99 (2002) 15621567. Fisher,M.A., Grimm,D., Henion,A.K., Elias,A.F., Stewart,P.E., Rosa,P.A., Gherardini,F.C., Borrelia burgdorferi sigma54 is required for mammalian infection and vector transmission but not for tick colonization, Proc. Natl. Acad. Sci. U. S. A, 102 (2005) 5162-5167. Fikrig,E., Chen,M.C., Barthold,S.W., Anguita,J., Feng,W., Telford,S.R., III, Flavell,R.A., Borrelia burgdorferi erpT expression in the arthropod vector and murine host, Mol. Microbiol., 31 (1999) 281290. Szczepanski,A., Furie,M.B., Benach,J.L., Lane,B.P., Fleit,H.B., The interaction between Borrelia burgdorferi and endothelium in vitro., J. Clin. Invest., 85 (1990) 1637-1647. Lux,R., Moter,A., Shi,W., Chemotaxis in pathogenic spirochetes: directed movement toward targeting tissues?, J Mol. Microbiol. Biotechnol., 2 (2000) 355-364. Shih,C.M., Chao,L.L., Yu,C.P., Chemotactic migration of the Lyme disease spirochete (Borrelia burgdorferi) to salivary gland extracts of vector ticks, Am. J. Trop. Med. Hyg., 66 (2002) 616-621. Berg,H.C., The Rotary Motor of Bacterial Flagella, Annu. Rev. Biochem., 72 (2003) 19-54. Szurmant,H., Ordal,G.W., Diversity in chemotaxis mechanisms among the bacteria and archaea, Microbiol. Mol. Biol. Rev., 68 (2004) 301-319. Wadhams,G.H., Armitage,J.P., Making sense of it all: bacterial chemotaxis, Nat. Rev. Mol. Cell Biol., 5 (2004) 1024-1037. Sourjik,V., Receptor clustering and signal processing in E. coli chemotaxis, Trends Microbiol., 12 (2004) 569-576. Parkinson,J.S., Complementation analysis and deletion mapping of Escherichia coli mutants defective in chemotaxis, J. Bacteriol., 135 (1978) 45-53. Bakker, R.G., Measuring chemotaxis in Borrelia burgdorferi the Lyme disease spirochete. Ph.D. Dissertation, West Virginia Univ. 2006. Shapiro,L., Losick,R., Dynamic spatial regulation in the bacterial cell, Cell, 100 (2000) 89-98. Lybarger,S.R., Maddock,J.R., Polarity in action: asymmetric protein localization in bacteria, J. Bacteriol., 183 (2001) 3261-3267. Shapiro,L., McAdams,H.H., Losick,R., Generating and exploiting polarity in bacteria, Science, 298 (2002) 1942-1946. Motaleb,M.A., Miller,M.R., Li,C., Bakker,R.G., Goldstein,S.F., Silversmith,R.E., Bourret,R.B., Charon,N.W., CheX is a phosphorylated CheY phosphatase essential for Borrelia burgdorferi chemotaxis, J. Bacteriol., 187 (2005) 7963-7969. Boesch,K.C., Silversmith,R.E., Bourret,R.B., Isolation and characterization of nonchemotactic CheZ mutants of Escherichia coli, J. Bacteriol., 182 (2000) 3544-3552.

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Targeted and Random Mutagenesis in Leptospira biflexa: Application for the Functional Analysis of Iron Transporters Hélène LOUVEL a, Simona BOMMEZZADRI a, Nora ZIDANE b, Paula RISTOW a, Zoé ROUY c, Claudine MEDIGUE c, Caroline BOURSAUX-EUDE d, Isabelle SAINT GIRONS a, Christiane BOUCHIER b, and Mathieu PICARDEAU a,1 a Laboratoire des Spirochètes, Institut Pasteur, Paris, France b Plate-Forme Génomique (PF1), Institut Pasteur, Paris ;France c Pasteur Genopole£ Ile de France, Genoscope and CNRS-UMR8030, Atelier de Génomique Comparative, Evry, France. d Plate-forme Intégration et Analyse Génomiques (PF4), Institut Pasteur, Paris, France Abstract. The genus Leptospira belongs to the order Spirochaetales and is composed of both saprophytic and pathogenic members, such as Leptospira biflexa and L. interrogans, respectively. A major factor contributing to our ignorance of spirochetal biology is the lack of methods for genetic analysis of these organisms. We have developed a system for transposon mutagenesis of L. biflexa using a mariner transposon, Himar1. This mutagenesis approach yields a randomly distributed set of insertion mutations throughout the genome, which can be screened for specific phenotypes. An analysis of transposon mutants has allowed the identification of genes required for diverse biological activities such as amino acid biosynthesis and metal transport. The development of numerous genetic tools for saprophytic species of Leptospira enables the use of these bacteria such as L. biflexa as a model bacterium We therefore sequenced the genome of the saprophyte L. biflexa consisting of a 3.6-megabase large chromosome and a 278kilobase small chromosome. Comparative genomics, in combination with gene inactivation, give us significant functional information on iron homeostasis in Leptospira. Keywords. Allelic exchange, transposon, auxotrophs, iron, manganese

Introduction The genetics of Leptospira remains at a very early stage compared to that of other bacterial species. The lack of genetic tools in pathogenic leptospira does not allow the 1

Corresponding author: Mathieu Picardeau. Laboratoire des Spirochètes, Institut Pasteur, 28 rue du docteur Roux, 75724 Paris Cedex 15, France. Tel: 33 (1) 45 68 83 68. Fax: 33 (1) 40 61 30 01. E-mail: [email protected].

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full characterization of genes of interest. In contrast, numerous tools for genetic manipulation of saprophytic Leptospira have been developped in our laboratory, enabling the use of these bacteria such as L. biflexa as a model bacteria. For example, we recently developed a mutagenesis system that used the transposable element Himar1, a member of the mariner family of transposons [1, 7]. Transposons of the mariner family have been used successfully for random mutagenesis of a diverse range of organisms, including eukaryotes, archaea, and both gram-positive and gram-negative bacteria. Unlike many eubacterial transposons, mariner elements can move from one DNA molecule to another by a ‘cut and paste’ mechanism dependent solely on transposase activity [10], without the requirement of host accessory factors. Therefore, we tested whether Himar1 elements can transpose in the saprophyte L. biflexa, and herein we demonstrate that to be the case. Iron plays a central role in many major biological processes such as the electron transport chains for most living cells, including Leptospira spp. In gram-negative bacteria, iron sources can be recognized by specific outer membrane receptors, called TonB-dependent receptors and then transported across the inner membrane by the FeoAB system or periplasmic-binding–protein-dependent ABC permeases. We recently characterized the fecA- and feoB-like genes, which are both involved in iron acquisition, by random transposon mutagenesis in L. biflexa [7]. Genome analysis of L. biflexa allows the identification of several genes that could be involved in iron uptake systems. To study the function of these genes, we generated several targeted mutants in L. biflexa, and their phenotypes were characterized. Since pathogenic Leptospira spp., as well as saprophytic species, need to obtain iron to grow in vitro, and probably in vivo in the host, a better knowledge of the iron uptake systems and their regulation is essential to understand the pathogenesis of this intriguing group of organisms.

1. The L. biflexa Sequencing Project Recently, the complete genome sequences of L. interrogans serovar Lai [9] and L. interrogans serovar Copenhageni [8] have been achieved. The two L. interrogans serovars contain a 4.3-Mb large circular chromosome and a 350-kb small circular chromosome with an average G+C content of approximately 35%. Although the two genomes exhibit 95% identity at the nucleotide level (99% identity for predicted protein coding genes that are orthologs), L. interrogans serovar Lai has nearly 1000 extra genes (4727 ORFs versus 3667 ORFs) in comparison to L. interrogans serovar Copenhageni [8]. This discrepancy may not reflect the reality, but rather be due to the annotation criteria used by the two genome projects [12]. The syntaxic re-annotation of the two genomes by using the MICheck software [4] estimated the number of ORFs to 3798 for L. interrogans serovar Lai and to 3651 for L. interrogans serovar Copenhageni. Whole genome comparative analyses revealed that the L. interrogans genomes are essentially syntenic, with the major of genomic difference consisting of one megabase inversion in the large chromosome [8]. The draft genome sequence (> 8x coverage) of the saprophyte L. biflexa serovar Patoc strain Patoc1 consists of a 3.6megabase large chromosome and a 278-kilobase small chromosome with a total of 3801 predicted coding genes, 60% of which are conserved with those of L. interrogans.

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H. Louvel et al. / Targeted and Random Mutagenesis in Leptospira biflexa

2. Targeted Mutagenesis in L. biflexa In this work, the ability of Leptospira to use various iron sources for its physiological role requirement was studied. We show that Leptospira spp. can acquire iron from different sources, including the siderophores aerobactin and ferrichrome. However, the uptake systems of these molecules remain to be determined. The outer membrane of Leptospira is relatively impermeable [5], and the acquisition of molecules may mainly rely on the active transport through dedicated outer membrane receptors. In gramnegative bacteria, the transport of iron sources into the periplasm against their concentration gradients is mediated by the inner membrane complex TonB, ExbB, and ExbD. The energy is transduced to the outer membrane receptor, also called TonBdependent receptor. Analysis of the genomes indicated that L. biflexa and L. interrogans contain eight and 12 genes, respectively, whose products share homology with TonB-dependent receptors. However, substrate specificity is difficult to characterize based solely on sequence analysis of the receptors. By random transposon mutagenesis in L. biflexa, we recently identified mutants (see below) with insertions in a gene (LEBIa1620) encoding a protein that shares homology with FecA [7]. Besides FecA, we have attempted to disrupt the other putative genes encoding TonB-dependent receptors by allelic exchange in L. biflexa. Gene inactivation of LEBIa2634, LEBIa2704, and LEBIa2712 resulted in a wild-type phenotype in iron-depleted medium supplemented with different iron sources. The lack of a phenotype in these mutants could be due to functional redundancy with another iron uptake system. Disruption of LEBIa3308 resulted in a mutant that was impaired in its ability to use desferrioxamine as an iron source. Interestingly, the mutant was also impaired in its ability to utilize ferrichrome, while the transport of aerobactin, another hydroxamate siderophore, was unaffected. These results are evidence that LEBIa3308 encodes the receptor protein for ferrioxamines and ferrichrome in L. biflexa. Finally, we failed to obtain double crossover events in LEBIa0145, LEBIa2087, and LEBIa3054. This may indicate that these genes are essential for the survival of L. biflexa. Leptospira have an absolute requirement for vitamin B12, which is usually transported via TonB-like systems in other gram-negative bacteria. Since vitamin B12 is a co-factor for enzymes of major biological processes, inactivation of its receptor should result in non viable mutants. Amino acid comparisons of TonB-dependent receptors of heme, hemoglobin, siderophores, and vitamin B12 revealed a highly conserved domain containing the FRAP and NPNL amino acid box [2, 11]. This conserved domain was found in some TonB-dependent receptors from Leptospira, including LEBIa2087 and LEBIa0145.

3. L. biflexa Mutants Isolated by Random Insertional Mutagenesis An important approach for investigating metabolism processes is the generation of large numbers of mutant bacteria. Electroporation of L. biflexa with plasmid vector pSC189 [3] containing both the hyperactive transposase C9 [6] and transposon terminal inverted repeats flanking a kanamycin resistance gene resulted to approximately 5000 transformants per Pg of DNA. To improve expression of the Himar1 transposase, the hyperactive transposase C9 was fused to spirochetal promoters. L. biflexa was electroporated with plasmids pSHT and pSFLT expressing the mariner transposase from the L. interrogans hsp10 and flgB promoters, respectively. We found that the transposition of Himar1 from donor plasmids pSHT and pSFLT was about 5-fold more

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efficient than that of Himar1 from pSC189, i.e., 2 to 3.5 104 transformants per Pg of DNA were obtained with plasmids pSHT and pSFLT. Southern blot hybridization and sequencing of the Himar1 insertion sites of 50 randomly chosen kanamycin-resistant colonies showed the transposition to be random and stable [7]. Similar to other hosts, the mariner transposon targets the dinucleotide sequence TA, which is duplicated upon cut-and-paste insertion. This target site is abundant in the L. biflexa genome with an average G+C content of approximately 39%. The availability of the whole genome sequence of L. biflexa should greatly facilitate genetic analyses in Leptospira. For example, genes disrupted by transposon insertion mutagenesis can be rapidly identified in mutants with interesting phenotypes through sequence analysis of the flanking regions and comparison with genome sequence. To investigate iron transporters, 6000 L. biflexa transposon mutants were screened onto medium with and without hemin, thus allowing the identification of 15 hemin-requiring mutants, and the putative genes responsible for this phenotype were identified. Twelve mutants had distinct insertions in a gene encoding a protein that shares homology with TonB-dependent receptor FecA, involved in ferric citrate transport. The L. biflexa fecA mutants were impaired in their ability to use iron citrate [7], and also iron sulfate and aerobactin as an iron source. Interestingly, among hydroxamate siderophores, aerobactin forms a distinct subfamily of siderophores that is derived from citrate, and therefore aerobactin and iron citrate share a similar structure that could be recognized by the same receptor. We also identified two mutants with a Himar1 insertion into feoAB-like genes, the product of which is required for ferrous iron uptake in many bacterial organisms [7]. Interestingly, another mutant exhibited a Himar1 insertion into a two-component system that could be involved in the regulation of iron uptake. Finally, by screening for manganese-requiring mutants, two new genes of unknow function, including a putative manganese transporter, were also identified. Phenotypic characterization of these mutants extends our understanding of the biology of Leptospira spp., which remains largely unknown.

4. Conclusions The availability of the whole-genome sequence of L. biflexa should shed light on the evolution of genomes in Leptospira and reveal ways in which virulent pathogens can evolve. The genome content reflects the bacterial lifestyles, and results from the distinct ongoing process of genome optimization in saprophytes and pathogens. The possession of specialized iron transport systems for the saprophyte L. biflexa and the pathogen L. interrogans may thus reflect the various iron sources they may encounter in their diverse habitats. Based on our findings, a model for iron uptake in Leptospira can reasonably be proposed (Figure 1). The analysis of the genome of the pathogen L. interrogans has allowed the identification of 12 putative TonB-dependent receptors, while L. biflexa possesses eight putative TonB-dependent receptors. This difference suggests that pathogenic species are able to use a wider panel of iron sources, in comparison to L. biflexa. The pathogens may also present redundancy in their genome content. Like in gram-negative bacteria, periplasmic binding proteins may shuttle ironcontaining complexes from TonB-dependent receptor to cytoplasmic membrane ATPbinding cassette (ABC) transporters, that in turn deliver them in the cytoplasm. The pathogen L. interrogans may obtain iron from its association with heme and thus

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Red blood cells

3308

?

2634

?

2712

?

3054

2087

0145

Outer mb

?

2704

?

Heme proteins

Desferrioxamine Ferrichrome

Iron sources

Iron citrate Aerobactin Iron salts

1620

Vitamin B12

Hemolysins

TonB-dependent recept

Siderophores RTX toxin transporter

Inner mb ExbB ExbD TonB

Hemin uptake system (HmuSTUV)

Siderophore typeABC transporter (?)

Metal type- ABC transporter

FeoAB

Figure 1. Schematic representation of iron acquisition systems in L. biflexa. Analysis of the draft genome sequence of the saprophyte L. biflexa suggests the presence of eight putative TonB-dependent receptors. The L. biflexa genome contains five TonB loci that could be involved in the formation of the ExbB-ExbD-TonB complex (only one TonB system is represented in this figure). By mutagenesis in L. biflexa, we isolated and characterized the function of two TonB-dependent receptors (LEBIa3308 and LEBIa1620) and the FeoAB system. Failure to obtain knock-out mutants and genetic organization suggest that LEBIa0145 is the TonBdependent receptor of either vitaminB12 or hemin. A metal type-ABC transporter was found in the L. biflexa genome and the HmuSTUV transport proteins may be involved in the periplasmic transport of hemin. An undescribed system may also exist for the periplasmic transport of siderophores (indicated in italic text). Leptospira could also release heme and hemoglobin from host red blood cells by the secretion of hemolysins. There is no evidence of siderophore synthesis by Leptospira spp. The molecules participating in each step of the transport process have not been identified and may involved reductases, periplasmic proteins, and permeases. In the cytoplasm, iron can be stored in bacterioferritin and Dps proteins.

secretes hemolysins to lyse red blood cells and liberates this metal. Surprisingly, L biflexa, a non pathogenic species, has putative hemin uptake and hemolysin secretion systems and may use a similar mechanism, a property hitherto described exclusively for pathogenic bacteria. However, the L. biflexa genome does not contain orthologs to the L. interrogans sphingomyelinase hemolysins, which may be involved in the typical vascular damage seen in the acute disease. It is rather questionable whether hemin is available to saprophytes during their living in the environment. The role of the genes of L. biflexa that encoded hemolysins and a putative hemolysin secretion system in iron acquisition remains to be elucidated. Leptospira also possess uptake systems that use siderophores produced by other bacteria or fungi. Bacterial iron homeostasis is best understood in E. coli, a bacterium phylogenetically distant from Leptospira. This study will serve as a basis for further study on iron acquisition systems in Leptospira. The application of our random transposon mutagenesis system to saprophytic and pathogenic strains will be particularly useful for discovering new genes involved in iron uptake and regulation.

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Antibiotic Resistance in Borrelia burgdorferi: Applications for Genetic Manipulation and Implications for Evolution D. Scott SAMUELS 1 Division of Biological Sciences and Biomolecular Structure and Dynamics Program, The University of Montana, Missoula, Montana, USA Abstract. Antibiotic resistance is not a clinical problem with Lyme disease, but it has been extensively employed to genetically dissect the causative agent Borrelia burgdorferi. The first selectable marker was a coumermycin A1-resistant gyrB allele, which encodes a subunit of DNA gyrase, a target of several antibiotics. The utility of coumermycin A1 resistance has been compromised by technical and genetic barriers; resistance to other antibiotics has replaced the gyrB marker. Fluoroquinolones are another class of antibiotics that target DNA gyrase, as well as its homolog topoisomerase IV. Fluoroquinolone resistance in B. burgdorferi maps to parC, which encodes a subunit of topoisomerase IV, suggesting that this enzyme is the primary target of fluoroquinolones in Borrelia. A fluoroquinoloneresistant parC allele has been fashioned into a counter-selectable marker, a genetic tool used to select for the loss of DNA. One of the second-generation selectable markers is a heterologous aadA gene that confers resistance to spectinomycin and streptomycin, which target the small subunit of the ribosome. Selection with spectinomycin failed due to a high frequency of mutants in the population. These had mutations in 16S rRNA and were able to compete with wild type in vitro. This lack of a significant fitness cost for the mutant may contribute to the spread of antibiotic resistance. Keywords. Antibiotic resistance, Borrelia burgdorferi, Lyme disease, genetics, mutants

Introduction Antibiotic resistance in clinical isolates is a colossal public health concern [1í3]. Fortunately, there have been no reported cases of antibiotic resistance in Lyme disease patients. However, antibiotic resistance has been used as a selectable marker for molecular genetic studies of Borrelia burgdorferi and, because of technical limitations, is likely the only viable genetic marker that can be used for selection. Rosa, Tilly, and

1 Corresponding author: Division of Biological Sciences, The University of Montana, 32 Campus Dr., Missoula, MT 59812-4824, USA: E-mail: [email protected].

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Stewart [4] is recommended as a recent and fairly comprehensive review of molecular genetics in B. burgdorferi. Antibiotics are broadly defined as organic substances that are either toxic or growth-inhibitory for organisms. Infectious disease therapy depends on the selective toxicity of antibiotics for the pathogen, so antimicrobial agents target a structure or function only found in a specific organism or group of organisms. For antibiotics that are specific for bacteria, these targets include cell wall synthesis (beta-lactams, glycopeptides, and cyclic lipopeptides), the cell membrane (polymyxins), protein synthesis (aminoglycosides, and the closely related spectinomycin, tetracyclines, macrolides, streptogramins, ketolides, and lincosamides), folate synthesis (sulfonamides and trimethoprim), RNA synthesis (rifamycins), DNA synthesis (fluoroquinolones and coumarins), and others [3, 5]. Many infectious diseases have been vanquished by these and other antibiotics, which have significantly extended and improved the lives of an enormous number of people. Unfortunately, resistance to these antibiotics is proving to be a “new apocalypse” [1]: methicillin-resistant Staphylococcus aureus, vancomycin-resistant S. aureus, vancomycin-resistant enterococci, macrolide-resistant Streptococcus, penicillin-resistant pneumococci, and multidrug-resistant tuberculosis [1í3, 6]. There are several causes for the escalation in antibiotic resistance, but the overuse of antibiotics, especially in agriculture, which accounts for well over half of the antibiotic use in the United States, is likely the primary factor [7]. A recent study found that the majority of soil bacteria sampled are resistant to many types of antibiotics: most bacterial strains tested were resistant to about seven different antibiotics, and not a single antibiotic had efficacy against all the bacterial strains [8]. Hence, antibiotic resistance is pervasive in the environment and will likely spread because of the promiscuity of bacteria sharing their genes [6, 9í12]. There are several mechanisms of antibiotic resistance: the antibiotic can be either kept out or pumped out of the cell, the antibiotic can be either destroyed or modified, or the target of the antibiotic can be changed. The mechanisms discussed in this chapter will focus on target mutation [5]. The mutated targets in antibiotic-resistant B. burgdorferi include DNA gyrase [13], its cousin topoisomerase IV [14], and the ribosome [15]. Two of these mutants have been developed into molecular tools for genetic experiments in the laboratory [16, 17], and the third has proved to be a hindrance [18].

1. History of Coumermycin A1 Resistance and its Application as the First Selectable Marker We isolated the first genetically defined antibiotic-resistant mutants of B. burgdorferi in 1993 [13]. Background (or “spontaneous”) mutants (as opposed to those induced with a mutagen) were selected with the coumarin antibiotic coumermycin A1, which targets the B subunit of DNA gyrase, a type II topoisomerase in prokaryotes [19, 20]. Type II topoisomerases are enzymes that alter DNA topology by breaking and resealing both strands of the double helix. DNA gyrase is a tetramer comprising two A subunits (GyrA) and two B subunits (GyrB). The A subunit is involved in the double-stranded nicking and resealing reactions, while the B subunit is responsible for providing energy through ATP hydrolysis. The gyrB and gyrA genes are in an operon proximal to the replication origin [21] near the center of the linear chromosome in B. burgdorferi [16, 22í24].

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D.S. Samuels / Antibiotic Resistance in Borrelia burgdorferi Thr-165 Arg-136 Gly-77

ATP

Asn-107 Asp-105

Figure 1. A region of the N-terminal domain of GyrB from Escherichia coli with a non-hydrolyzable ATP analog and residues homologous to those altered in coumarin-resistant mutants highlighted in black: Gly-77, Asp-105, Asn-107, Arg-136, and Thr-165 are homologous to Gly-74, Asn-102, Gly-104, Arg-133, and Thr162 in B. burgdorferi. Coordinates are courtesy of Dale Wigley [26].

DNA gyrase has proved to be a popular drug target because of its essential nature as well as structural and functional differences with the eukaryotic type II topoisomerases that were exploited. The clinically effective fluoroquinolone antibiotics target the A subunits of DNA gyrase and its homolog topoisomerase IV (see section 2 below). On the other hand, coumermycin A1 and the other coumarin antibiotics (such as novobiocin) never achieved widespread clinical application because of several pharmacological limitations, such as solubility and toxicity [19, 20]. We mapped the coumermycin A1-resistant mutations to a conserved arginine residue in GyrB [13]. This residue (Arg-133 in B. burgdorferi is homologous to Arg-136 in Escherichia coli) makes contact with coumarin antibiotics [25], which interact with GyrB at the ATPbinding site (Figure 1), and is mutated in several other coumarin-resistant prokaryotes (Table 1) [27í34]. Note that this arginine is numbered 133 in our publications (GenBank accession AF017075) [13, 16], but corresponds to Arg-138 in the B. burgdorferi genome sequence (BB0436) [22]. This discrepancy is because our collaborator Wai Mun Huang observed that there is no reasonable Shine-Dalgarno sequence for a ribosome binding site except downstream of the start codon assigned during genome annotation. 1.1. gyrB Mutants Resistant to Coumermycin A1 Arg-133 was mutated to either isoleucine or glycine in coumermycin A1-resistant B. burgdorferi; the mutants had a mild growth phenotype [13]. Curiously, the homologous arginine (or lysine in some bacterial species) is mutated to serine, cysteine, histidine, leucine, isoleucine, glutamine, glutamic acid, and threonine in other coumarin-resistant microorganisms (Table 1) [27í34]. (Three of the nine species in which coumarinresistant mutations have been isolated are spirochetes!) The particular mutations depended on which of the six arginine codons was present in a particular species: the AGA of B. burgdorferi gyrB is mutated to ATA or GGA in background mutants [13], although mutations to AGU or AGC (serine), AAA (lysine), and ACA (threonine) would only require a single base change (and, as we discovered, do confer resistance to coumermycin A1 [35]). These observations provoked the question of which amino acid

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substitutions would confer resistance. We used site-directed mutagenesis of this codon to demonstrate that any substitution conferred resistance: the 19 single Arg-133 mutants conferred between sixfold and 560-fold resistance to coumermycin A1 [35].

Table 1. GyrB mutations conferring coumarin resistance. Organism

Mutation (conserved Arg or Lys in bold)

Reference

Escherichia coli

Arg-136 to Ser, Cys, His, Leu, or Ala; Asp-73 to Glu; Gly-77 to Ala or Ser; Ile-78 to Ala or Leu; Gly-164 to Val; or Thr-165 to Ala or Val

[27, 28, 36]

Haloferax alicantei

Arg-137 to His, Asp-82 to Gly, Ser-122 to Thr (triple mutant)

[29]

Streptococcus pneumoniae

Ser-127 to Leu

[37]

Staphylococcus aureus

Arg-144 to Ser or Ile; Gly-85 to Ser; Ile-102 to Ser; Ser-128 to Leu; Thr-173 to Asn or Ala; or Ile-175 to Thr; or double mutants, including with Ile-56 to Ser

Bartonella bacilliformis

Arg-184 to Gln; Gly-124 to Ser; or Thr-214 to Ala or Ile

[30]

Treponema denticola

Lys-136 to Glu or Thr

[31]

Brachyspira hyodysenteriae

Gly-78 to Ser or Cys; or Thr-166 to Ala

[38]

Thermoplasma acidophilum

Arg-136 to His

[33]

Borrelia burgdorferi

Arg-133 to Gly or Ile, or any other substitution; or double and triple mutants, including with Asn-102 to Asp and/or Gly-104 to Asp; or other mutations at Gly-74 or Thr-162

[32, 34]

[13, 35]

We next hypothesized that a second-site mutation would increase resistance. To test this hypothesis, we plated Arg-133 to Ile mutants, which were marked with a second silent mutation in the Arg-133 to Ile codon (ATT instead of the background single ATA mutation), in higher levels of coumermycin A1. Two types of second-site mutations were isolated: either Asn-102 to Asp or Gly-104 to Asp. These residues are opposite Arg-133 on a lid covering the ATP-binding site (Figure 1) [26], but move distal to the protein when a coumarin binds [25]. Both mutants yielded higher levels of resistance, and the experiment yielded another question: would all three mutations confer even greater resistance? The advantage of a “super-resistant” allele for developing coumermycin A1-resistant gyrB into a selectable marker would be that one could use very high levels of coumermycin A1 to minimize the number of background mutants in genetic experiments. The additional motivation was that more mutations might decrease homologous recombination of the marker into the chromosomal gyrB locus. Therefore, one of the second-site mutations was transformed into the other double mutant. The resulting triple mutant did in fact have even higher levels of resistance. Note that the colloquial description of this triple mutant allele, “NGR,” is derived from the one letter codes for the three amino acids that were mutated

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(N102D/G104D/R133I) and became a shortcut when writing the mutations on a multitude of culture tubes every week. 1.2. Development and Downfall of gyrB as a Selectable Marker The final step in fashioning the triple mutant gyrB allele into a selectable marker for genetic studies was to map the transcriptional start site for gyrB (and presumably the gyrBA operon [24]) in collaboration with Rich Marconi [39]. The gyrB cassette was amplified as the upstream regulatory region, and the entire open reading frame with the appropriate restriction enzymes incorporated into the primers. These coumermycin A1resistant gyrB cassettes were used for many years as the only selectable marker available. Despite problems and inconveniences, the coumermycin A1-resistant gyrB was seminally employed to demonstrate genetic transformation [16] and recombination of DNA into a heterologous site [40]. In addition, coumermycin A1 resistance was used to show that short oligonucleotides could serve as transformation substrates for bacteria [41]. Besides for illustrating the feasibility of reverse genetic experiments, several genes were disrupted with gyrB to probe their in vitro function, including ospC [42], guaB [43], oppAIV [44], gac [45], rpoS [46], and chbC [47]. The problems of the gyrB marker include extensive screening of transformants, which is due to homologous recombination into the chromosomal gyrB locus [40, 45], the large size, which we propose prevented its use in demonstrating transduction despite exhaustive efforts by former graduate student Christian Eggers [48], and pleiotropic effects, which are due to perturbed levels of DNA supercoiling. Regarding the latter issue, gyrB mutants, which have relaxed DNA supercoiling [27], have increased expression of groEL [49] and ospC [50]. Furthermore, our preliminary data suggest that the coumermycin A1-resistant gyrB allele suppressed the phenotype of at least one mutant in our laboratory. We had discovered Gac, an architectural DNA-binding protein in B. burgdorferi [24] and were curious about its function. The gac gene is uniquely embedded in the gyrA gene, which encodes the A subunit of DNA gyrase, so generating a null gac mutant without disrupting the essential gyrA gene was a challenge. The gyrB marker was used to mutate the first two methionine codons of gac, resulting in abolition of Gac synthesis without dramatically affecting GyrA [45]. Unfortunately, the gac mutants had no recognizable phenotype. However, more recent efforts using a kanamycin resistance cassette (see below) resulted in gac mutants with a growth phenotype that was readily suppressed during outgrowth of the cultures. We are currently introducing the gac mutations linked to kanamycin resistance into coumermycin A1-resistant cells to test our hypothesis that the gyrB mutation suppresses the gac mutant phenotype (and we are creating a conditional gac mutant using an inducible promoter). Gac has HU-like activity [24] and there is precedence for suppressing an HU mutant defect with a coumarin-resistant gyrB allele [51, 52]. Other experiments have revealed genetic suppression of cell division gene defects by gyrB mutations [53, 54]. 1.3. Further Developments in Genetic Manipulation A major breakthrough was when Jim Bono, a postdoctoral fellow in Patti Rosa’s laboratory, fused a strong constitutive B. burgdorferi promoter to an exogenous gene, aphI, which confers resistance to the aminoglycoside kanamycin [55]. Concurrently, Felipe Cabello’s laboratory discovered that the ermC gene conferred erythromycin

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resistance to B. burgdorferi, and they fused a native promoter to a gfp allele [56] (although that allele does not function well in B. burgdorferi [57]). The new antibiotic resistance markers shattered the barriers to many genetic experiments (see [4] for review) and the promoter fusion method, first introduced to B. burgdorferi by Chuck Sohaskey when he was in Alan Barbour’s laboratory [58], has been wielded several times since [17, 18, 57, 59í64]. A caveat regarding the ermC marker is that some Institutional Biosafety Committees have balked at its use because erythromycin can be prescribed to treat Lyme disease in certain situations, although the discovery of natural erythromycin-resistant mutants should allay these concerns [65]. However, two other hybrid selectable markers, conferring resistance to streptomycin [18] and gentamicin [59], are now popular with molecular borreliologists. The other vital advance was the construction [57, 66], or identification [56], of shuttle vectors that replicate in B. burgdorferi. Currently, there are a suite of plasmids available for genetic studies: these are based on either pGK12, which is derived from the Lactococcus lactis plasmid pWV01 [56], pBSV2, which is derived from cp9 [66], or pCE320, which is derived from a cp32 [57]. Some of these plasmids are compatible during replication in B. burgdorferi [18]. More recently, several next-generation genetic tools have been developed: transposon mutagenesis [62, 63], inducible promoters [64, 67, 68], and a counter-selectable marker [17].

2. Fluoroquinolone Resistance and its Application as a Counter-selectable Marker Many, but not all, bacteria have two type II topoisomerases: DNA gyrase and topoisomerase IV. DNA gyrase introduces negative supercoiling into DNA, and topoisomerase IV decatenates replicated DNA as well as relaxes supercoiled DNA [69]. Topoisomerase IV is also a tetramer comprised of two A subunits, called ParC, and two B subunits, called ParE (for their role in partitioning of DNA to daughter cells). Fluoroquinolones, potent antibiotics that are widely used, target type II topoisomerases [20, 70]. Resistance to fluoroquinolones usually maps to a small region in the A subunits of DNA gyrase (GyrA) and topoisomerase IV (ParC). The primary target of fluoroquinolones, defined as the type II topoisomerase that is mutated in firststep fluoroquinolone-resistant strains, is usually DNA gyrase in gram-negative bacteria and topoisomerase IV in gram-positive bacteria, although this also depends on the particular fluoroquinolone [20, 70]. To our knowledge, no data had ever been reported on the primary target in spirochetes. Therefore, we used a genetic approach to identify the antibiotic target by selecting for mutants of B. burgdorferi that are resistant to various fluoroquinolone antibiotics [14]. 2.1. Fluoroquinolone Mechanism of Action How fluoroquinolones work is not completely understood [71, 72]. DNA topoisomerases catalyze changes in DNA topology by a nucleophilic attack with their active site tyrosine on the phosphodiester bond between nucleotides. A transesterification results in one or two covalent bonds between one or both strands and the topoisomerase. Type II topoisomerases, DNA gyrase, and topoisomerase IV attack both strands of the DNA substrate. These enzyme-linked DNA strands serve as a gate to pass other DNA strands resulting in the eponymous changes in DNA topology. The topoisomerase mechanism is completed by a second transesterification that restores the

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integrity of the DNA. The fluoroquinolone antibiotics are thought to stabilize the covalent complex between the type II topoisomerase and DNA. These result in blockage of transcribing RNA polymerases and replicating DNA polymerases as well as in double-stranded breaks. Therefore, these antibiotics are thought to turn topoisomerases into poisons. An antibiotic-sensitive enzyme in the cell is postulated to be sufficient to cause DNA damage and cell death [71, 72]. Consequently, resistant mutations in gyrA are recessive [73]. The story with topoisomerase IV and parC mutations is not as straightforward. Results in E. coli have demonstrated codominance and gene dosage effects [74, 75]. However, parC is not the primary target of the fluoroquinolones in E. coli, although experiments in Staphylococcus aureus, where topoisomerase IV is the primary target, demonstrated fluoroquinolone-resistant parC was dominant [76]. A caveat to this latter report is that parC is the second gene in an operon with parE, and fluoroquinolone-resistant parEC was recessive, suggesting that expression of parC alone may be attenuated and result in lack of genetic interaction. 2.2. parC Mutants Resistant to Fluoroquinolones Mutants are difficult to obtain because fluoroquinolone resistance is recessive (which is one reason that fluoroquinolones are clinically successful chemotherapeutic agents). We isolated several fluoroquinolone-resistant mutants of B. burgdorferi: all of them had one of five different mutations (at three codons) in parC, the gene encoding the A subunit of topoisomerase IV, and none had mutations in gyrA [14]. This provided strong support that topoisomerase IV is the primary target of fluoroquinolone antibiotics in B. burgdorferi. The mutations were Thr-69 to Lys or Arg, Ser-70 to Pro, and Glu-73 to Gly or Lys. Thr-69 and Glu-73 are highly conserved residues that are hot spots for fluoroquinolone-resistant mutations in other bacteria (although the homolog of Thr-69 is a Ser in almost all other bacteria). The parC mutants, like the gyrB mutants, had a mild growth phenotype; they were between fourfold and 75-fold more resistant than wild-type, depending on the particular mutation and fluoroquinolone tested, with the exception that none of the mutants were resistant to the fluoroquinolone ciprofloxacin [14]. In addition, we were not able to isolate ciprofloxacin-resistant mutants. We do not know why the mutants are susceptible to ciprofloxacin (class II), but resistant to the more the potent fluoroquinolones sparfloxacin (class III), moxifloxicin (class IV), and Bay-Y3118 (experimental) [77].

Figure 2. Alignment of a portion of the ParC and GyrA proteins from B. burgdorferi (Bb), Streptococcus pneumoniae (Sp), and E. coli (Ec). The numbers at the top refer to the amino acid residues of the ParC protein of B. burgdorferi and indicate residues that are mutated in the fluoroquinolone-resistant strains (Thr69, Ser-70, and Glu-73). The asterisk (*) indicates the highly conserved serine that confers fluoroquinolone resistance when mutated; the plus (+) indicates the conserved acidic residue that is also often mutated. This figure is reprinted from Galbraith et al., Antimicrob. Agents Chemother. (2005) [14].

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Threonine, at position 69 in ParC from B. burgdorferi, is a conservative substitution for serine, but the homologous residue in GyrA is Gln-86 (Figure 2). Wildtype B. burgdorferi are less susceptible to fluoroquinolones than most other bacteria [77, 78]. We hypothesize that the presence of a glutamine, instead of the conserved serine that is often mutated in fluoroquinolone-resistant strains, is responsible for both the natural lack of fluoroquinolone susceptibility and the primary target preference for topoisomerase IV [14]. 2.3. Development of parC as a Counter-Selectable Marker Recent advances (described above), including new selectable markers for introducing recombinant DNA, have allowed researchers to exploit the awesome power of genetics to study B. burgdorferi. However, another essential component of a genetic system, besides useful selectable markers, is a counter-selectable marker. This allows us to select for the loss of genetic material from the experimental organism. A counterselectable marker has been developed for Leptospira [79]. One strategy for counterselection involves using a merodiploid with a recessive antibiotic-resistance allele on the chromosome and a dominant antibiotic-sensitive allele on a plasmid. The merodiploid is susceptible to the antibiotic, unless the plasmid is lost. Therefore, plasmid loss is selected with the antibiotic. Although the story is complicated, we hypothesized that fluoroquinolone resistance in B. burgdorferi is recessive, because mutants were not readily isolated compared to other antibiotics such as coumermycin A1, spectinomycin, and the aminoglycosides [13, 15]. Therefore, we constructed a counter-selectable marker for B. burgdorferi using parC alleles. In Borrelia, parC appears to be in an operon with parE and lack its own promoter [22]. We have previously used the gyrBA promoter to drive transcription of a selectable marker (see above). Therefore, we fused the gyrBA promoter to the parC open reading frame. This hybrid fluoroquinolone-sensitive cassette was cloned into the shuttle vectors pBSV2 [66] and pKFSS1 [18], generating the plasmids pBSCSM and pKFCSM that confer resistance to kanamycin and streptomycin, respectively, and susceptibility to fluoroquinolones [17]. As a proof of principle, pBSCSM was transformed into the Glu-73 to Lys parC mutant that is resistant to several fluoroquinolone antibiotics [14] and the presence of the plasmid was selected for with kanamycin. The transformants were then plated with or without a fluoroquinolone antibiotic (Bay-Y3118) and with or without kanamycin. Kanamycin selects for the presence of the plasmid, and Bay-Y3118 selects against the presence of the plasmid. Ten colonies were picked from the kanamycin plate and ten from the Bay-Y3118 plate; no colonies were obtained in the presence of both kanamycin and Bay-Y3118. Plasmid maintenance was assayed by xenodiagnosis in E. coli and by PCR by using plasmid-specific primers. All ten colonies from the kanamycin plate possessed the plasmid and none of the ten colonies assayed from the Bay-Y3118 plate demonstrated evidence of the plasmid [17]. As controls, the parental plasmid pBSV2, which lacks gyrBAp::parC, does not confer fluoroquinolone susceptibility to the parC mutant, and pBSCSM cannot be selected against in the parental wild-type strain. pKFCSM has not yet been assayed, but its parental plasmid pKFSS1, which confers resistance to spectinomycin and streptomycin, is less stable than pBSV2 [18], leading us to hypothesize that it may be more readily cured upon counter-selection with fluoroquinolone. These preliminary data suggest that the hybrid counter-selectable marker functions in the cell and support our hypothesis that fluoroquinolone-resistant parC is recessive.

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3. Spectinomycin Resistance and its Implications for the Evolution of Antibiotic Resistance We continued to develop and refine genetic tools to study B. burgdorferi on a molecular level. In the 1990s, coumermycin A1 resistance was the only available selectable marker, and shuttle vectors had neither been identified nor constructed. Our experimental excursions eventually led us to assemble a hybrid spectinomycin- and streptomycin-resistant cassette [18], now one of several selectable markers in the molecular toolbox for borreliologists [16, 55, 56, 59, 60]. This antibiotic resistance marker has been used to disrupt genes [80-83] and to complement mutants [67, 84í88]. 3.1. Development of aadA as a Selectable Marker We originally attempted to construct a shuttle vector by cloning the coumermycin A1resistant selectable marker into a variety of broad-host-range plasmids: pRKY55 (a derivative of pTJS133) [89], pUFR047 (a derivative of pSa747) [90], pBBR1MCS [91], and pDSK600 and pVLT35 (derivatives of pRSF1010) [92, 93]. None of these recombinant plasmids replicated independently in B. burgdorferi for us. However, when we transformed with pVLT35 carrying a coumermycin A1-resistant gyrB, a single crossover recombination event resulted in the insertion of the intact recombinant plasmid in the B. burgdorferi chromosome and the consequent duplication of the gyrB gene [94], which has been observed by others [95]. The recombined plasmid also carried the aadA gene [96], which confers resistance to spectinomycin and streptomycin. Preliminary data suggested that the transformant had low-level resistance to these antibiotics, which motivated us to fuse a B. burgdorferi promoter to the aadA gene, as previously described [55], and clone it onto the extant shuttle vector pBSV2 [66]. The new plasmid, pBV102, conferred a high level of resistance to both spectinomycin and streptomycin to B. burgdorferi. Therefore we replaced the aphI open reading frame, which encodes kanamycin resistance, on pBSV2 with the aadA open reading frame, thus creating pKFSS1 [18]. pKFSS1 conferred resistance to spectinomycin and streptomycin and could be comaintained in transformants with its parental plasmid pBSV2 or with pCE320; selection of B. burgdorferi transformants with spectinomycin failed, resulting in many colonies that did not contain the plasmid [18]. Note that the hybrid cassette also confers resistance to spectinomycin and streptomycin in E. coli, but many common laboratory strains are resistant to streptomycin. Because of these limitations, spectinomycin is used to select in E. coli, and streptomycin is used to select in B. burgdorferi. 3.2. Spectinomycin-Resistant Mutants Have a Low Fitness Cost We were curious why spectinomycin selection failed, yielding large numbers of spectinomycin-resistant B. burgdorferi colonies [18]. We first hypothesized that these clones had a mutation in the small subunit of their ribosomes, a well-established target for spectinomycin. Spectinomycin-resistant mutations have been described in the S5 protein or the 16S rRNA of several organisms. All of the B. burgdorferi spectinomycinresistant clones had mutations in the 16S rRNA gene, encoding either A1185 to G or C1186 to U [15]. These mutants were over a 1000-fold more resistant to spectinomycin than wild type. We also isolated mutants resistant to the closely related aminoglycoside

D.S. Samuels / Antibiotic Resistance in Borrelia burgdorferi

8 6 4 2 0 WT

16S 16S 16S C1186U A1185G A1402G

S12 K88E

Antibiotic-resistant colonies (%)

B.

10

Doubling time (h)

A.

50

65

100

generations

50 40 30 20 10 0 WT + WT + C1186U A1185G

WT + K88E

WT + A1402G

Figure 3. Growth rate and competition of antibiotic resistant mutants of B. burgdorferi compared to wildtype (WT). A. The doubling time of the wild-type strain is not significantly different from the doubling times of the antibiotic-resistant mutants, although the kanamycin- and gentamicin-resistant mutant (16S A1402G) grows slightly slower; values are means plus the standard error of the means (SEM) for a total of five replicates in three independent experiments. B. The two spectinomycin-resistant mutants (C1186U and A1185G), but neither the streptomycin-resistant mutant (K88E) nor the kanamycin- and gentamicin-resistant mutant (A1402G) successfully compete with the wild-type strain during 50 to 100 generations of coculture; values are means ± SEM for three to five independent experiments.

antibiotics kanamycin, gentamicin, and streptomycin, which also target the small subunit of the ribosome. The kanamycin-resistant and gentamicin-resistant clones both had an A1402 to G mutation in their 16S rRNA and were about 100-fold resistant to either antibiotic [15]. The clones selected in streptomycin were about tenfold resistant and had mutations in rpsL, encoding Lys-88 to either Arg or Glu in the S12 protein [15]. All of these mutations in B. burgdorferi had precedence in other organisms. The frequency of the spectinomycin-resistant mutants was 6 × 10–6, which was about 100fold higher than the frequency of the aminoglycoside-resistant mutants. We next hypothesized that there was a large subpopulation of spectinomycinresistant cells with a 16S rRNA mutation in the parental strain, a highly passaged B31, and that the high frequency was a result of a lower fitness cost for the 16S rRNA C1186 to U and A1185 to G mutations conferring spectinomycin resistance. Again, there was precedence in other organisms, historically in E. coli and more recently in Chlamydia psittaci [97]. Growth assays (Figure 3A) demonstrated that the spectinomycin-resistant mutants grew at the same rate as wild-type in vitro without selection; however, a streptomycin-resistant strain with a Lys-88 to Glu mutation in the S12 protein also grew at the same rate [15]. To assess fitness, competition assays followed mixed cultures of wild type and antibiotic-resistant mutants over 50 or 100 generations [15]. The competition assays (Figure 3B) showed that the spectinomycinresistant mutants were maintained in the co-culture at close to the starting frequency for 50 generations and were still a significant presence in the culture after 100 generations, while the aminoglycoside-resistant mutants were greatly diminished at 50 generations and absent from the culture at 100 generations [15]. These results suggested that there was little fitness cost associated with spectinomycin resistance, especially with the C1186 to U mutation.

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3.3. Antibiotic Resistance in the Clinic and in the Laboratory Antibiotic resistance has not yet become a recognized problem in treating Lyme disease. In addition, spectinomycin and the other antibiotics discussed in this chapter are not used to treat B. burgdorferi infections. However, our results indicate that B. burgdorferi can become resistant by mutation of antibiotic targets. Other resistance mechanisms, such as preventing an antibiotic from entering a cell, pumping an antibiotic out of a cell, or inactivating an antibiotic, have also not been identified in clinical isolates; however, there is no a priori reason that antibiotic-resistant mutants cannot emerge in Lyme disease patients. Moreover, our data on spectinomycin resistance suggest that certain antibiotic-resistant mutants have minimal fitness cost and thus may be able to survive even in the absence of the antibiotic, which may have a profound effect on the evolution and dissemination of resistance [98, 99]. Although discussion of antibiotic resistance in Lyme disease patients is purely conjectural, there are profound implications for developing genetic markers used for experimentally manipulating B. burgdorferi. Resistance to aminoglycosides [18, 55, 59, 60] has been extensively exploited in these endeavors [4]. The high frequency of spectinomycin-resistant mutants in a population of B. burgdorferi cells constrains selection with the aadA cassette to E. coli [18] and serves as a caveat in augmenting the genetic tools for progressively elaborate molecular experiments.

Acknowledgments I thank Christian Eggers and Amanda Ng for their thoughtful and critical reading of this manuscript; the past and current members of my laboratory, especially Janet Alverson, Sharyl Bundle, Dan Criswell, Betsy Eggers, Christian Eggers, Kristi Frank, Kendal Galbraith, Mike Gilbert, Scott Knight, Craig Kuchel, Meghan Lybecker, Kathy Mach, Mike Mazzotta, Elizabeth Morton, Amanda Ng, Virginia Tobiason, and Beth Todd; and my colleagues, especially Steve Lodmell, Mike Minnick, Nyles Charon, Mike Norgard, Justin Radolf, Kit Tilly, Rich Marconi, Xiaofeng Yang, George Chaconas, Phil Stewart, Chuck Sohaskey, Wai Mun Huang, Claude Garon, Cathy Lawson, Thad Stanton, Melissa Caimano, Bob Gilmore, Darrin Akins, Jon Skare, Aravinda de Silva, John Leong, Ira Schwartz, Mathieu Picardeau, Jim Miller, Steve Norris, Alan Barbour, David Haake, Erol Fikrig, Tom Schwan, Patti Rosa, Joe Hinnebusch, Karl Drlica, Tony Maxwell, Peter Heisig, and Christian Eggers. Our research is supported by a grant from the National Institutes of Health (AI051486) and was supported by a grant from the National Science Foundation (MCB-9722408).

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Development of Treponeme Genetic Systems a

Howard K. KURAMITSU a,1 and Caroline E. CAMERON b Department of Oral Biology, State University of New York at Buffalo, Buffalo, NY, USA b University of Washington, Seattle, WA 98195, USA Abstract. Electroporation of Treponema denticola allowed for construction of monospecific mutants in this oral spirochete. The subsequent construction of shuttle plasmids enabled the expression of heterologous genes in these organisms. More recently, a comparable expression system has been developed in T. phagedenis. Keywords. Treponema denticola, shuttle plasmids, electroporation, mutants, Treponema phagedenis

Introduction Treponemes have been recognized as being responsible for a number of human diseases including syphilis [1]. In addition, a variety of these organisms are associated with the common human malady periodontitis [2]. The relatively fastidious nature of these organisms has made it difficult to cultivate and examine them in the laboratory. This has been especially true in attempting to apply molecular genetic approaches in defining the virulence properties of these organisms. This review will describe the development of genetic systems in the past decade for examining the physiological properties of selected treponemes. One of the spirochetes associated with periodontitis, Treponema denticola, can be cultivated in the laboratory and has served as a model organism for noncultivable members of this genus [2]. Biochemical approaches identified several potential virulence factors of these organisms including a major outer surface protein, Msp [3], as well as a serine protease, PrtP [4]. However, the absence of mutants defective in these proteins made it difficult to verify the role of each of these factors in pathogenicity. For this reason, initial attempts were carried out to develop a gene transfer system in these organisms.

1 Corresponding Author: Howard K. Kuramitsu, Department of Oral Biology, SUNY at Buffalo, Buffalo, NY 14214; E-mail: [email protected].

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1. Gene Transfer in T. denticola Initially, markers for detecting transfer of heterologous plasmids into T. denticola were sought by examining the antibiotic sensitivity of two strains of the organism, ATCC 33520 and 35405 [5]. This analysis suggested that either erythromycin or chloramphenicol might allow for the detection of gene transfer into these organisms. Therefore, the broad-host range plasmid encoding chloramphenicol resistance, pKT210, was electroporated into strain 33520 and transformants were identified [5]. Although some modification of the plasmid was detected, this demonstration indicated that it was possible to introduce genetic material into T. denticola following electroporation.

2. Mutant Construction in T. denticola The demonstration of gene transfer into T. denticola 33520 suggested that electroporation might be useful to construct monospecific mutants in these organisms. Therefore, since the background of spontaneous chloramphenicol resistant mutants in T. denticola appeared to be higher than erythromycin resistant mutants, the ermFermAM cassette [6] was utilized to interrupt T. denticola genes. Initially, mutagenesis of the flgE gene of strain 35405 was attempted since inactivation of this gene coding for the flagellar hook protein would lead to an easily recognizable nonmotile phenotype. Following electroporation of an flgE gene fragment interrupted with the erm cassette into strain 35405, Ermr colonies were identified that appeared to be nonmotile on agarose plates [7]. Confirmation of the correct double cross-over integration event indicated that this procedure could be used to construct monospecific mutants in T. denticola. Subsequently, a number of different laboratories have used this procedure to construct defined mutants of the organism [8]. Most recently, this approach has been utilized to identify T. denticola genes that are involved in synergistic biofilm formation with Porphyromonas gingivalis [9]. This was also one of the first reports examining biofilm formation by a spirochete. The availability of a system to inactive genes in T. denticola also allows for the identification of factors that are involved in the potential virulence of these organisms. For example, inactivation of the prtP gene coding for the serine protease dentilisin of strain 35405 followed by inoculation into mice has demonstrated that the protease is a likely virulence factor in these organisms [10].

3. Heterologous Gene Expression in T. denticola Since a number of pathogenic treponemes cannot be readily cultivated in the laboratory, the ability to express genes from these organisms in a genetically tractable treponeme could provide a means to investigate genes from the former organisms. Therefore, the utilization of T. denticola for such purposes was investigated. In order to construct a shuttle plasmid for use in T. denticola, a replication region from a naturally occurring plasmid was sought. In this regard, two plasmids were identified, pTD1 and pTS1, and characterized in two different strains of T. denticola [11, 12]. Therefore, based upon the nucleotide sequence of plasmid pTS1 [13], a putative replication region was identified in this plasmid and was used to construct a shuttle plasmid that could be utilized in both T. denticola and Escherichia coli [14]. The plasmid also contained the ermFermAB

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cassette immediately downstream from the promoter region of the T. denticola prtB gene. Using this shuttle plasmid, pKMR4PE, it was possible to express the flaA gene from T. palllidum in T. denticola 33520 [14]. More recently, Limberger and coworkers [15] have constructed a similar plasmid expressing chloramphenicol resistance and have expressed the fliG gene from T. pallidum in strain 33520. Interestingly, shuttle plasmids could be successfully introduced into strain 33520, which harbors the naturally occurring plasmid pTD1, but not in strain 35405, which does not contain a plasmid. However, the molecular basis for this difference has not yet been elucidated. It was also of interest to determine if nontreponemal genes could be expressed in T. denticola as reporter genes. Therefore, the lacZ gene from E. coli coding for ȕgalactosidase activity was engineered downstream of the erm cassette in plasmid pKMR4PE and introduced into strain 33520. On X-gal agarose plates, the transformants appeared dark blue while the control strain was colorless (Figure 1). Direct assays for enzyme activity also showed a significant increase in the strain harboring the lacZ-containing plasmid relative to 33520 with the shuttle plasmid alone. These results indicated that some nontreponemal genes could also be expressed in T. denticola and that the lacZ gene could be used as a reporter gene in these organisms. More recently, the same shuttle plasmid was utilized to express the green-fluorescence protein gene in T. denticola 33520 although the resulting fluorescence was not as stable nor as strong as the protein expressed in E. coli [16]. However, it may be possible to express stronger fluorescence in these organisms using more recently constructed gfp genes.

4. Complementation Analysis in T. denticola One of the advantageous of developing a shuttle plasmid system is the ability to express genes in the organism for complementation of either mutations or naturally occurring defects. For complementation of Ermr mutants in strain 33520, a shuttle plasmid based upon the pKMR4PE plasmid but expressing coumerymycin resistance was developed [17]. This plasmid was utilized to express the T. denticola flgE gene in the strain 33520 flgE mutant and was demonstrated to restore the motility of the mutant. More recently, a similar shuttle plasmid expressing a T. pallidum fliG gene was utilized to complement a fliG mutation in strain 33520 [15]. Therefore, these systems can be valuable in characterizing genes isolated from noncultivable treponemes such as T. pallidum. Because of the inability to establish the pTS1-based shuttle plasmids in strain 35405, it was not possible to complement mutants constructed in this strain. A comparable interference strategy was also used to confirm a role for the leucine-rich repeat protein, LrrA, in the motility of T. denticola [18]. When the lrrA gene from strain 35405 was introduced into strain 33520, which is motile and does not contain a homolog of the lrrA gene, the resultant transformant was attenuated in motility. This confirmed a role for the LrrA protein in the motility of T. denticola as suggested from the properties of an lrrA mutant in strain 35405.

74

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A

B

Figure 1. lacZ expression from T. denticola 33520. A, colonies on X-gal agarose plates containing pKMR4PE-lacZ; B, shuttle plasmid alone. Arrows indicate location of colonies.

5. Expression of Heterologous Treponeme Genes in T. phagedenis Since T. pallidum and T. denticola have been isolated in humans, it was of interest to develop a gene transfer system in a treponeme which is not a normal human colonizer since such an organism would be expected to be physiologically distinct from those inhabiting the human host. T. phagedenis is one such organism and also is very similar to T. denticola in its G+C ratio [19]. Therefore, the shuttle plasmids developed for use in T. denticola might also function in T. phagedenis as well. Utilizing T. phagedenis Kazan, it was indeed possible to demonstrate the transformation of shuttle plasmid pKMR4PE into this organism (Yamada and Kuramitsu, unpublished results). Therefore, the same shuttle plasmid system used in T. denticola was evaluated in T. phagedenis for expression of heterologous treponeme genes. One of the potential virulence properties of T. pallidum is its ability to attach to the extracellular matrix, ECM, including the glycoprotein laminin [20]. Recently, one of our laboratories has isolated a gene from T. pallidum coding for the laminin-binding adhesin Tp0751 [20]. Confirmation of the role of this adhesin in such binding was sought by expression of a laminin-binding domain of Tp0751 in the non-lamininbinding T. phagedenis. Using a derivative of the T. denticola-E. coli shuttle plasmid pKMR4PE, a 159 bp DNA fragment from the tp0751 gene containing its ribosome binding site, putative signal sequence, and amino acids involved in laminin binding [21] was introduced into the shuttle plasmid. Transformation of T. phagedenis following the electroporation protocol developed for T. denticola [14] allowed for the identification of transformants harboring the correct chimeric plasmid. Northern blot analysis allowed for the identification of the mRNA corresponding to the T. pallidum insert (data not shown). To determine if T. phagedenis expressing the laminin-binding domain of Tp0751 could mediate binding to laminin, laminin-coated glass slides were incubated with the transformant and binding compared with the organism containing only the shuttle plasmid under dark field microscopy. These results indicated that the construct expressing the laminin-binding domain, but not that containing only the shuttle plasmid, bound to the laminin-coated slides. Quantitation of these results by

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75

counting the number of organisms in six separate fields revealed that there was a significant increase in attachment when the Tp0751 laminin-binding domain was expressed in T. phagedenis (Figure 2). These results confirmed the laminin-binding activity of the Tp0751 adhesin and also indicated that T. phagedenis can serve as a convenient treponeme host for the expression of genes from some noncultivable treponemes.

Number of Treponemes/Field

100

WT pKMR 751/pKMR

75

p<0.0001

50

25

0 BSA

laminin

Coating Reagent Figure 2. Expression of the Tp0751 laminin-binding domain in T. phagedenis.

6. Summary The development of gene transfer systems for use in T. denticola and T. phagedenis now makes it possible to carry out genetic analysis of these organisms as well as comparable studies for noncultivable treponemes. In the later regard, the expression of several genes from T. pallidum in both of these transformable treponemes has demonstrated the feasibility of this approach. It will be of interest to determine if similar gene transfer systems can be developed for other cultivable treponemes. Since the shuttle plasmids utilized in expressing heterologous genes in T. denticola could not be established in strain 35405, it also will be of interest to determine the molecular basis for this property. This further suggests that for some treponemes additional shuttle plasmids derived from naturally occurring plasmids other than pTS1 may need to be developed. In addition, it will be of interest to determine if additional genetic techniques, such as transposon mutagenesis, can be developed in the treponemes. As the genetic analysis of treponemes is in its infancy, we look forward to further developments in this important area of research.

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Acknowledgments We wish to gratefully acknowledge the contributions of H. Li, S. Chauhan, B. Chi, A. Ikegami, M. Yamada, N. Charon, J. Ruby, R. Limberger, N. Brouwer, L. Tisch and P. Cullen to the studies cited in this review. The work cited from our laboratories was supported in part by NIH grants DE09821 (HKK) and AI51334 (CEC).

References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21]

C.W. Penn and D.G. Prtichard. The spirochetes. In Tarpley and Wilson’s Principles of bacteriology, virology, and immunology. Vol. 2. pp. 603–628. Edward Arnold, London. 1990. W.J. Loesche and B. Laughon. Role of spirochetes in periodontal disease. In R.J. Genco and S.E. mergenhagen (eds). Host-parasite interactions in periodontal disease. pp. 62–75. American Society for Microbiology, Washington, D.C. 1982. M. Haapasalo, K.H. Muller, V.J. Uitto, W.K. Leung, and B.C. McBride. Characterization, cloning, and binding properties of a major 53-kilodalton Treponema denticola surface antigen. Infect. Immun. 60 (1992), 2058–2065. V.J. Uitto, D. Grenier, E.C. Chan, and B.C. McBride. Isolation of a chymotrypsinlike enzyme from Treponema denticola. Infect. Immun. 56 (1988), 2717–2722. H.Li and H.K. Kuramitsu. Development of a gene transfer system in Treponema denticola by electroporation. Oral Microbiol. Immunol. 11 (1996), 161–165. H.M. Fletcher, H.A. Schenkein, R.M. Morgan, K.A.Bailey, C.R. Berry, and F.L. Macrina. Virulence of a Porphyromonas gingivalis W83 mutant defective in the prtH gene. Infect. Immun. 63 (1995), 1521– 1528. H. Li, J. Ruby, N. Charon, and H. Kuramitsu. Gene inactivation in the oral spirochete Treponema denticola: construction of a flgE mutant. J. Bacteriol. 178 (1996), 3664–3667. H.K. Kuramitsu. Genetic manipulation of cultivable treponemes. In S. Lukehard and J. Radolf (eds). Molecular biology and pathogenisis of treponemeal infections. Horizon Press, London (in press). M. Yamada, A. Ikegami, and H.K. Kuramitsu. Synergistic biofilm formation by Treponema denticola and Porphyromonas gingivalis. FEMS Microbiol. Lett. 250 (2005), 271–277. K. Ishihara, H.K. Kuramitsu, T. Miura, and K. Okuda. Dentilisin activity affects the organization of the outer sheath of Treponema denticola. J. Bacteriol. 180 (1998), 3837-3844. A. Ivic, J. MacDougall, R.R.B Russell, and C.W. Penn. Isolation and characterization of a plasmid from Treponema denticola. FEMS Microbiol. Lett. 78 (1991), 189–194. E.C. Chan, A. Klitorinos, S. Gharbia, S.D. Caudry, M.D. Rahal, and R. Siboo. Characterization of a 4.2 kb plasmid isolated from peridontopathic spirochetes. Oral Microbiol. Immunol. 11 (1996), 365–368. S. Chauhan and H.K. Kuramitsu. Sequence analysis of plasmid pTS1 from oral spirochetes. Plasmid 51 (2004), 61–65. B. Chi, S. Chauhan, and H. Kuramitsu. Development of a system for expressing heterologous genes in the oral spirochete Treponema denticola and its use in expression of the Treponema pallidum flaA gene. Infect. Immun. 67 (1999), 3653–3656. L.L. Slivienski-Gebhart, J. Izard, W.A. Samsonoff, and R.J. Limberger. Development of a novel chloramphenicol resistance expression plasmid used for genetic complementation of a fliG mutant in Treponema denticola. Infect. Immun. 72 (2004), 5493–5497. I.Saint Girons, B. Chi, and H. Kuramitsu. Development of shuttle vectors for spirochetes. J. Mol. Microbiol. Biotechnol. 2 (2000), 443–445. B. Chi, R.J. Limberger, and H.K. Kuramitsu. Complementation of a Treponema denticola flgE mutant with a novel coumermycin A1-resistant Treponema denticola shuttle vector system. Infect. Immun. 70 (2002), 2233–2237. A. Ikegami, K. Honma, A. Sharma, and H.K. Kuramitsu. Multiple functions of the leucrine-rich repeat protein LrrA of Treponema denticola. Infect. Immun. 72 (2004), 4619–4627. I. Olsen, B.J. Paster, and F.E. Dewhirst. Taxonomy of spirochetes. Anaerobe 6 (2003), 39–57. C.E. Cameron. Identification of a Treponema pallidum laminin-binding protein. Infect. Immun. 71 (2003), 2525-2533. C.E. Cameron, N.L. Brouwer, L.M. Tisch,and J.M.Y. Kuroiwa. Defining the interaction of the Treponema pallidum adhesin Tp0751 with laminin. Infect. Immun. 73 (2005), 7485–7494.

Part 2 Genomics and Diversity

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Comparative Genomics of Borrelia burgdorferi Sherwood R. CASJENS a,1 , Wai Mun HUANG a, Eddie B. GILCREASE a, Weigang QIU b, William D. MCCAIG b, Benjamin J. LUFT c, Steven E. SCHUTZER d and Claire M. FRASERe a Pathology Department, University of Utah School of Medicine, Salt Lake City, UT 84132 b Department of Biological Sciences, Hunter College of the City University of New York, 695 Park Avenue, New York City, NY 10021 c Department of Medicine, Health Science Center, Stony Brook University, Stony Brook, NY 11794 d Department of Medicine, New Jersey Medical School, 185 South Orange Avenue, Newark, NJ 07103 e The Institute for Genomic Research, 9712 Medical Center Drive, Rockville, MD 20850 Abstract. We have determined draft whole genome sequences for B. burgdorferi isolates N40 and JD1 and the plasmids from isolate 297. The comparison of these draft sequences sheds considerable light on a number of aspects of their biology including horizontal exchange of genetic information, plasmid diversity, lipoprotein variability, and plasmid partitioning. We find that plasmids cp9, cp26, the cp32s, and lp54 are quite constant in gene organization among the four strains, but the remaining linear plasmids are quite variable in organization. We analyze the replication, partitioning, and compatibility gene cluster on the plasmids from these four strains and propose to use them in a universal B. burgdorferi plasmid nomenclature system. Keywords. Borrelia burgdorferi, Lyme disease, plasmids, partitioning, genome sequences

Introduction Comparison of whole genome sequences or “comparative genomics,” both within and between species, is becoming a critical part of a true understanding of genome sequences. This strategy allows the discovery of invariant (and therefore most likely evolutionarily important) features of the compared genomes, allows discovery of variable sequences that permit tracking of evolutionary branches within species, and 1 Corresponding Author: Pathology Department, University of Utah Medical School, 30 North 1900 East, Salt Lake City, UT, USA, 84103; E-mail: [email protected]

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gives critical insight into evolutionary mechanisms. In addition, genome comparison between closely related genomes can often illuminate inaccuracies in the prediction of genes and other features in genomes. The nucleotide sequence of the genome of the type strain of the Lyme disease spirochete Borrelia burgdorferi B31 was completed about five years ago [1, 2]. That genome sequence and associated experimental analysis of the genome demonstrated that the B. burgdorferi genome is very unusual, as it is made up of many more replicons (DNA molecules) than other bacteria, a number of which are linear with closed hairpin ends. In addition to its 910 kbp linear chromosome, the sequenced culture of strain B31 carries twelve linear and ten circular plasmids [1, 2], and two additional circular plasmids are now thought to have been lost between strain B31’s original isolation and its genome sequencing [3í5]. This work confirmed Barbour’s [6] original groundbreaking observations and following studies in many laboratories that Borrelia isolates universally harbor numerous linear and circular plasmids [7í17]. However, because of the many paralogous sequences present among the plasmids, a lack of information on the range of plasmid variation among isolates, and the fact that many of the plasmids are similar in size (and thus not separable in electrophoresis gels), members of the Borrelia genus are currently in an unenviable position. In order to know the complete complement of plasmids present in any given strain, a complete genome sequence must be determined. Thus strain B31 remains the only Borrelia isolate for which all resident plasmids have been identified. Although the strain B31 genome sequence has been of great value to Lyme disease research, many questions remain regarding the molecular pathogenesis of Lyme disease, the interaction of the bacteria with their arthropod tick vector Ixodes scapularis, as well as the diversity and evolutionary history of the Lyme disease bacteria. The B. burgdorferi isolates that are most commonly used in laboratory studies are isolates B31, N40, and 297 [18í20]. Both N40 and 297 have been anecdotally found to have significantly different plasmid contents from strain B31, so lack of knowledge of their genome sequences has been a hindrance to some aspects of Lyme disease research.

1. Draft Sequences of Three Borrelia burgdorferi Genomes In order to begin to attack the Lyme agent unknowns mentioned above, we have determined draft sequences of the complete genomes of B. burgdorferi strains N40 and JD1 and determined a draft sequence of the plasmids of strain 297. These three strains were chosen for sequencing because they include the two commonly used laboratory strains N40 and 297 and because they represent a reasonable sample of the known B. burgdorferi diversity. With that of strain B31, the four sequenced genomes include members of all three rRNA spacer sequence types [21], four of the eight different pulsed-field gel DNA pattern types [22], and four of the nineteen OspC types [23] that have been described (Table 1). Strain 297 was isolated from a human patient with Lyme disease, while the others are tick isolates; all were isolated in the northeastern part of United States of America, a region with a high frequency of human Lyme disease. Large and small insert DNA libraries were constructed with DNAs from each of the three strains, and random shotgun sequencing was carried out until an average of at least 8-fold coverage was obtained; then the TIGR ASSEMBLER computer program was used to assemble as much of the sequence as possible [1,25]. We have previously reported an analysis of some of the nucleotide polymorphisms among orthologous

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81

sequences in these draft genome sequences, which strongly indicates that B. burgdorferi has a history that includes a substantial amount of horizontal transfer of genetic information [26]. These draft genome sequences have a number of limitations relative to completed genome sequences [27], and completion of the N40, JD1, and 297 genome sequences is currently underway. In this report, we describe several preliminary observations derived from these draft genome sequences. Since these are draft sequences, it remains possible that some of our preliminary conclusions will require modification when the genomes are completed. To infer more robust information from these draft sequences, unique DNA probes were identified from each of the contigs, and these probes were used in Southern analyses [28] to determine the size of intracellular DNA molecule from which each of the bulk sequencing contigs was derived (data not shown). The combined observations show that strains B31, N40, JD1, and 297 carry 21, 16, 20, and 19 plasmids, of which 12, 8, 11, and 9 are linear, respectively (Table 2). Many of the plasmids resident in any Borrelia strain can be lost with passage in culture [7, 29í35]. The fact that DNA was sequenced from very low passage cultures and parallel cultures are infectious in the laboratory suggests that at least most of their native plasmids should be present in the genome sequences.

Table 1. B. burgdorferi isolates used in this study. Isolate

PFG type 1

rRNA spacer type 2

OspC type 3

Source

Reference

B31

B

I

A

Ixodes tick/NY

[18]

N40

E

III

E

Ixodes tick/NY

[19]

JD1

C

II

D

Ixodes tick/MA

[24]

297

A

II

K

Human/CT

[20]

1

There are at least eight B. burgdorferi pulsed field gel (PFG) pattern types [9, 22]. There are three rRNA spacer types as determined by Wormser et al. [21]. 3 There are at least nineteen OspC types as determined by Wang et al. [23]. 2

In agreement with conclusions from their previously reported chromosomal macrorestriction maps [9], the linear large chromosomes of the B. burgdorferi strains B31 and JD1 are co-linear at the “draft sequence” level of analysis (no N40 map has been determined, and the 297 chromosome was not sequenced). The B31, N40, and JD1 chromosomes have no whole gene insertions or deletions when compared to one another, with the exception of the extreme right end, which carries differing amounts of plasmid-like DNA in each strain [36, 37] (to be described in detail elsewhere). This is in agreement with the report by Glöckner et al. [38] that the chromosome of Borrelia garinii strain PBi is co-linear with that of B. burgdorferi B31 except for only eight insertions and six deletions that are greater than 100 bp in length (the longest of which is 1878 bp). This observation places the B. burgdorferi large chromosome among the most evolutionary stable chromosomes of all the bacteria for which comparative genomics has been done. The three chromosomes are all between 0.46% and 0.61% different from one another in nucleotide sequence in pair-wise comparisons [26].

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82

2. Plasmid Conservation Among B. burgdorferi Strains Some of the plasmids are known to be quite highly conserved and carry few sequences that are very closely related to sequence on other plasmids [2]. These plasmids, cp9, cp26, and lp54, are present in our “draft” sequences as complete, closed sequences (except for the tips of the linear plasmid lp54). Not surprisingly, since they both have previously been found to be present with similar structures and sequences in essentially all other natural isolates (e.g., [11, 44]), and cp26 is thought to be essential [45], we find that linear plasmid lp54 and circular plasmid cp26 are the most highly organizationally conserved among the B. burgdorferi plasmids. They are both present in B31, N40, JD1, and 297 with no gene content differences detected at the “draft sequence” level of analysis. Plasmid cp9 has been previously sequenced in B. burgdorferi strains B31 [2] and N40 [46] and B. garinii IP21 [47] and found to be quite similar in all three isolates. Among our three new B. burgdorferi draft genome sequences, only strain N40 carries a plasmid that is similar to cp9 of B31. Our N40 8722 bp cp9 sequence has seven differences from the previously reported N40 cp9 sequence [46] (two 1 bp insertions, one of which truncates the eppA gene in the previous sequence; four 1 bp differences; and an A at the “A or G” at the previously published position 7315). Figure 1 shows that the B31 and N40 plasmids have very similar gene organizations; however, one gene (BBC10) and part of another (BBC08) that are present on the B31 cp9 have been replaced in N40 cp9 by several hundred bp of apparently non-protein-coding DNA (this report and [46]). Also in agreement with previous observations [3, 48í50], we find that all four of these strains carry multiple circular plasmids in the cp32 family (Table 2), with generally similar organization, although N40 and 297 each carry two cp32-like plasmids that appear to have suffered deletions in the 8 to 12 kbp range relatively recently in evolutionary time (see also [39, 40, 42]), and JD1 has a plasmid that is made up of two different fulllength cp32s fused together into a 61.6 kbp circular plasmid. Table 2. Number of plasmids carried by four B. burgdorferi isolates. B31

N40

Total plasmids

21

Linear plasmids

12

Circular plasmids

9

297

16

20

19

8

11

9

8

9

10

lp54

1

+

+

+

+

cp26

+

+

+

+

+

+

-

-

3

4

cp9 cp32s 1

JD1

2

8

6

8

5

94

lp54 and cp26 are the most highly conserved plasmids among these B. burgdorferi isolates. A “+” indicates the presence of very similar plasmids in all four strains; a “–” indicates no homolog is present. 2 The multiple cp32s present in each isolate are highly conserved in overall structure, but have some regions of higher variability and/or non-homology [1, 3]. 3 Those present in the culture of strain B31 whose genome was sequenced; this value includes the “cp32” that is integrated into linear plasmid lp56 [1]. 4 Two cp32 plasmids in N40 and 297 have (different) substantial deletions (see also [39, 40]). The cp32 contents of strains N40 and 297 have been previously studied by several laboratories [e.g., 40-43]. 5 One JD1 circular plasmid is a fusion of two cp32-type plasmids.

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83

B31 cp9 PFam per cent identity

57

50 49

161

eppA 95

55

rev 63

96

C01

C02 C03

C05

C06

C08

C10

C11

1000

2000

3000

4000

5000

6000

7000

165 C12 8000

9000

95.6 1000

73.8 96.8

2000

3000

99.0

N40 cp9

80.3 90.4

4000

5000

6000

99.4 7000

99.1

8000

Figure 1. Matrix comparison of cp9 plasmids from B. burgdorferi strains B31 and N40. A scanning window of 17 matches out of 23 bp was used in the computer program DNA STRIDER [57]. Open reading frames are indicated as block arrows, pointing in the direction of transcription. Above the B31 open reading frames the paralogous family [1] to which the genes belong is indicated. Light gray genes are involved in plasmid replication and partitioning (see text and [40]), and white genes encode proteins that have been found to elicit antibodies in infected rabbits and humans. The latter genes, BBC06 and BBC10, have been named eppA and rev, respectively, in those previous studies [58í60]. On the left, per cent nucleotide sequence identity values are shown for each apparently intact N40 gene relative to its B31 orthologue .

Even the organizationally highly conserved plasmids lp54, cp9 and cp26, have regions that are more substantially more variable than the bulk of the genomes [26]. For example, Figure 1 shows that although the protein coding regions of B31 cp9 and N40 cp9 are 93.9% identical overall in nucleotide sequence similarity, the similarities of the different orthologous cp9 genes range from 73.8% to 99.1%. On plasmid lp54 genes BBA07, BBA24, BBA68, BBA69, and BBA70 are more variable than the rest of the plasmid [26]. Table 3 shows the relationships among the highly variable ospC gene (on circular plasmid cp26) in the four sequenced genomes compared the relationships among the rest of the cp26 plasmids; ospC may well be the most variable locus in the B. burgdorferi genome [22, 26, 51í54]. Such uneven distributions of levels of similarity are often considered to be evidence of past horizontal exchange events

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84

between homologous sequences that have been separated for long evolutionary times, but in the case of ospC a balancing selection for diversity is almost certainly at least partly responsible [23, 26, 55].

Table 3. The hypervariable region on plasmid cp26. ospC gene

cp26 minus ospC1

B31 vs. N40

17.4%2

1.18%2

B31 vs. JD1

13.5%

1.29%

B31 vs. 297

14.5%

1.54%

N40 vs. JD1

16.2%

1.38%

N40 vs. 297

14.2%

1.56%

JD1 vs. 297

13.5%

0.83%

1

The entire cp26 plasmids with the coding region of the ospC gene (BBB19) removed.

The “non-lp54” linear plasmids in all four of the genome sequences have some properties that are similar to those of lp54, cp9, and cp26 discussed above, in that they are mosaics or patchworks of nucleotide sequences that are highly similar (97í100% identical) to B31 plasmid sequences that are intermixed with regions that are homologous but much less similar. However, like their B31 relatives, these plasmids also appear to carry a substantial amount of non-protein-coding DNA sequences and numerous pseudogenes (see discussion of this aspect of the strain B31 plasmids in refs. [1, 4, 56]). When these “non-lp54” plasmids are compared to the B31 plasmids, there are two additional important differences. First, there are some regions (a relatively small fraction of the total) that are unrelated to any B31 sequence, and the organization of the sequences that are highly similar to B31 sequences is often different from that of strain B31. There appear to have been numerous rearrangements and exchanges of genetic information among these linear plasmids since their last common ancestor. Although these N40, JD1 and 297 linear plasmids have substantial organizational differences when compared to the strain B31 linear plasmids, a detailed description of their gene organizations must await the currently underway upgrading of the draft sequences to final, “closed” genome sequences.

3. Plasmid Genes As mentioned above, the less variable plasmids, lp54, cp9, cp26, and the multiple cp32s have very similar gene content and organization in each of the four of the B. burgdorferi isolates under consideration in this report, and the remaining linear plasmids appear to be somewhat variable in number (Table 1) and structure among B. burgdorferi isolates. One of the important features of these “variable” linear plasmids in strain B31 is that they carry numerous paralogous lipoprotein gene families [1], many of which are known to be important in the bacteria’s interaction with its hosts. Since the linear plasmids in strains N40, JD1 and 297 carry largely sequences that are

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homologous to these B31 plasmids, the three draft sequences also contain similar but non-identical paralogous families (PFam’s) of many of these genes. Table 4 shows that four of these gene families, PFams 12, 44, 52, and 60 (as defined in [1]), all contain multiple members in each of the four genomes. Although the numbers of genes present does not vary widely, there are differences among B. burgdorferi strains. (These numbers could change upon finalization of the genome sequences). The biological roles of these four protein families remain unknown, although family 44 is known to be antigenic in infected humans [61].

Table 4. Sizes of several plasmid-borne lipoprotein paralogous gene families. B31

N40

JD1

297

PFam121

3

3

3

3

PFam44

3

3

3

3

PFam52

3

1

3

2

PFam60

8

6

7

8

vls2

1

1

1

1

1

The four paralogous gene families are defined as in Casjens et al. [1]. These values could change upon completion of the N40, JD1, and 297 genome sequences. They include only apparently intact genes and do not include the various apparently truncated or otherwise disrupted, paralogous “pseudogenes” that are also present in these genomes. 2 These values are the number of vls cassette regions present (each region contains multiple cassettes) [62, 63].

4. Plasmid Partitioning The sequence of the 21 B. burgdorferi B31 plasmids revealed that all of the plasmids, both linear and circular, that are larger than 20 kbp carry paralogous four-gene clusters. These genes are members of the paralogous families named PFam32, PFam49, PFam50, and PFam57/62 (PFam57 and PFam62 as originally defined are, in fact, related to one another and so can be considered subgroups of the larger family “PFam57/62” [1]). Each of the four plasmids smaller than 20 kbp, lp5, lp17, lp21 (really about 19 kbp in length), and cp9 carries only a subset of these genes. Figure 2 diagrams the arrangement of these gene clusters on the strain B31 plasmids. Of the four gene families present in these clusters, only the PFam32 members have convincing similarity to other genes in the current nucleotide sequence database, and they are homologous to parA genes, which play an important role in the partitioning of many bacterial plasmids. The function of ParA proteins has been recently reviewed by Leonard [64]. Since PFam32 homologues encode proteins that are important in the successful partitioning of replicated daughter plasmids into the two daughter cells during the subsequent cell division, these highly conserved clusters were proposed to be involved in plasmid replication, partitioning and/or compatibility [1,3,50,65]. Subsequent experimental work has supported these predictions. These gene clusters or

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parts of them, especially the PFam57/62 members, are capable of programming autonomous plasmid replication in Borrelia cells. Rosa, Chaconas, and Radolf, and their co-workers have shown: (i) that DNA constructs (both linear and circular) containing these gene clusters from strain B31 plasmids lp17, lp25, and lp28-1, as well as cp9, from strain N40 and cp32-3 from strain CA-11.2A, are capable of autonomous replication in Borrelia cells; (ii) that such constructs are not able to co-establish residency (i.e., are incompatible) with native plasmids that carry the same partition gene cluster; and (iii) that PFam57/62 genes are essential for plasmid replication [45, 66í70].

cp26

cp9

7 x cp32 lp5

lp17 lp21 lp25 lp28-1

family 49 family 32 family 50 family 57/62

lp28-2 lp28-3 lp28-4 lp36 lp38 lp54 lp56

Figure 2. Putative plasmid replication/partitioning genes on the B. burgdorferi B31 plasmids. Genes from four paralogous families thought to be involved in plasmid replication and partitioning are indicated by different symbols. Squares represent genes that appear to be intact, and rectangles represent pseudogene relatives. Black and gray boxes enclose the PFam32 genes and their clustered partition genes derived from linear and circular plasmids, respectively, that encode the proteins used in the neighbor-joining analyses in Figure 3 and Figure 4 (see below and text). This figure is modified from Figure 8 of Casjens et al. [1].

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The draft genome sequences of B. burgdorferi strains N40, JD1 and 297 show that they carry similar plasmid-borne replication/partitioning genes in very similar clustered arrangements to those in strain B31. A strong prediction of the idea that these genes are involved in plasmid replication, partitioning, and compatibility is that no cell should be able to stably maintain multiple plasmids that have very similar partitioning gene clusters. We therefore compared the sequences of the proteins encoded by the plasmidborne PFam32 genes in the four genome sequences; if, as is sometimes found, there are multiple PFam32 genes on any plasmid (see Figure 2), only the one from the intact cluster was included in the analysis (but see lp56 below). As we have noted before [1], there are numerous fragmented pseudogenes that are related to these four paralogous families scattered across the linear strain B31 plasmids; we have also ignored all PFam32 fragments in the above analysis. The B31 and N40 cp9 plasmids and the B31 lp5 carry no PFam32 gene and so were also not included. With only one exception, the neighbor-joining PFam32 protein trees in Figure 3 and Figure 4 show that this is indeed the case on linear and circular plasmids in all four B. burgdorferi strains that have genomes with complete or draft sequences. Stevenson and Miller [42] have previously shown that this is the case for a number of these cp32 plasmids. The one exception noted above is the two B31 PFam32 proteins of plasmids cp32-2 and cp32-7 in the “cp32-2/7” group in Figure 4; this appears to be the “exception that proves the rule,” in that although both the B31 cp32-2 and B31 cp32-7 plasmids carry PFam32 genes of the same sequence type, these two plasmids have, in fact, never been found in the same cell [1, 3, 42]. The most likely explanations for this are that either the original isolate of strain B31 was a mixture of cells that contained either cp32-2 or cp32-7 (but not both in any cell, or, since the cp32 plasmids are thought to be bacteriophage genomes [71í74], one of these two plasmids was established in a B31 subculture that had lost the other plasmid when it was delivered by a phage virion during handling of the strain after its initial isolation. Thus, the PFam32 proteins fall into a limited number of very robust “sequence type” clusters with very convincing bootstrap values. When all the PFam32 proteins from Figure 3 and Figure 4 are used to build a single large neighbor-joining tree (not shown), the same situation is observed, that is, none of the (still very robust) clusters from the Figure 3 and Figure 4 trees overlap or merge. By the criteria discussed above, we find 27 different PFam32 sequence types, 14 types on linear plasmids, and 13 types on circular plasmids to be present in the four B. burgdorferi genomes. Interestingly, strain B31 linear plasmid lp28-1 carries two clusters of the putative partition genes, but one of them is missing its PFam57/62 member (cluster not boxed in Figure 2). Thus, this PFam32 gene, BBF12 present in the incomplete cluster, was not included in the analysis of Figure 3; however, if it is included it is a 28th PFam32 type and so could represent yet another putative compatibility type that is apparently not present in intact form in any of the four strains analyzed to date. Grimm et al. [69] were able to drive autonomous replication of a circular plasmid with the partition gene clusters from the B31 linear plasmids lp25 and lp28-1, suggesting that there is no obligate mechanistic reason that the genes that drive linear plasmid replication and partitioning should be systematically different from those that

88

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Figure 3. Neighbor-joining tree of PFam32 protein amino acid sequences from B. burgdorferi linear plasmids. CLUSTAL X [75,76] was used to generate the tree; the numbers on the branches represent bootstrap values from 1000 trials. At the right each protein is indicated by a symbol that denotes the Borrelia isolate and the plasmid its gene resides on. The plasmid names are from Casjens et al. [1] and Table 5.

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Figure 4. Neighbor-joining tree of PFam32 proteins amino acid sequences from B. burgdorferi circular plasmids. The tree was generated as in Figure 3, and the cp32 names are those of Stevenson and Miller [42]. At the right each protein is indicated by the Borrelia isolate and the plasmid on which its gene resides.

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drive circular plasmid replication and partitioning. In a neighbor-joining tree of all the PFam32 genes in Figure 3 and Figure 4, the circular and linear plasmid-derived gene types were not robustly separated onto a linear derived branch and a circular derived branch (not shown); thus we also have not yet found anything about the PFam32 gene sequences that suggests that there are mechanistic differences during the segregation of linear and circular plasmids. In addition, the strain B31 linear plasmid lp56 is certainly a composite plasmid in which a 30 kbp cp32 type plasmid has integrated rather recently into a 23 kbp linear plasmid resulting in a 53 kbp linear plasmid [1]. Therefore, this plasmid has two PFam32 genes, one from each of the plasmid-derived gene (BBQ08) in the linear plasmid tree. The PFam57/62 gene associated with BBQ08 on B31 lp56 was broken during the insertion event, so if there is specificity of interaction among the four proteins encoded by a given partition cluster, the cluster from the inserted (circular) cp32 may be driving the replication and partitioning of the linear lp56 plasmid, again suggesting no critical mechanistic differences between linear and circular plasmids in this regard. On the other hand, individual putative PFam32 genes specificity types (clusters in Figure 3 and Figure 4) were never found to include partitioning proteins from both circular and linear plasmids, suggesting that there has not been recent functional exchange of these genes between linear and circular plasmids. This in turn might suggest some subtle differences between linear and circular plasmid partitioning that could be important evolutionarily but are not yet noticeable in the laboratory.

5. Borrelia Plasmid Naming Conventions Historically, Borrelia plasmids have been named according to their topology (linear “lp” and circular “cp”) and their approximate size in kbp; thus, for example, linear strain B31 plasmid lp54 is about 53,600 bp in length. It would be possible to continue this tradition and simply name the plasmids in other strains according to their size; however, this nomenclature scheme has the following serious problems: (i) the fact that most linear plasmids are in the 24 to 31 kbp size range and nearly all known cp32’s are between 29 and 32 kbp means that independent names based on size are severely limited in number; (ii) there are numerous and substantial linear plasmid organizational differences among strains (above); and (iii) there have been many apparently homologous recombination events among the cp32 plasmids (to be reported in detail elsewhere [42]). Thus, a nomenclature scheme based only on plasmid size means that identically named plasmids in different strains would very often have essentially nothing in common (i.e., plasmid size is usually not a useful indicator of any aspect of plasmid gene content). In order to give plasmids names that carry some biological meaning, we propose that when possible, names be given to Borrelia plasmids according to the type of partition genes, in particular the type of PFam32 gene that they carry. Of the four partition cluster gene families, PFam32 and PFam49 are somewhat more diverse than PFam57/62 or PFam50 (data not shown), so we chose PFam32 to give robust separation into sequence types. To date, the only sequenced plasmids that cannot be named by this scheme are the very short plasmids cp9 and lp5, since they do not carry a PFam32 gene. The cp9s sequenced from strains B31 [1], N40 (this report and [46]), B. garinii strain IP21 [47], and B. bissettii strain DN127 cl-9 (our unpublished data; strains JD1 and 297 do not carry a cp9-like plasmid) seem to be quite highly conserved

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and not exchanging genetic information “randomly” with other non-homologous plasmids (above), so naming plasmids “cp9” that are homologous to B31’s cp9 is not likely to be confusing, and the latter plasmid, lp5, has only been found in strain B31 to date. This nomenclature strategy has previously been suggested for the cp32 plasmids by Stevenson and Miller [42], and we show here that it can be applied to all known B. burgdorferi plasmids that carry a PFam32 gene. Obviously, this nomenclature scheme has the mild disadvantage that the numbers in new plasmid names will often not accurately reflect their sizes (for example, the plasmids with strain B31 lp38 type PFam32 genes in strains JD1 and 297 are actually about 24 kbp in length). Nonetheless, we feel that the benefits of such a scheme (names that systematically convey information about the replication, partitioning, and compatibility properties of the plasmid) outweigh these minor points. We also note that, when Borrelia genomes are sequenced, the exact lengths of the linear replicons are not automatically known, since terminal hairpin-ended DNA fragments cannot ligate to both sides of the opening in the circular plasmid vector DNA and so are not present in the circular plasmid-borne DNA libraries used for bulk sequencing [2, 36, 77, 78]. Thus, although their size can be estimated in electrophoresis gels, the lengths of even “sequenced” linear replicons in the size range of the Borrelia plasmids are usually not known to better than about ±1 kbp, unless labor-intensive, directed measures are taken to determine the sequences out to the tips of the linear replicons [1]. In such a PFam32-based nomenclature scheme, one could either give the plasmids completely new names or consider the B31 plasmids as “reference plasmids” and use them as the basis for future names. The Lyme disease research community has become accustomed to using the strain B31 plasmids in the five to eight years since their publication. Thus, we suggest that the second option be utilized. Therefore, the three new PFam types (putative specificities) found in this report were named “lp28-5,” “lp28-6,” and “lp28-7” (Table 5 and Figure 3). These are all linear plasmids in the 27 to 32 kbp range, and their sizes vary among the four strains when they are present (this variation in size is present in the previously known B31-based plasmids as well—see lp28-4 in Table 5, for example). Rather than name these new specificities according to the size of the founding member, we chose to retain the “lp28” names to indicate that these plasmids have the same general relationships as the four B31 “lp28” plasmids, namely, that they are on average in the same size range and they carry largely genes from the same set of paralogous gene families as the B31 “lp28” plasmids.

Table 5. Variation in size and new plasmid types in four B. burgdorferi strains. Plasmid

B31 1

N40

JD1

297

lp28-4

27

30

32

31

lp28-5

2

-

28

27

29

lp28-6

-

-

27

27

lp28-7

-

-

31

-

37

31

24

24

lp36 1

2

Plasmid sizes in kbp. The measurements are from Southern analysis of DNAs displayed by pulsedfield electrophoresis gels [79]. “-” indicates the plasmid is not present in the indicated isolate.

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6. Summary We have determined draft sequences for B. burgdorferi isolates N40, JD1, and 297. The comparison of these draft sequences increases our understanding of a number of aspects of their biology including plasmid diversity, lipoprotein variability, and plasmid partitioning. As these sequences are finished into “complete genome sequences,” additional, more precise information on plasmid organizational relationships and total gene content variability will certainly be useful in the study of the molecular basis of Lyme disease.

Acknowledgements This work was supported by a grant from the Lyme Disease Association, Inc. (Jackson, NJ) and by grants AI37256 (BJL), GM60654 (WGQ) and AI49003 (SRC) from the National Institutes of Health.

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[38] G. Glockner, R. Lehmann, A. Romualdi, S. Pradella, U. Schulte-Spechtel, M. Schilhabel, B. Wilske, J. Suhnel & M. Platzer. (2004) Comparative analysis of the Borrelia garinii genome. Nucleic Acids Res 32, 6038í6046. [39] B. Stevenson, S. Casjens, R. van Vugt, S. F. Porcella, K. Tilly, J. L. Bono & P. Rosa. (1997) Characterization of cp18, a naturally truncated member of the cp32 family of Borrelia burgdorferi plasmids. J Bacteriol 179, 4285í4291. [40] M. J. Caimano, X. Yang, T. G. Popova, M. L. Clawson, D. R. Akins, M. V. Norgard & J. D. Radolf. (2000) Molecular and evolutionary characterization of the cp32/18 family of supercoiled plasmids in Borrelia burgdorferi 297. Infect Immun 68, 1574í1586. [41] D. R. Akins, M. J. Caimano, X. Yang, F. Cerna, M. V. Norgard & J. D. Radolf. (1999) Molecular and evolutionary analysis of Borrelia burgdorferi 297 circular plasmid-encoded lipoproteins with OspEand OspF-like leader peptides. Infect Immun 67, 1526í1532. [42] B. Stevenson & J. C. Miller. (2003) Intra- and interbacterial genetic exchange of Lyme disease spirochete erp genes generates sequence identity amidst diversity. J Mol Evol 57, 309í324. [43] X. Yang, T. G. Popova, K. E. Hagman, S. K. Wikel, G. B. Schoeler, M. J. Caimano, J. D. Radolf & M. V. Norgard. (1999) Identification, characterization, and expression of three new members of the Borrelia burgdorferi Mlp (2.9) lipoprotein gene family. Infect Immun 67, 6008í6018. [44] K. Tilly, S. Casjens, B. Stevenson, J. L. Bono, D. S. Samuels, D. Hogan & P. Rosa. (1997) The Borrelia burgdorferi circular plasmid cp26: conservation of plasmid structure and targeted inactivation of the ospC gene. Mol Microbiol 25, 361í373. [45] R. Byram, P. E. Stewart & P. Rosa. (2004) The essential nature of the ubiquitous 26-kilobase circular replicon of Borrelia burgdorferi. J Bacteriol 186, 3561í3569. [46] P. E. Stewart, R. Thalken, J. L. Bono & P. Rosa. (2001) Isolation of a circular plasmid region sufficient for autonomous replication and transformation of infectious Borrelia burgdorferi. Mol Microbiol 39, 714í721. [47] J. J. Dunn, S. R. Buchstein, L. L. Butler, S. Fisenne, D. S. Polin, B. N. Lade & B. J. Luft. (1994) Complete nucleotide sequence of a circular plasmid from the Lyme disease spirochete, Borrelia burgdorferi. J Bacteriol 176, 2706í2717. [48] B. Stevenson, K. Tilly & P. A. Rosa. (1996) A family of genes located on four separate 32-kilobase circular plasmids in Borrelia burgdorferi B31. J Bacteriol 178, 3508í3516. [49] B. Stevenson, W. Zückert & D. Akins. (2000) Repetition, conservation, and variation: the multiple cp32 plasmids of Borrelia species. J. Molec. Microbiol. Biotech., in press. [50] W. R. Zuckert & J. Meyer. (1996) Circular and linear plasmids of Lyme disease spirochetes have extensive homology: characterization of a repeated DNA element. J Bacteriol 178, 2287í2298. [51] G. Seinost, D. E. Dykhuizen, R. J. Dattwyler, W. T. Golde, J. J. Dunn, I. N. Wang, G. P. Wormser, M. E. Schriefer & B. J. Luft. (1999) Four clones of Borrelia burgdorferi sensu stricto cause invasive infection in humans. Infect Immun 67, 3518í3524. [52] M. Theisen, B. Frederiksen, A. M. Lebech, J. Vuust & K. Hansen. (1993) Polymorphism in ospC gene of Borrelia burgdorferi and immunoreactivity of OspC protein: implications for taxonomy and for use of OspC protein as a diagnostic antigen. J Clin Microbiol 31, 2570í2576. [53] M. Y. Alghaferi, J. M. Anderson, J. Park, P. G. Auwaerter, J. N. Aucott, D. E. Norris & J. S. Dumler. (2005) Borrelia burgdorferi ospC heterogeneity among human and murine isolates from a defined region of northern Maryland and southern Pennsylvania: lack of correlation with invasive and noninvasive genotypes. J Clin Microbiol 43, 1879í1884. [54] T. Lin, J. H. Oliver, Jr. & L. Gao. (2002) Genetic diversity of the outer surface protein C gene of southern Borrelia isolates and its possible epidemiological, clinical, and pathogenetic implications. J Clin Microbiol 40, 2572í2583. [55] W. G. Qiu, D. E. Dykhuizen, M. S. Acosta & B. J. Luft. (2002) Geographic uniformity of the Lyme disease spirochete (Borrelia burgdorferi) and its shared history with tick vector (Ixodes scapularis) in the Northeastern United States. Genetics 160, 833í849. [56] S. Casjens. (1999) Evolution of the linear DNA replicons of the Borrelia spirochetes. Current Opinion in Microbiology 2, 529í534. [57] S. E. Douglas. (1994) DNA Strider. A Macintosh program for handling protein and nucleic acid sequences. Methods Mol Biol 25, 181í194. [58] C. I. Champion, D. R. Blanco, J. T. Skare, D. A. Haake, M. Giladi, D. Foley, J. N. Miller & M. A. Lovett. (1994) A 9.0-kilobase-pair circular plasmid of Borrelia burgdorferi encodes an exported protein: evidence for expression only during infection. Infect Immun 62, 2653í2661. [59] J. C. Miller & B. Stevenson. (2003) Immunological and genetic characterization of Borrelia burgdorferi BapA and EppA proteins. Microbiology 149, 1113í1125.

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[60] J. T. Skare, D. M. Foley, S. R. Hernandez, D. C. Moore, D. R. Blanco, J. N. Miller & M. A. Lovett. (1999) Cloning and molecular characterization of plasmid-encoded antigens of Borrelia burgdorferi. Infect Immun 67, 4407í4417. [61] S. Feng, S. Das, T. Lam, R. A. Flavell & E. Fikrig. (1995) A 55-kilodalton antigen encoded by a gene on a Borrelia burgdorferi 49-kilobase plasmid is recognized by antibodies in sera from patients with Lyme disease. Infect Immun 63, 3459í3466. [62] J. R. Zhang, J. M. Hardham, A. G. Barbour & S. J. Norris. (1997) Antigenic variation in Lyme disease borreliae by promiscuous recombination of VMP-like sequence cassettes. Cell 89, 275í-285. [63] J. R. Zhang & S. J. Norris. (1998) Genetic variation of the Borrelia burgdorferi gene vlsE involves cassette-specific, segmental gene conversion. Infect Immun 66, 3698í3704. [64] T. A. Leonard, J. Moller-Jensen & J. Lowe. (2005) Towards understanding the molecular basis of bacterial DNA segregation. Philos Trans R Soc Lond B Biol Sci 360, 523í-535. [65] J. Garcia-Lara, M. Picardeau, B. J. Hinnebusch, W. M. Huang & S. Casjens. (2000) The role of genomics in approaching the study of Borrelia DNA replication. J Mol Microbiol Biotechnol 2, 447í454. [66] C. Beaurepaire & G. Chaconas. (2005) Mapping of essential replication functions of the linear plasmid lp17 of B. burgdorferi by targeted deletion walking. Mol Microbiol 57, 132í142. [67] C. H. Eggers, M. J. Caimano, M. L. Clawson, W. G. Miller, D. S. Samuels & J. D. Radolf. (2002) Identification of loci critical for replication and compatibility of a Borrelia burgdorferi cp32 plasmid and use of a cp32-based shuttle vector for the expression of fluorescent reporters in the Lyme disease spirochaete. Mol Microbiol 43, 281í295. [68] D. Grimm, A. F. Elias, K. Tilly & P. A. Rosa. (2003) Plasmid stability during in vitro propagation of Borrelia burgdorferi assessed at a clonal level. Infect Immun 71, 3138í-3145. [69] D. Grimm, C. H. Eggers, M. J. Caimano, K. Tilly, P. E. Stewart, A. F. Elias, J. D. Radolf & P. A. Rosa. (2004) Experimental assessment of the roles of linear plasmids lp25 and lp28-1 of Borrelia burgdorferi throughout the infectious cycle. Infect Immun 72, 5938í5946. [70] P. E. Stewart, G. Chaconas & P. Rosa. (2003) Conservation of plasmid maintenance functions between linear and circular plasmids in Borrelia burgdorferi. J Bacteriol 185, 3202í3209. [71] C. H. Eggers & D. S. Samuels. (1999) Molecular evidence for a new bacteriophage of Borrelia burgdorferi. J Bacteriol 181, 7308í7313. [72] C. H. Eggers, B. J. Kimmel, J. L. Bono, A. F. Elias, P. Rosa & D. S. Samuels. (2001) Transduction by IBB-1, a bacteriophage of Borrelia burgdorferi. J Bacteriol 183, 4771í4778. [73] C. H. Eggers, S. Casjens & D. S. Samuels. (2001). Bacteriophages of Borrelia burgdorferi and other spirochetes. In The Spirochetes: Moelcuar and Cellular Biology (Saier, M. & Garcia-Lara, G., eds.), pp. 35í-44. Horizon Scientific Press, Wymondham, U.K. [74] C. Ojaimi, C. Brooks, S. Casjens, P. Rosa, A. Elias, A. Barbour, A. Jasinskas, J. Benach, L. Katona, J. Radolf, M. Caimano, J. Skare, K. Swingle, D. Akins & I. Schwartz. (2003) Profiling of temperatureinduced changes in Borrelia burgdorferi gene expression by using whole genome arrays. Infect Immun 71, 1689í1705. [75] F. Jeanmougin, J. D. Thompson, M. Gouy, D. G. Higgins & T. J. Gibson. (1998) Multiple sequence alignment with Clustal X. Trends Biochem. Sci. 23, 403í405. [76] J. D. Thompson, D. G. Higgins & T. J. Gibson. (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 22, 4673í4680. [77] J. Hinnebusch, S. Bergstrom & A. G. Barbour. (1990) Cloning and sequence analysis of linear plasmid telomeres of the bacterium Borrelia burgdorferi. Mol Microbiol 4, 811í820. [78] J. Hinnebusch & A. G. Barbour. (1991) Linear plasmids of Borrelia burgdorferi have a telomeric structure and sequence similar to those of a eukaryotic virus. J Bacteriol 173, 7233í7239. [79] S. Casjens & W. M. Huang. (1993) Linear chromosomal physical and genetic map of Borrelia burgdorferi, the Lyme disease agent. Mol Microbiol 8, 967í-980.

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Treponema Genomics George M. WEINSTOCK a,b,1 , David ŠMAJS c, Petra MATƞJKOVÁ c, Michal STROUHAL c, Thomas J. ALBERTd, Steven J. NORRIS e, Timothy PALZKILL a,b and Erica J. SODERGREN a a Human Genome Sequencing Center, Baylor College of Medicine, One Baylor Plaza, Houston, Texas 77030, USA b Department of Molecular Virology and Microbiology, Baylor College of Medicine, One Baylor Plaza, Houston, Texas 77030, USA c Department of Biology, Faculty of Medicine, Masaryk University, Tomešova 12, 60200 Brno, Czech Republic d NimbleGen Systems Inc., 1 Science Court, Madison, Wisconsin 53711, USA e Department of Pathology and Laboratory Medicine, University of Texas Medical School at Houston, P.O. Box 20708, Houston, Texas 77030, USA Abstract. Treponemes include the agents of syphilis, yaws, and other diseases, but as a group are fastidious organisms that often can only be grown in animals. This has hampered molecular analysis, but much progress is being made since genome sequences for these bacteria have been completed. The current state of the art involves comparative genomics, microarrays, whole genome clone sets, and other tools to unravel the biology of these pathogens. Keywords. DNA sequence, comparative genomics, syphilis, yaws, microarray

Introduction Treponema pallidum subspecies pallidum is the causative agent of the sexually transmitted disease syphilis. It is a member of the spirochetes, a well-defined group of bacteria classified by their morphology. Understanding T. pallidum pathogenesis is limited by the novel properties of this bacterium. T. pallidum is an obligate human pathogen that has not been cultivated continuously under in vitro conditions [1]. The inability to culture the bacterium prevents use of standard genetic approaches and makes biochemistry challenging. Furthermore, T. pallidum is thought of as a “stealth” pathogen, since it can persist in a host for decades [2]. This capability undoubtedly is related to a relatively low protein content of the cell surface, as judged by freeze fracture microscopy and flagella that are hidden in the periplasm from the host defenses. Following the spread of syphilis throughout Europe in the 16th century, it took over 400 years for T. pallidum to be identified as the syphilis agent. In 1905 Schaudinn 1 Corresponding Author: George M. Weinstock, Human Genome Sequencing Center Baylor College of Medicine, One Baylor Plaza, Houston, TX, USA; E-mail: [email protected]

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and Hoffman made this identification, and 2005 marked the centenary of this milestone [3]. The genome sequence of T. pallidum [4] was one of the first bacteria to be completed and greatly enhanced analysis of T. pallidum’s capabilities. The exclusive human pathogenicity and growth dependence were found to derive from a relatively small chromosome (1.14 Mbp) containing 1039 predicted ORFs. Examination of the sequence suggested potential virulence factors of T. pallidum [5, 6]. At least 70 genes (including genes for potential cell surface proteins) are candidates for possible roles in pathogenesis of T. pallidum infection. Considering that about 40% of the T. pallidum genes do not have homology with genes of known functions, there are likely to be many more undiscovered virulence factors. These genes are treponeme-specific and may give the organism its unique properties. Learning about these novel genes is one of the major challenges in furthering our understanding of this microbe. One of the major findings from the genome sequence was the tpr genes. This family of a dozen genes only showed sequence similarity to another treponemal gene, the major sheath/surface protein of T. denticola [7, 8]. Thus these genes were in the treponeme-specific class and, given that msp had virulence properties, were likely virulence factors themselves.

1. Post-Sequencing Genomic Analysis Many bacterial genome projects follow a standard pattern. For pathogens, the series of activities is listed below: 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Sequence genome. Predict all genes. Annotate/compare genes to databases. Clone all genes. Produce all proteins. Identify antigens for vaccines and diagnostics. Make microarrays. Make knockout mutations. Make interaction maps. Compare genomes.

The initial description of the genome sequence and subsequent analyses [4–6, 9] included steps 1, 2, and 3. The construction of a complete clone set (step 4) was accomplished [10] when over 90% of the T. pallidum genes were individually cloned into expression vectors in E. coli. Demonstration of expression of these genes (step 5) was by fusion of the treponemal gene to the reporter gst and detection of the resulting hybrid proteins with GST antibody. This clone set was valuable for a number of purposes: antigen identification, a convenient source of genes for microarrays, functional annotation of polypeptides (e.g., by identifying affinity ligands with phage display technology), and for protein purification. The entire set of about 900 clones expressed in E. coli was screened for antigens using antisera from T. pallidum infected rabbits [11]. One hundred and six antigens were identified, of which 85 had not been previously described. This gene-by-gene method of finding protein antigens is much more sensitive than the traditional

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approaches of making shotgun libraries in high copy number expression vectors and then doing colony immunoblots. That approach suffers from a number of biases, including overrepresentation of the most immunoreactive clones, cloning biases when proteins are toxic to the host, and the need to build very large clone libraries to obtain good sampling of the genome. The clone set approach used for T. pallidum, on the other hand, looks at each gene separately, providing maximum sensitivity with minimum bias. Two other observations are of interest. About one-half of the clones are likely exported proteins based on their sequences, and this enrichment for noncytoplasmic proteins indicates that the immune response is targeting proteins on the cell surface. Another observation is that different sets of antigens react when rabbit sera from different times after infection are used. Thus there is a temporal pattern of expression, immunogenicity, or host response. When human syphilitic sera were used, a distinct set of antigens was likewise observed [12]. The clone set was also used to build a microarray of the full set of T. pallidum genes [13]. This demonstrated that all predicted genes do in fact give rise to transcripts. Thus even though a large fraction of the predicted genes do not match sequences in databases, this is not because they are not real genes. The next step, making knockout mutations, is not feasible with T. pallidum since the bacterium cannot be grown outside of animals. Studies of interactions (step 9) are in progress (T. Palzkill, unpublished). However, much work is ongoing in the final step, comparative genomics.

2. Comparative Genomics of Treponemes Since the first insight into T. pallidum genomics came from comparison with the msp gene of T. denticola, it was of interest to delve more deeply into this genome. T. denticola is an oral spirochete that has been associated with periodontal disease. It can be cultured in vitro and has been used for biochemical studies, genetics (knockout mutants can be constructed), and studies of its interaction with host cells. The sequence of T. denticola was determined and analyzed [14] in comparison to T. pallidum. While the two bacteria are both spirochetes, much of their genomic structure is different. The T. denticola genome is over twice the size of T. pallidum and has a very different base composition (much more AT rich). There is almost no conservation of gene order, and the two chromosomes do not align to any appreciable extent. The extra genes of T. denticola presumably allow it to be less fastidious and compete with the extensive microbial population in the oral cavity.

Organism

Disease

T. pallidum subsp. pallidum

Venereal syphilis

T. pallidum subsp. pertenue

Yaws

T. pallidum subsp. endemicum

Endemic syphilis (bejel)

T. carateum

Pinta (not yet grown in animals)

T. paraluiscuniculi

Venereal spirochetosis (rabbits)

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Another important comparison is between other T. pallidum subspecies and closely related species. The syphilis strain is T. pallidum subsp. pallidum. But there are a number of other subspecies, listed in the Table. Diseases caused by these organisms are quite distinct, yet at the DNA level these organisms are nearly indistinguishable, being >95% identical based on hybridization. One hopes that in the future, comparison of these sequences will shed light on which virulence factors are responsible for the unique host ranges and infection phenotypes. In an initial study, using restriction enzyme digestion of segments of the genome produced by long-range PCR as the basis for comparison, only four differences were found between the syphilis and yaws bacteria, and each of these were only single genes (E. Sodergren et al., unpublished). This result is consistent with the high degree of conservation seen by hybridization and suggests that the phenotypic differences may reside at the single nucleotide level. Moreover, all four differences were in or adjacent to tpr genes, fortifying the notion that these are changes affecting virulence genes that give the distinctive treponemal infections. Similar results were found in studies of other treponemes in the table (M. Strouhal et al., unpublished). To probe the differences more deeply, hybridization to oligonucleotide arrays was performed. In this case the oligonucleotide array was constructed at NimbleGen and comprised 29-mers spaced seven nucleotides apart along the genome. Comparison of two syphilis strains showed slightly more than 300 single nucleotide polymorphisms (SNPs), while comparison of syphilis to yaws bacteria showed of the order of 1000 SNPs and at least several times this number were found in comparing the human and rabbit syphilis strains (P. Matejkova et al., unpublished). For a sense of scale, the syphilis-yaws comparison indicates about 1 SNP per gene, yet only a subset of these are expected to cause nonconservative changes in protein structure, so only a fraction of the genes are affected.

3. Conclusions In the century since T. pallidum was identified there have been notable successes in the struggle to control syphilis and understand this bacterium. Notable among these are the invention of the first antimicrobial treatments, such as Salvarsan, by Paul Ehrlich, the discovery of the exquisite sensitivity of T. pallidum to penicillin, and even a Nobel Prize in 1927 for treatment of advanced syphilis by infection with Plasmodium. However, the genome sequence published in 1998 ushered in a whole new era, the early stages of which are described here. We can certainly expect the future to hold further discoveries about not only syphilis and T. pallidum, but also the other fascinating close relatives.

References [1] [2] [3] [4]

Cox, D.L. Culture of Treponema pallidum. Methods Enzymol, 1994. 236: 390–405. Norris, S.J., et al. Treponema and other human host-associated spirochetes, in Manual of Clinical Microbiology, P.R. Murray, Editor. 2003, ASM Press: Washington, D.C., Chapter 61. Waugh, M. The centenary of Treponema pallidum: on the discovery of Spirochaeta pallida. Skinmed, 2005. 4(5): 313–315. Fraser, C.M., et al. Complete genome sequence of Treponema pallidum, the syphilis spirochete. Science, 1998. 281(5375): 375–388.

100 [5] [6] [7] [8] [9] [10] [11] [12] [13] [14]

G.M. Weinstock et al. / Treponema Genomics Weinstock, G.M., et al. The genome of Treponema pallidum: new light on the agent of syphilis. FEMS Microbiol Rev, 1998. 22(4): 323–332. Weinstock, G.M., et al. Identification of virulence genes in silico: infectious disease genomics, in Virulence Mechanisms of Bacterial Pathogens, K.A. Brogden, et al., Editor. 2000, ASM Press: Washington, D.C., 251–-261. Mathers, D.A., et al. The major surface protein complex of Treponema denticola depolarizes and induces ion channels in HeLa cell membranes. Infect Immun, 1996. 64(8): 2904–2910. Fenno, J.C., K.H. Muller, and B.C. McBride Sequence analysis, expression, and binding activity of recombinant major outer sheath protein (Msp) of Treponema denticola. J Bacteriol, 1996. 178(9): 2489–2497. Norris, S.J., D.L. Cox, and G.M. Weinstock. Biology of Treponema pallidum: correlation of functional activities with genome sequence data. J Mol Microbiol Biotechnol, 2001. 3(1): 37–62. McKevitt, M., et al. Systematic cloning of Treponema pallidum open reading frames for protein expression and antigen discovery. Genome Res, 2003. 13(7): 1665–1674. McKevitt, M., et al. Genome scale identification of Treponema pallidum antigens. Infect Immun, 2005. 73(7): 4445–4450. Brinkman, M.B., et al. Reactivity of antibodies from syphilis patients to a protein array representing the Treponema pallidum proteome. J Clin Microbiol, 2006. 44(3): 888–891. Smajs, D., et al. Transcriptome of Treponema pallidum: gene expression profile during experimental rabbit infection. J Bacteriol, 2005. 187(5): 1866–1874. Seshadri, R., et al. Comparison of the genome of the oral pathogen Treponema denticola with other spirochete genomes. Proc Natl Acad Sci U S A, 2004. 101(15): 5646–5651.

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Comparative Analysis of Pathogenic Leptospira Genomes Richard L. ZUERNER a,*,1 , Dieter M. BULACH b,c,*, Torsten SEEMANN c, Ross L. COPPELb,c,d and Ben ADLER b,c,d a Bacterial Diseases of Livestock Research Unit, National Animal Disease Center, USDA Agricultural Research Service, Ames, IA 50010, USA b Australian Bacterial Pathogenesis Program, Department of Microbiology, Monash University, VIC 3800, Australia c Victorian Bioinformatics Consortium, Monash University, VIC 3800, Australia d Australian Research Council Centre of Excellence in Structural and Functional Microbial Genomics, Department of Microbiology, Monash University, VIC 3800, Australia Abstract. The presence of two chromosomes makes Leptospira unusual amongst its closest relatives in the bacterial world. The Leptospira genome is in a state of flux, as indicated by the presence of many chromosomal rearrangements that alter genetic organization between individual serovars. It is therefore somewhat remarkable that at least two Leptospira loci (LPS biosynthetic genes and the S10spc-alpha ribosomal protein operon) form large, extended operons that are among the longest bacterial operons reported to date. Insertion sequences (IS) that are distributed throughout Leptospira genomes contribute to the formation of rearrangements. These elements can transpose and disrupt the integrity of genes, or alternatively, can activate cryptic genes by providing promoter activity to genomic sequences downstream of the insertion site. Bioinformatics and experimental functional analyses were used to characterize the L. interrogans genomes and thus gain insight into this organism’s biology. Quantitative analysis of the L. interrogans serovars Lai and Copenhageni genomes showed that these bacteria are proficient in environmental sensing and response, and in nutrient transport. These data support epidemiological evidence that L. interrogans is transmitted primarily by passage through environmental sources. Few pseudogenes were detected in either strain, suggesting that there is sufficient selective pressure to maintain a highly functional genome. However, several genes were identified that are complete in one strain but have frameshifts in the other that may affect phenotype. Further differences in phenotype may also result from gene acquisition, and we found several large, serovar-specific gene clusters. Analysis of an ECF locus from L. interrogans serovar Pomona is used to show how RT-PCR and expression vectors can be used to localize promoters in L. interrogans. Antisera produced against recombinant fusion proteins were used to detect invasion of lung, liver, and kidney during experimental infection of hamsters with serovar Pomona. These data are consistent with some of the clinical manifestations of severe leptospirosis and

1 Corresponding Author: Richard L. Zuerner, Bacterial Diseases of Livestock Research Unit, National Animal Disease Center, USDA Agricultural Research Service, Ames, IA 50010, USA. * Denotes first co-authors.

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R.L. Zuerner et al. / Comparative Analysis of Pathogenic Leptospira Genomes help to illustrate how genomic analysis can aid in the understanding of these pathogenic bacteria. Keywords. Leptospira, genomics, insertion sequences, recombinant proteins

Introduction Leptospirosis is one of the most common and widespread zoonotic diseases known [1]. This disease is caused by pathogenic species of Leptospira; most mammalian species are susceptible to infection [2]. Infection occurs through exposure of contaminated body fluids, principally urine from infected animals, enabling the bacteria to gain access through mucous membranes or abrasions. Following dissemination through the blood, Leptospira concentrates in liver and kidney. Release of viable bacteria through the urine is an important part of the transmission process, leading to infection of other animals by either direct exposure, or by contamination of water sources that are then a broader source of infection [2]. Periodic epidemics of human leptospirosis coincide with seasonal flooding in urban areas and are principally associated with environmental contamination from urine originating from chronically infected maintenance hosts [3]. Leptospirosis is manifested in one of two forms, as either a chronic infection with low mortality or an acute infection with high mortality [2]. Chronic Leptospira infections in maintenance host species result in little apparent disease in adults but can induce reproductive failure (abortion, stillbirth, and weak offspring) during gestation. Normal maintenance hosts are a persistent source of infection for wildlife, domesticated species, and humans. For example, rats chronically infected with L. interrogans are one of the most common sources of environmental contamination leading to human leptospirosis outbreaks during urban flooding [3]. Chronic Leptospira infections in livestock, especially cattle, can result in economic losses due to reduced herd vitality and lost milk production, and may be a source of health risk for individuals in the livestock industry. Accidental exposure of non-maintenance hosts to contaminated fluids, either directly or from environmental sources, can result in acute infection. The clinical symptoms resulting from accidental exposure to pathogenic Leptospira can range from a mild, influenza-like disease to an acute, severe infection resulting in death from multiple organ failure. The same strains that cause chronic infections in their normal maintenance hosts can cause severe infections in nonmaintenance hosts [2]. Progress in analyzing pathogenic Leptospira has been slow for several reasons. Leptospira is fastidious, and these bacteria utilize fatty acids and fatty alcohols as carbon and energy sources. Although some strains can grow in a defined, protein-free medium, protein, in the form of bovine serum albumin, is often needed to bind free fatty acids to overcome toxicity [4]. Leptospira grows slowly, and primary isolation from clinical samples is often difficult. Few tools for genetic manipulation of pathogenic strains are available and are limited to transposon mutagenesis [5, 6]. Methodology enabling the generation of gene-specific mutations is still lacking, and there are no methods for genetic complementation. Thus, genetic analysis of pathogenic Leptospira has relied on gene isolation and sequencing, and more recently, genomic analysis. In this paper, we discuss developments in the genomic analysis of pathogenic Leptospira leading to new information on the biology of these bacteria.

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1. Genetic Organization Although Leptospira shares a common ancestry with Brachyspira, Borrelia, and Treponema, genetic organization among these bacteria is quite different. The L. interrogans genome is nearly 5 Mbp in length and is composed of two circular chromosomal replicons present at nearly equal molar ratios [7]. Early studies suggested that the larger chromosomal replicon (CI) resembled a typical bacterial chromosome, containing all copies of ribosomal rRNA genes and a typical replication origin [8, 9]. The small chromosomal replicon (CII) is a constant feature among Leptospira, and these early studies showed the presence of a unique copy of asd, a gene essential for cell wall biosynthesis [8]. Further characterization of both replicons lagged until genomic sequencing data for two L. interrogans serovars, Lai and Copenhageni, were reported [10, 11]. Through genomic sequence analysis, we now have a much better and detailed understanding of the genetic content of L. interrogans; these data provide a foundation for identifying virulence traits, antigenic proteins, and metabolic potential that previous mapping studies could not generate. Genomic sequence analysis revealed that all tRNA and most housekeeping functions are encoded on the CI replicon [10, 11]. As with many other bacteria, the Leptospira replication origin coincides with a sharp deflection in the GC content of the leading strand, also known as GC skew. Although there is a sharp deflection in the GC skew in the CII replicon, the genetic content surrounding this region is different from that of CI. Instead of genes typical of chromosomal replication origins (for example, dnaA and gidA) on CII, the GC skew is located adjacent to parA and parB, two genes associated with plasmid and chromosomal partitioning. This finding suggests that L. interrogans utilizes the parAB gene system for segregation of the small chromosome.

2. Genomic Analysis To provide an accurate and detailed comparison of the L. interrogans serovar Lai and Copenhageni genome sequences, we modified the annotation by applying consistent criteria for all encoded proteins, including determining the consensus amino terminus through BLASTP analysis, and removed several small putative coding sequences (CDS) that lacked credible upstream translation initiation sequences. By revising the annotation of these two genomes, we established a format useful for comparison of additional Leptospira genome sequences as they become available. Furthermore, this process created a platform for consistent annotation of Leptospira genome sequences, making comparative analysis easier and more accurate. The revised annotation of both strains is summarized in Table 1. The CDS line in this table shows what we define as the “functional genome” in comparison to the total coding potential of these bacteria (that is, no pseudogenes, gene fragments, or transposable elements). The L. interrogans genome contains a small proportion (2–3%) of pseudogenes. By comparison, many pathogenic bacteria have a significantly higher proportion of pseudogenes. This finding suggests that there is strong selective pressure on L. interrogans to maintain gene integrity. Both strains have IS elements distributed around the CI replicon; serovar Lai also has IS elements on CII [10, 11]. Based on hybridization analysis, the L. interrogans serovar Pomona genome has more copies of

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Table 1. Essential features of the L. interrogans genome. Serovar Copenhageni

Serovar Lai

Feature Size (Kb)

CI

CII

CI

CII

4,280

350

4,330

359

Protein coding sequences CDS 1

3,110

274

3,114

272

With assigned function

1,813

159

1,799

157

Conserved hypothetical

496

35

497

34

Unique hypothetical

801

80

818

81

Pseudogenes

65

5

109

17

Transposases

76

0

90

11

3,251

279

3,313

300

37

0

37

0

2

0

1

0

Total Transfer RNA genes Ribosomal RNA genes 23S

1

16S

2

0

2

0

5S

1

0

1

0

not including transposases or pseudogenes

the elements IS1500 [12], IS1501 (data not shown), and IS1502 [9] than either serovar Lai or Copenhageni. This may contribute to the large number of rearrangements that differentiate these strains (see below). Quantitative analysis of L. interrogans CDS was done using the clusters of orthologous genes (COGs) approach [13, 14]. The L. interrogans genome encodes a large number of proteins associated with environmental sensing and response, and diverse transport functions (Figure 1). These findings are consistent with the survival requirements of bacteria that routinely pass through water or other environmental niches between mammalian hosts. Two of the largest groups in L. interrogans are the poorly characterized conserved hypothetical proteins (R and S). Few genes in either serovar are unique. In serovar Lai, there are 54 serovar-specific genes not present in serovar Copenhageni. Conversely, there are 31 serovar Copenhageni-specific genes not present in serovar Lai. In both strains, several of the serovar-specific genes are found as clusters, with some extending approximately 40 kb in length. The presence of these clustered, serovar-specific genes suggests they may have been acquired through horizontal gene transfer. In both serovars, most of the unique CDS encode hypothetical proteins, so it is unclear how they may

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Figure 1. COG analysis of the L. interrogans genome. Each segment represents the proportion of the genome encoding proteins in each COG category. General COG categories (starting at the top of the chart and going clockwise) include: information storage and processing (COG J, A, K, L, and B), cellular processes and signaling (COG D, Y, V, T, M, N, U, and O), metabolism and transport (COG C, G, E, F, H, I, P, and Q), and poorly characterized groups R and S.

affect phenotype. Copenhageni-specific genes that have putative functions include a ferrodoxin related protein, tautomerase, and an AcrR family transcription factor, while serovar-specific genes in Lai include a nucleotidyltransferase, proteic killer gene system, and a gene encoding a stability toxin. Further diversification between serovars Lai and Copenhageni is achieved through the generation of pseudogenes. There are only six pseudogenes common to both serovars, with all other CDS being intact in the other serovar. Many of the serovarspecific pseudogenes occur in CDS encoding hypothetical proteins, making it difficult to assess their potential role in altering phenotype. However, several serovar-specific pseudogenes are in CDS encoding proteins having presumed functions based on their homology with better-characterized proteins. For example, serovar Copenhageni has frameshift mutations (pseudogenes) in genes encoding glutathione transferase, LigC, a TonB dependent receptor, and a flavoprotein. Pseudogenes in serovar Lai are found in CDS encoding a TPR-repeat protein, a Na+/H+ antiporter, a phospholipase D like lipoprotein, an adenylate-guanylate cyclase, and a phospholipid synthase. Mutations in some of these genes, for example, mutation of the adenylate-guanylate cyclase may have broad impact on gene expression, as this group of proteins participates in global gene regulation.

3. Genetic Rearrangements Several studies have shown that genetic organization in L. interrogans is quite variable. The larger (CI) replicon undergoes extensive rearrangements (inversions, deletions, and

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Figure 2. The CI replicons from L. interrogans serovars Pomona and Icterohaemorrhagiae are represented as horizontal lines. The lines drawn between the two maps denote the relative position of identical genes. Lines that intersect or diverge indicate rearrangements between the two chromosomal replicons. The sigE locus is localized in serovar Pomona near rrlB.

insertions), which dramatically alter global genetic organization of closely related strains [8]. Figure 2 is a comparison of the genetic maps of two serovars of L. interrogans showing the presence of extensive rearrangements throughout the CI replicon. This propensity for rearrangement is likely to be limited by viability, given the absence of rearrangements in two key loci. Specifically, there is no evidence of rearrangement in the LPS [15] and S10-spc-alpha [16] loci, each forming large operons that generate some of the longest bacterial transcripts known. Studies comparing the restriction endonuclease digestion patterns of L. interrogans serovar Pomona isolates showed the presence of several polymorphisms that may affect important phenotypic characteristics including antigenicity, or adaptation to a particular host species [17]. DNA hybridization studies indicate that serovars Icterohaemorrhagiae and Pomona share about 84% similarity in genetic composition [18] and have therefore undergone considerable divergence. In contrast, the two L. interrogans strains for which genomic sequencing has been completed, serovars Lai and Copenhageni, share about 98% of their genetic composition [10, 11, 18]. Comparison of the CI replicons from L. interrogans serovars Lai and Copenhageni reveal the presence of a large inversion (Figure 3A) that coincides with a rearrangement between two copies of an IS element [10]. The role of IS elements in generating chromosomal rearrangements in L. interrogans was first suggested in a study describing IS1500 [12]. Localization of some IS1500 insertions showed these often coincide with regions of the genome that have undergone large rearrangements [12]. These data are also consistent with studies that

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Figure 3. Linear representation of the genomic sequences, both CI (A) and CII (B), of L. interrogans serovars Lai (top) and Copenhageni (bottom). Regions of sequence similarity were identified by using Megablast and visualized using ACT. Black lines denote regions of highly similar sequence between the map lines drawn in grey.

have used IS elements as epidemiological tools to identify serovars [17]. In contrast to the variability seen in the CI replicon, genetic organization in CII appears to be more stable (Figure 3B), and recombination between the CI and CII replicons is not apparent.

4. Insertion Sequences The role of IS elements in altering chromosome organization in Leptospira spp. has been suggested by several studies. IS elements are a group of elements that can transpose to new sites in the genome [19]. Many IS elements follow a classic structural organization, consisting of a central “unique” region that is often flanked by terminal inverted repeats. This central region often contains one or more genes that encode proteins by catalyzing the transposition of the element [19]. Most IS elements generate a small duplication at the integration site. This sequence duplication results from enzymatic repair to staggered ends formed at the insertion site during transposition. Insertion of an IS element into a gene usually inactivates that gene and can often cause polar mutations within operons by disrupting transcription of genes located downstream of the insertion site [19]. Some IS elements can promote transcription of sequences adjacent to the insertion site, thereby activating cryptic genes [20]. Insertion elements may also influence genetic organization by providing targets for homologous recombination. Several IS elements from Leptospira spp. have been described, including IS1500 [12], IS1501 [21], IS1502 [9], IS1533 [22], and an IS5-like element [23]. Identification of the IS3-like element, IS1500, resulted in the development of a useful tool for epidemiological studies [17]. Variation in the hybridization patterns or PCR products using IS1500-based probes suggests that these elements are active and can transpose in L. interrogans. We recently demonstrated the spontaneous transposition of IS1501 during analysis of in vitro selected antigenic variants of L. interrogans serovar Pomona [21]. In that study, novel transpositions were discovered in an LPS biosynthetic gene,

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Figure 4. Transposition of IS1501 into the LPS biosythesis locus of L. interrogans serovar Pomona. Antigenic variants of L. interrogans serovar Pomona were selected in vitro and novel copies of IS1501 were identified. A common insertion in OrfP35 (black arrow), a gene near the 5' end of the LPS locus was identified in both variants. The insertion site is shown as an open box in the parent locus, and the novel insertion of IS1501 is shown as a grey arrow in the mutant locus. Sequences at the parent and mutant loci are shown with the 3-bp duplication highlighted in uppercase letters.

and alterations in the LPS were detected. IS1501 generates a 3-bp duplication of the target site during transposition (Figure 4). Transcriptional analysis at two of the insertion sites showed that IS1501 directs transcription into adjacent downstream sequences. Therefore, this element is capable of acting as a mobile promoter. Because of this promoter activity, IS1501 can disrupt the coding region of a gene without inducing a downstream polar mutation.

5. Functional Genomics To illustrate the application of genomic data, we describe two studies whereby genes identified through genome sequencing formed the basis for further analyses leading to a better understanding of Leptospira biology. First, we describe a locus cloned from L. interrogans serovar Pomona that has homology to the ECF (extra cytoplasmic factor) family of proteins [24, 25] and may play a role in regulating cellular responses to the extracellular environment. Second, we describe studies leading to identification and characterization of the outer membrane protein LipL21 and its expression during experimental infection. These studies applied sequence data to develop strategies that would be untenable in a pre-genomics era. Many bacterial species regulate genes associated with the extracellular environment by coordinating the process of transcription initiation. Often, the promoters of these genes are recognized by alternative ı factors belonging to the ECF family [24, 25]. ECF-regulated genes can affect host-parasite interactions, and there is a growing body of literature showing that inactivation of ECF encoding genes can reduce bacterial virulence. For example, inactivation of either of two ECF-like genes of Salmonella typhimurium results in significantly reduced virulence [26, 27]. During an ongoing project in use to identify genes in L. interrogans through sequencing random clones, we identified a genetic locus that spans genes encoding an ECF-like protein and two potential transmembrane proteins. Analysis of this locus may help us analyze how L. interrogans responds to its extracellular environment to regulate gene expression.

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Figure 5. A map of the 3,659 bp HaeIII fragment from L. interrogans serovar Pomona. Key restriction sites are noted above the horizontal line representing the fragment. Closed arrows indicate genes encoded on the fragment and open boxes indicate the various plasmid inserts mentioned in the text. Note that fragment 669595 contains the promoters for both hsa and sigE.

BLAST analysis of sequences obtained from randomly picked cloned fragments of the L. interrogans serovar Pomona genome led to identification of an ECF encoding gene on plasmid pKB25. Analysis of this sequence revealed an open reading frame that encoded a novel ECF (extra cytoplasmic factor) related protein and was designated sigE. This gene corresponds to loci LA0876 and LIC12757 from L. interrogans serovars Lai and Copenhageni, respectively. Sequences flanking sigE were isolated using a PCR-based genome walking method [16], resulting in the isolation of clones 671RH and 673RH. In total, these three cloned fragments spanned a 3,659 bp HaeIII fragment containing four genes (Figure 5). Two genes occur downstream of sigE, both lacking homologs in GenBank, and were designated dshA and dshB (for downstream of igE hypothetical gene). In L. interrogans serovar Pomona, these genes may represent pseudogenes due to the presence of an inframe stop codon that is not present in orthologous genes from serovars Lai (LA0878) and Copenhageni (LIC12756). Upstream of sigE is a gene with a region of limited similarity (E value = 0.083) to the Leptospira borgpetersenii ysp1 gene, a gene adjacent to a sphingomyelinase encoding gene, sphA [28]. The L. interrogans gene is designated hsa (for homolog to sphA adjacent gene) and corresponds to LA0875 (serovar Lai) and LIC12758 (serovar Copenhageni). Analysis of L. interrogans RNA by RT-PCR showed that sigE, dshA, and dshB are co-transcribed (data not shown). This transcript may end at a potential transcription termination stem-loop structure is predicted to occur downstream of dshB. The sigE and hsa genes are separated by 62 bp and are divergently transcribed. Promoter activity for both sigE and hsa was tested in Escherichia coli by inserting a 457 bp amplicon with the sigE-hsa intergenic region and part of both coding regions upstream of the promoterless chloramphenicol acetyltransferase gene of pKK232-8 [29]. This insert, placed in both orientations in pKK232-8, conferred chloramphenicol resistance while the intact vector did not, showing that promoter activity for both genes functioned in E. coli.

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Figure 6. Comparison of the SigE protein to ECF-ı factors. Proteins were aligned with SigE and shaded either black (identical residue in all sequences), dark grey (some sequences share an identical residue with SigE), or light grey (some sequences share a similar residue with SigE). Sequences shown with accession numbers are: Lin_sigE (this work, AF143504.); Bac_sigV (Bacillus subtilis SigV, O05404); Bac_sigX (B. subtilis SigX, P35165); Bac_yhdM (B. subtilis YhdM, CAA74497); Cac_sigX (Clostridium acetobutylicum SigX, AAC12856); Pae_sigX (Pseudomonas aeruginosa SigX, AAD11567); and Rho_rpoE (Rhodobacter spharoides RpoE, AAB17906). A predicted helix-turn-helix structure is found in segment 4.2.

The SigE protein shares several motifs with ECF-ı factors (Figure 5). Like other ECF-ı factors, SigE lacks region 1, a region thought to inhibit DNA binding in the absence of core RNA polymerase [25]. The most conserved portion of ı factors, including SigE, is region 2, which is subdivided into four regions: region 2.1 may bind core RNA polymerase; region 2.2 is a conserved domain of unknown function; region 2.3 may affect DNA melting; and region 2.4 is thought to bind to the -10 sequence of ECF-regulated promoters (Figure 5) [25]. Like other ECF-ı factors, region 3 of SigE is much smaller than primary ı-factors. In primary ı-factors, region 3 residues may bind DNA and interact with the core RNA polymerase [15]. Region 4 is thought to bind –35 sequences and often has a DNA binding helix-turn-helix motif [25, 30]. SigE also has a predicted helix-turn-helix motif within region 4 (Figure 6). The Hsa and DshA proteins each have regions with predicted transmembrane domains and these proteins may be exposed to the extracytoplasmic environment. There are three potential transmembrane domains in Hsa and one in DshA. The DshB protein had no recognizable motifs that might suggest function. Alignment with homologs from L. interrogans serovars Lai and Copenhageni suggest this gene in serovar Pomona may be a pseudogene created by introduction of an in-frame stop codon. Further analysis is required to determine the cellular locations and possible functions of these proteins. The identification of an ECF-ı factor-encoding gene in L. interrogans suggested that this bacterium uses alternative sigma factors to control transcription of selected genes. This is a key finding in understanding how Leptospira control gene expression, especially in response to alterations in the extracellular environment. Moreover, considering the current understanding of fluid nature of genome layout in Leptospira, this strategy may be an effective strategy to maintain control of genes as they are moved to different locations. In the context of bacterial pathogenesis, the identification and characterization of genes controlled by ECF-ı factors in Leptospira may lead to the

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identification of novel virulence factors, given the importance of this class of genes in other pathogenic bacteria. In a second example using genomics to characterize pathogenic Leptospira, a draft sequence of L. borgpetersenii was used as a database to search for matches to peptide sequences obtained from the L. interrogans serovar Lai outer membrane [31], resulting in the identification of the LipL21 CDS [32]. PCR primers were designed that were used to amplify a portion of the LipL21 CDS leading to the construction of a Histagged fusion protein that was produced in E. coli. His-tagged LipL21 fusion protein was gel purified and used to immunize rabbits and the production of homologous

A

C

B

D

Figure 7. Detection of L. interrogans serovar Pomona in hamster tissue. Paraffin-embedded tissue from hamster lung (A), kidney (C), and liver (D) after infection with L. interrogans serovar Pomona were processed for antigen detection. Sections were incubated with anti-LipL21, followed by a secondary antibody tagged with a fluorescent dye, then illuminated with ultraviolet light. Tissue nuclei were counterstained with DAPI. Arrows point to fluorescence due to the presence of LipL21. For comparison, a section of lung stained with silver and examined by light microscopy (B) reveals the presence of L. interrogans.

antisera. Subsequent immunoblot analysis of Leptospira lysates showed that LipL21 is a major outer membrane protein that is well conserved across pathogenic Leptospira species, but not produced by saprophytic species [32]. Immunoblot analysis using patient sera and sera from experimentally infected hamsters revealed the presence of anti-LipL21 antibodies. These data suggested that LipL21 is expressed during infection. To directly test for LipL21 expression during mammalian infection, we used anti-LipL21 antisera in indirect immunofluorescence studies to analyze tissue sections from hamsters experimentally infected with L. interrogans serovar Pomona. Analysis of lung, kidney, and liver tissue showed the presence of Leptospira in all tissues by

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silver stain (Figure 6 and data not shown), and the LipL21 antisera bound to bacteria in each of these tissues, showing that this protein is expressed during infection in several different organs. The bacteria distribute differently in these tissues, being localized in tight junctions between hepatocytes, having a diffuse distribution in the kidney, or present in the alveolar space of the lung. Detection of Leptospira in the lung, in particular, is consistent with pulmonary hemorrhage, a clinical manifestation severe leptospirosis [33].

6. Conclusions Genome sequence analysis of pathogenic Leptospira is providing valuable data that can be rapidly applied to understand more about the biology of these bacteria. Accurate comparison of genomic sequence data relies on establishing common criteria for gene identification and the realization that genome annotations are dynamic data sources that need periodic revisions as new information becomes available. By revising genome annotations of L. interrogans serovars Copenhageni and Lai, we created databases by using common criteria useful for comparative analysis with existing genome sequences and established a foundation for future studies as more bacterial genomes become available. Gene discovery combined with functional analysis is making it possible to explore the biology of pathogenic Leptospira and helping to identify potential vaccine candidates and potential virulence factors. Combined, these studies will facilitate rational, technology-based development of new methods for the control and prevention of leptospirosis.

Acknowledgements We thank Paul Cullen and David Haake for the generous gift of anti-LipL21 sera, Paul Hauer for providing hamster tissues, and David Alt, Ami Frank, Richard Hornsby, and Amanda Toot for their excellent technical support throughout this project.

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(1999) Comparative analysis of the LPS biosynthetic loci of the genetic subtypes of serovar Hardjo: Leptospira interrogans subtype Hardjoprajitno and Leptospira borgpetersenii subtype Hardjobovis. FEMS Microbiol. Lett. 177, 319– 326. Kazmierczak, M.J., Wiedmann, M. and Boor, K.J. (2005) Alternative sigma factors and their roles in bacterial virulence. Microbiol. Molec. Biol. Rev. 69, 527–543. Lonetto, M.A., Brown, K.L., Rudd, K.E. and Buttner, M.J. (1994) Analysis of the Streptomyces coelicolor sigE gene reveals the existence of a subfamily of eubacterial RNA polymerase ǻ factors involved in the regulation of extracytoplasmic functions. Proc. Natl. Acad. Sci. USA 91, 7573–7577. Fang, F.C., Libby, S.J., Buchmeier, N.A., Loewen, P.C., Switala, J., Harwood, J. and Guiney, D.G. (1992) The alternative sigma factor katF (rpoS) regulates Salmonella virulence. Proc. Natl. Acad. Sci. USA 89, 11978–11982.

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R.L. Zuerner et al. / Comparative Analysis of Pathogenic Leptospira Genomes Humphreys, S., Stevenson, A., Bacon, A., Weinhardt, A.B. and Roberts, M. (1999) The alternative sigma factor, sigmaE, is critically important for the virulence of Salmonella typhimurium. Infect. Immun. 67. Segers, R.P., Drift, A.V.D., Nijs, A.D., Corcione, P., Zeijst, B.A.V.D. and Gaastra, W. (1990) Molecular analysis of a sphingomyelinase C gene from Leptospira interrogans serovar hardjo. Infect. Immun. 58, 2177–2185. Brosius, J. (1984) Plasmid vectors for the selection of promoters. Gene 27, 151–160. Lonetto, M., Gribskov, M. and Gross, C.A. (1992) The V70 Family: Sequence Conservation and Evolutionary Relationships. J. Bacteriol. 174, 3843–3849. Cullen, P.A., Cordwell, S.J., Bulach, D.M., Haake, D.A. and Adler, B. (2002) Global analysis of outer membrane proteins from Leptospira interrogans serovar Lai. Infect. Immun. 70, 2311–2318. Cullen, P.A., Haake, D.A., Bulach, D.M., Zuerner, R.L. and Adler, B. (2003) LipL21 is a novel surface-exposed lipoprotein of pathogenic Leptospira species. Infect. Immun. 71, 2414–2421. Trevejo, R.T., Rigau-Perez, J.G., Ashford, D.A., McClure, E.M., Jarquin-Gonzalez, C., Amador, J.J., de los Reyes, J.O., Gonzalez, A., Zaki, S.R., Shieh, W.J., McLean, R.G., Nasci, R.S., Weyant, R.S., Bolin, C.A., Bragg, S.L., Perkins, B.A. and Spiegel, R.A. (1998) Epidemic leptospirosis associated with pulmonary hemorrhage-Nicaragua, 1995. J. Inf. Dis. 178, 1457–1463.

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Leptospira interrogans: Genomics and “Immunomics” Ana L. T. O. NASCIMENTO 1 Centro de Biotecnologia, Instituto Butantan, São Paulo, SP, Brazil Abstract. Leptospirosis, an emerging infectious disease, is a worldwide zoonosis of human and veterinary concern. Caused by pathogenic spirochaetes of the genus Leptospira, the disease presents greater incidence in tropical and subtropical regions. Humans can be infected by exposure to chronically infected animals and their environment. The genome sequence of Leptospira interrogans serovar Copenhageni was recently reported. It contains a broad array of genes encoding for regulatory system, signal transduction, and methyl-accepting chemotaxis proteins, conforming to the organism’s ability to respond to diverse environmental stimuli. A large number of exported lipoproteins and transmembrane outer membrane proteins were identified that may be involved in leptospiral pathogenesis and protective immunity. Comparative analysis with the Leptospira interrogans serovar Lai genome revealed that, despite genetic similarity, there are structural differences, including a large chromosomal inversion. The leptospiral genome sequence, combined with bioinformatics tools, offered a unique opportunity to search for immune targets to be used for vaccine or diagnostic kit development. Out of a hundred recombinant proteins tested, sixteen were recognized by antibodies present in sera from patients diagnosed with leptospirosis and might be useful for these purposes. The most important results obtained within genome sequences, comparative genomics, and outer membrane genome-derived protein expressed in E. coli are reviewed here. Keywords. Genome, Leptospira, leptospirosis, vaccine, diagnosis, recombinant proteins

Introduction Spirochetes are motile, helically shaped bacteria, which include the genera Leptospira, Leptonema, Borrelia and Treponema. Borrelia and Treponema are the causative agents of Lyme disease, relapsing fever, and syphilis. Leptospira consists of a genetically diverse group of pathogenic and non-pathogenic or saprophytic species [1]. Leptospirosis is an emerging infectious disease of human and veterinary concern. The disease presents greater incidence in tropical and subtropical regions [1, 2]. The transmission of leptospirosis has been associated with exposure of individuals in close proximity to wild or farm animals [3]. Recently the disease became prevalent in cities 1 Corresponding author: Ana L.T.O. Nascimento, Centro de Biotecnologia, Instituto Butantan, Avenida Vital Brazil, 1500, CEP 05503-900, São Paulo, SP, Brazil. E-mail: [email protected]

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with sanitation problems and large populations of urban rodent reservoirs that contaminate the environment through their urine [4]. The incidence of leptospirosis remains underestimated in part due to the broad spectrum of signs and symptoms that patients may present. Children primarily show fever, vomiting, headache, diarrhea, and abdominal and generalized muscle pain, whereas adults have fever, headache, anorexia, muscle pain, and constipation [4, 5]; 5 to 15% of the cases evolve more severely, presenting hemorrhages with renal and hepatic failure, a condition known as Weil's syndrome [4], with a mortality rate of 5 to 40%. Leptospirosis also has a great economic impact in the agricultural industry since the disease affects livestock, inducing abortions, stillbirths, infertility, reduced milk production, and death [3, 4]. Environmental control measures are difficult to implement because of the long-term survival of pathogenic leptospires in soil and water and the abundance of wild and domestic animal reservoirs [1]. Currently available veterinary vaccines are based on inactivated whole cell or membrane preparations of pathogenic leptospires. These types of vaccine confer protective responses through induction of antibodies against leptospiral lipopolysaccharide [6]. However, these vaccines fail to induce long-term protection against infection and do not provide cross-protective immunity against leptospiral serovars not included in the vaccine preparation. The large number of pathogenic serovars (>230) imposes a major limitation on the production of a multiserovar component vaccine and the development of immunization protocols based on whole cell or membrane preparations. Protein antigens conserved among pathogenic serovars may contribute to overcome these limitations. The genome sequences of L. interrogans serovars Copenhageni and Lai have been published [7, 8], and comparative genome analysis between the two serovars was performed [9]. The main features found through genome analysis of the L. interrogans serovar Copenhageni that should contribute to the understanding of leptospiral physiology and pathogenesis are reviewed here. The results obtained through “genome data mining”, gene cloning, and protein expression are discussed.

1. Genome Features The L. interrogans serovar Copenhageni genome consists of two circular chromosomes with a total of 4,625,429 base pairs: chromosome I (CI) with 4,277,169 bp and chromosome II (CII) with 350,181 bp [7, 9]. The L. interrogans genome has one rrf gene, two rrl genes and two rrs genes coding for 5S, 23S, and 16S rRNA, respectively. As in other parasitic strains, L. interrogans serovar Copenhageni has only one rrf (5S) gene, which is located close to the origin replication region as described before for other strains of L. interrogans [7, 10]. Phylogenetic studies based on 16S rDNA sequences, using Leptonema as an outgroup, were performed, and the resultant analysis showed that Leptospira are split into two well-supported monophyletic groups, one of them formed by the pathogenic strains (e.g., L. interrogans) and the other formed by the non-pathogenic strains (e.g., L. biflexa). At the base of the clade of the pathogenic strains, L. inadai and L. fainei form a well-supported assemblage. Considering a constant divergence rate of 1 to 2% per 50 million years for the 16s rDNA [11], separation time between the two main assemblages (L. interrogans versus L. biflexa) was estimated to be approximately 590 to 295 million years ago [7].

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Based upon the localization of the gene metF, which encodes for the essential enzyme methylene tetrahydrofolate reductase [12], the small replicon of L. interrogans was previously suggested to be a second chromosome. A thorough genome sequence annotation revealed that genes encoding enzymes for metabolic pathways, such as glycolysis and the tricarboxylic acid cycle, as well as the enzymes for biosynthesis of amino acids and cofactors, are also spread between the two chromosomes. Sequence analysis of CII shows that an almost complete operon of genes coding for protoheme biosynthesis pathway is present (hemAIBCENYH). Although no homolog of the gene encoding for uroporphyrinogen III synthetase (hemD) was found, experimental evidence has shown that the hemC gene is able to cope with hemD activity [13]. L. interrogans therefore has the capability to synthesize protoheme de novo. In addition, thirteen genes clustered in CII coding for the cobalamin biosynthesis pathway were identified (cobC, cobD, cbiP, cobP, cobB, cobO, cobM, cobJ, cbiG, cobI, cobL, cobH, cobF) [7]. Orthologs of cobGKN genes, known to be involved in the cobalamin pathway [14], were not found. However, predicted coding sequences inside this operon in CII (LIC20133 and LIC20135) could perform these steps. Other genes present in the genome, such as LIC11145, LIC13354, LIC12391, and LIC10522, could also cope with these activities. The presence of cysG, in CI, may also be a cobalt-inserting enzyme in the B12 pathway [7, 15]. The remaining genes involved in this biosynthesis were found in CI (cysG/hemX/cobA, cobT/cobU, cobS). Indeed, comparable growth curves for L. interrogans serovar Lai, L. interrogans serovar Copenhageni, and L. biflexa serovar Patoc were obtained in either the presence or absence of vitamin B12 on the EMJH culture medium [7]. Thus L. interrogans, unlike the spirochetes Borrelia burgdoferi and Treponema pallidum, have the complete repertoire of genes for de novo synthesis of protoheme and cobalamin. The functional link between the two replicons strengthens the concept that the small replicon is indeed a second chromosome.

2. Comparative Genomics 2.1. L interrogans serovars Copenhageni and Lai Comparative genome analysis of L interrogans serovars Copenhageni and Lai showed that they are highly conserved and similar in size (~ 4.3 Mb and ~ 350 kb for CI and CII, respectively). The average nucleotide identity between the two serovars is 95%; the average nucleotide identity between pairs of predicted protein coding genes that are orthologs is 99%; and the numbers of ortholog pairs are 3079 and 261 for CI and CII, respectively [9]. Chromosomes CII in both serovars are collinear, but for the CI chromosome, a large inversion was detected. Flanking the inversion breakpoints, two identical copies of an IS element, in opposite orientation, were identified in serovar Lai. In serovar Copenhageni, 56 shotgun clones that span the inversion breakpoints and are anchored in non-repetitive portions of the sequence unequivocally confirmed the assembly. Taken together, the data suggested that the rearrangement took place in the Lai genome [9]. In addition, a 54 kb insertion was found in the Lai genome, accounting for the differences in genome organization. The Copenhageni and the Lai genomes contain 3728 and 4768 predicted open reading frames, respectively. The difference in the number of structural genes is mainly because the Brazilian Sequencing Group did not consider predicted coding sequences less than or equal to 150 bp in length that lacked significant homologs. The Copenhageni and the Lai genomes contain 64 and

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118 unique predicted coding genes, respectively. Eighty-one out of a hundred-eighteen structural genes unique to serovar Lai are located in the 54 kb insertion mentioned above [9]. 2.2. L. interrogans and Spirochetes A comparison between the L. interrogans genome and the genomes of the spirochetes B. burgdorferi and T. pallidum yielded the following results: 1167 (31%) of the genes in L. interrogans Copenhageni are found in B. burgdorferi and/or in T. pallidum; 666 (41%) of the genes in B. burgdorferi are found in the Copenhageni genome; 589 (57%) of the genes in T. pallidum are found in the L. interrogans genome. Three hundred sixty-two predicted genes were found to be shared among all three spirochaetes, of which 45 are hypothetical [7] (detailed list available at http://aeg.lbi.ic.unicamp.br/world/lic/). A thoroughly analysis of these common genes should contribute to the understanding of spirochete biology. 2.3. Energy Metabolism L. interrogans does not utilize glucose as a carbon and energy source under normal laboratory conditions [4], and it was thought that the glycolysis pathway was absent or incomplete. Surprisingly, the genome sequence revealed a complete glucose utilization route. Thus, the question arises as to why leptospires do not metabolize glucose as their energy source. In an attempt to answer this question, the genome was thoroughly searched, and only one glucose uptake system, a glucose/sodium symporter that is dependent on a sodium gradient across the bacterial membrane, was found within the leptospiral genome. In addition, no sugar ABC-transporter was identified, and an incomplete phosphoenolpyruvate-protein phosphotransferase system (PTS) was present with no B or C sugar permease components [9]. Taken together, these observations could explain the difficulties in utilization of glucose as a source of energy under certain growth conditions. Leptospira utilizes beta-oxidation of long-chain fatty acids as the major energy and carbon source [16], and as expected, a complete route was found. Glycerol metabolism genes are also present, including those encoding a glycerol-3-phosphate transporter, a glycerol uptake facilitator protein, glycerokinases, and a glycerol-3-phosphate dehydrogenase, which suggested that glycerol and fatty acids are obtained through phospholipid degradation. Two key enzymes of the pentose-phosphate pathway are missing, glucose-6-phosphate 1-dehydrogenase and 6-phosphogluconate dehydrogenase. L. interrogans probably obtains most of its reducing power for macromolecular biosynthesis via a membrane nicotinamide nucleotide transhydrogenase driven by a proton motive force [9]. L. interrogans generates ATP using an F0F1 type ATPase that is encoded by several genes organized in a single operon, atpBEFHAGDC. This operon has the same genetic organization as most of the eubacteria, in contrast to the ATP synthases found in B. burgdorferi and T. pallidum, which are of the V1V0-type [17, 18].

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3. Regulatory Systems and Signal Transduction A vast array of genes encoding for regulatory systems that enables Leptospira to respond to environmental signals was identified in the genome. There are 80 genes encoding components of the phosphorylation-mediated signal transduction pathway: 29 histidine kinases (HK), 30 response regulators (RR), and 18 hybrid kinase/regulators (HK/RR). Nineteen of the histidine kinases are located in the inner membrane, nine are cytoplasmic, and one is probably found in the periplasm, as predicted by the PSORT program [7, 19]. The response regulators are the cytoplasmic effectors of the message, which become functional after being phosphorylated by the cognate histidine kinase. The RRs may possess a second effector domain, which will perform its ultimate function, such as the DNA-binding helix-turn-helix domain (HTH). Other domains found in L. interrogans RRs are the GGDEF and EAL motifs, which correspond to putative diguanylate cyclase and phosphodiesterase domains, respectively [7].

4. Motility and Chemotaxis The motility and chemotaxis apparatus of L. interrogans is complex as its genome contains at least 79 putative motility-associated genes. All genes are well conserved among L. interrogans, T. pallidum, and B. burgdorferi, and 42 genes were found to be common to all three genera. However, the leptospiral genome contains multiple copies of a number of motility-associated genes, accounting in part for the higher number. In addition, the L. interrogans genome contains 11 putative genes encoding methylaccepting chemotaxis proteins (MCPs), which is roughly twice as many as in T. pallidum and B. burgdorferi. Forty-eight of the 79 motility-associated genes are located in 14 gene clusters varying in size from two to eight genes. Thus, as in T. pallidum and B. burgdorferi, the majority of the structural and functional motility genes are organized in operons. However, it seems that the operons suffered extensive rearrangements because they are usually smaller, corresponding to portions of the major Treponema and Borrelia operons [7, 9]. These differences might be associated with the high capacity of pathogenic leptospires to survive and adapt to a variety of environments and hosts.

5. Outer Membrane Proteins An important focus of current leptospiral research is the identification of outer membrane proteins (OMPs). Due to their location, leptospiral OMPs are likely to be relevant in host-pathogen interactions and consequently have the potential ability to stimulate heterologous immunity. The L. interrogans genome contains at least 263 predicted genes encoding for potentially surface-exposed integral membrane proteins, 250 of which were previously unknown. The leptospiral genome contains homologues of SecY and other secretory proteins involved in exporting proteins with signal peptides across the cytoplasmic membrane. Genes encoding for signal peptidases I (LIC11233, LIC10478), the lipoprotein biosynthesis pathway (LIC11063, LIC12389, and LIC12556, LIC13250), and the proteins involved in transport and incorporation of

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lipoproteins into the outer membrane (LIC12545, LIC10232, LIC12429, and LIC10233) were identified [7, 9]. One hundred eighty-four predicted coding sequences in the L. interrogans genome were found to have a lipoprotein signal peptidase cleavage site [9]. As shown by their lipobox motifs, all the predicted lipoprotein-coding sequences conform to the rule previously described [9, 20, 21]. Eighty-four outer membrane proteins with transmembrane domains were identified, including TolC orthologs, factor CzcC, TonB-dependent outer membrane proteins, and porin [9, 22]. Outer membrane proteins that are conserved among pathogenic serovars may serve as vaccine that avoids the limitations of currently available whole-cell preparations.

6. Searching for Immune Targets: “Immunomics” The advent of whole-genome sequencing has made an impressive impact on the microbial field landscape. The complete genomic sequence of Neisseria meningitidis serogroup B offered a new strategy for the identification of vaccine candidates [23]. This landmark approach, called reverse vaccinology, has been applied in the last few years, revolutionizing the vaccine research area [24, 25]. The design of vaccines is based on bioinformatic tools for the prediction of potential antigens in silico, hence narrowing down the universe to be tested. In addition, this approach has the advantage of revealing proteins independent of their abundance and without the need of growing the microorganism in vitro [26]. A first high-throughput screening aimed at identifying candidates for vaccine or diagnostic test evaluation was recently published [27]. These studies described 16 new leptospiral membrane-associated proteins selected from the genome of L. interrogans serovar Copenhageni [7, 9]. The rationale for the choice of the predicted coding sequences, described by Gamberini and colleagues [27], is that surface-associated molecules are potential targets for inducing immune responses. The PSORT program (http://psort.nibb.ac.jp/) [19] was used to predict the localization of the coded proteins within the bacterium. Public and custom sequence-specific search algorithms were used for identification of sequence motifs including lipoprotein cleavages sites, transmembrane domains and signal peptides (http://www.cbs.dtu.dk/services/TMHMM) (http://www.cbs.dtu.dk/services/SignalP) [28, 29]. Putative proteins, homologous to surface proteins previously characterized as virulence factors in other organisms, were searched for by blast analysis (http://www.ncbi.nlm.nih.gov/BLAST/) [30]. The in silico approach resulted in a large number of genes covering ~20 % of the total number of predicted proteins in the genome. From these sequences, the selection was focused mainly on hypothetical, unknown proteins, having either signal peptide sequences or lipobox motifs [7, 9]. Genes encoding proteins with known cytoplasmic functions were excluded. Of the 206 selected coding sequences, more than 97% were amplified. The correct sequences were confirmed by DNA sequencing and 175 genes (84%) were effectively cloned into pENTR. The DNA inserts were transferred by recombination from pENTR to pDEST17 expression vector. This E. coli vector expresses the recombinant proteins with six histidine residues at the N-terminus, which allows a rapid purification of the protein by metal chelation chromatography. By using this approach, the authors have successfully expressed and purified 150 recombinant proteins. Purified proteins were screened for reactivity by immunoblotting with serum from patients diagnosed with leptospirosis. The recombinant lipoprotein LipL32 was employed as a positive control

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since it was shown to be immunogenic and highly conserved among Leptospira pathogenic serovars [31, 32]. Antibodies present in serum from convalescent leptospirosis patients recognized 16 proteins of 100 tested. These comprise: four probable new lipoproteins, six leptospiral conserved hypothetical proteins, four conserved hypothetical proteins, one hypothetical protein, and one peptidoglycanassociated membrane protein [27].

Whole genomic sequence

Computer prediction of surface-exposed proteins (2000ORFs)

In silico selected protein candidates (206 ORFs)

Cloning and express recombinant proteins: 175 ORFs cloned; 150 recombinant proteins expressed

Immunoblotting: 16 proteins

Figure 1. Strategy used for identification of vaccine candidates from the whole genome sequence.

Protein expression in the most prevalent pathogenic serovars of L. interrogans is an important requirement for leptospiral vaccine candidates. The conservation of the selected proteins was evaluated against protein extracts from several L. interrogans serovars: Canicola, Icterohaemorrhagiae, Copenhageni, Bratislava, Hardjo, Autumnalis, Pomona, Pyrogenes, Grippotyphosa, and the nonpathogenic strain L. biflexa serovar Patoc. Four of 10 proteins tested proved to be highly conserved among the pathogenic leptospires. Most interesting, none of these four proteins were present in the nonpathogenic L. biflexa strain, suggesting that they may be relevant for pathogenesis [27]. The protective immune activity of these proteins is currently under investigation in our

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laboratories. Figure 1 summarizes the experimental approach used in the work performed by Gamberini and colleagues [27].

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[24] Wizemann TM, Heinrichs JH, Adamou JE, et al. (2001) Use of a whole genome approach to identify vaccine molecules affording protection against Streptococcus pneumoniae infection. Infect. Immun. 69:1593–8. [25] Rappuoli, R. (2001) Reverse vaccinology, a genome-based approach to vaccine development. Vaccine 19: 2688–2691. [26] Adu-Bobie, J., Capecchi, B., Serruto, D., Rappuoli, R., and Pizza, M. (2003) Two years into reverse vaccinology. Vaccine 21: 605-610. [27] Gamberini M, Gomez RM, Atzingen MV, Martins EA, Vasconcellos SA, Romero EC, Leite LC, Ho PL, Nascimento AL. (2005) Whole-genome analysis of Leptospira interrogans to identify potential vaccine candidates against leptospirosis. FEMS Microbiol Lett. 244:305-13. [28] Krogh A., Larsson B., von Heijne G.and Sonnhammer EL.(2001) Predicting transmembrane protein topology with a Hidden Markov Model: Application to complete genomes. J. Mol. Biol. 305: 567-580. [29] Bendtsen JD., Nielsen H., von Heijne G. and Brunak S. (2004) Improved Prediction of Signal Peptides: SignalP 3.0. J. Mol. Biol. 340: 783–795. [30] Altschul FF., Madden TL., Schäffer AA., Zhang J., Zhang Z., Miller W. and Lipman DJ. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 25 (17): 3389–3402. [31] Dey, S., Mohan, C.M., Kumar, T.M., Ramadass, P., Nainar, A.M., Nachimuthu, K. (2004) Recombinant LipL32 antigen-based single serum dilution ELISA for detection of canine leptospirosis. Vet. Microbiol. 103: 99-106. [32] Haake, D.A., Suchard, M.A., Kelley, M.M., Dundoo, M., Alt, D.P., Zuerner, R.L. (2004) Molecular evolution and mosaicism of leptospiral outer membrane proteins involves horizontal DNA transfer. J. Bacteriol. 186: 2818-28.

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Genotypic Variation and Borrelia burgdorferi Pathogenesis Ira SCHWARTZ a,b,1 , Guiqing WANG a, Radha IYER a, Caroline OJAIMI a, Darya TEREKHOVA a, Sabina SANDIGURSKY a, Gary P. WORMSER b and Dionysios LIVERIS a a Department of Microbiology & Immunology, New York Medical College, Valhalla, NY 10595, USA b Division of Infectious Diseases, Department of Medicine, New York Medical College, Valhalla, NY 10595, USA Abstract. Lyme disease, the most commonly reported arthropod-borne disease in the United States, is caused by infection with the spirochete, Borrelia burgdorferi. The acute stage of the infection has a varied presentation, ranging from mild, localized disease (characterized by a skin lesion) to highly symptomatic, disseminated disease. We hypothesize that this is due, in part, to B. burgdorferi genotypic variation. By combining PCR amplification of the 16S-23S rDNA spacer with restriction fragment length polymorphism (RFLP) analysis, several genotypes were identified among B. burgdorferi clinical isolates obtained from either skin or blood of early Lyme disease patients. Hematogenous dissemination in humans is associated with a distinct genotype and disease severity and spirochete burden was also associated with this same genotype in a murine model of Lyme disease. A genomic approach was undertaken to elucidate the differences in genome content and/or gene expression that may result in disease variability. Comparative transcriptional profiling of two clinical isolates with distinct genotypes (invasive and attenuated) was performed using whole genome arrays. A total of 78 ORFs had significantly different expression levels in the two isolates. Nearly 25% of the differentially expressed genes are predicted to be localized on the cell surface, implying that these two isolates have considerably different cell surface properties. Comparative genome hybridization demonstrated that genotypic variation largely results from differences in plasmid content and/or sequence and revealed several plasmid-encoded candidate genes that are uniquely absent in attenuated strains. A number of genes identified in these investigations are currently under further study by genetic analysis to substantiate a possible role in virulence. Keywords. Borrelia burgdorferi, pathogenesis, comparative genomics

1

Lyme

disease,

genotypic

variation,

Corresponding Author: Department of Microbiology & Immunology, New York Medical College, Valhalla, NY 10595 USA; E-mail: [email protected], Phone: 914-594-4658.

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Introduction Lyme borreliosis is a global arthropod-borne zoonosis, occurring throughout the Northern hemisphere [1, 2]. It is caused by infection with spirochete Borrelia burgdorferi sensu lato [2, 3]. This group of organisms is comprised of at least 12 genomic species [3í5]. Three of these are responsible for the majority of human disease worldwide; B. burgdorferi sensu stricto in the US and Europe and Borrelia garinii and Borrelia afzelii in Eurasia [3, 6, 7]. Globally, it has been noted that symptoms of Lyme borreliosis vary with geographic location; neurologic and chronic skin manifestations predominating in Europe and arthritic symptoms being more common in the US [1, 2]. It has been proposed that these differences may result from differences in pathogenesis between the infecting spirochetal species [3, 8, 9]. Lyme disease is the most frequently reported arthropod-borne disease in the US and all Lyme disease is caused by B. burgdorferi sensu stricto (referred to simply as B. burgdorferi throughout this chapter) [3, 10]. A characteristic skin rash, erythema migrans (EM) occurs in >75% of patients [1]. Furthermore, 70-80% of patients with EM are symptomatic at presentation (i.e., report at least one symptom such as fever, headache, neck or joint pain) and over 50% can be shown to have disseminated infection based on having a positive blood culture for B. burgdorferi or the presence of multiple EM skin lesions [11]. Thus, early Lyme disease in the US has a varied presentation, ranging from no symptoms to disseminated infection with multiple symptoms. Variation in symptom profile or in spirochete dissemination could be related to differences in the host and/or in the infecting strain of B. burgdorferi. One hypothesis is that the distinctiveness in disease manifestations may be the result of genotypic variation of the infecting strain of B. burgdorferi. A corollary of this hypothesis is that not all B. burgdorferi genotypes circulating in nature are capable of causing human disease.

1. Molecular Typing of B. burgdorferi Many methods have been employed for the molecular typing of B. burgdorferi. These include serotyping, multi-locus enzyme electrophoresis, ribotyping, pulsed-field gel electrophoresis (PFGE), randomly amplified polymorphic DNA and sequencing of various target genes (reviewed in [3]). We combined PCR amplification of the proximal portion of the 16S-23S rDNA spacer with restriction digestion of the

Table 1. Genotype distribution of B. burgdorferi isolates cultivated from skin or blood of early Lyme disease patients.1 RST1

RST2

RST3

Total

Skin

81 (28%)

129 (44%)

83 (28%)

293

Blood

53 (42%)

57 (45%)

17 (13%)

127

1

Cultivated from patients presenting with EM at the Lyme Disease Diagnostic Center, New York Medical College, 1991-2002.

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amplified product to produce a rapid assay that could be employed with infected wildlife, tick and human specimens and did not require prior cultivation of B. burgdorferi or genomic DNA isolation [12, 13]. Initially, PCR-RFLP typing of 217 B. burgdorferi isolates originally cultivated for skin biopsies or blood of early Lyme disease patients showed three major RFLP types which are designated RST1, RST2and RST3 [14]. Currently, 420 B. burgdorferi isolates cultivated from early Lyme disease patients with EM between 1991-2002 have been typed by this method and the results are presented in Table 1. The distribution of genotypes was significantly different in skin and blood (P<.001). RST1 isolates were over-represented among blood isolates, whereas RST3 strains were under-represented.

2. Hematogenous Dissemination is Associated with Genotype Is infection due to a specific B. burgdorferi genotype correlated with signs or symptoms in early Lyme disease? To investigate this further, 104 adult patients with EM and a positive skin biopsy culture were studied [15]. A highly significant association between infecting RST genotype and spirochetemia (as determined by positive blood culture) was found (P <.001). Patients were 13.8-fold more likely to be spirochetemic if the B. burgdorferi isolated from their EM lesions was of the RST1 genotype than the RST3 genotype (Table 2). Furthermore, the same association existed for the presence of multiple EM lesions (P=.045), a manifestation believed to arise by hematogenous dissemination of B. burgdorferi. Patients with RST1 infection in skin were 4.6-fold more likely to present with multiple EM than were patients with an RST3 skin infection (Table 2). No other significant associations between RST genotype and clinical signs or symptoms were found except for a history of fever or chills (P=.033). The results are consistent with a specific genotype of B. burgdorferi influencing Lyme disease pathogenesis and, in particular suggested an association between RST1 skin infection and hematogenous dissemination.

Table 2. Comparison of blood culture yield and presence of multiple EM lesions with genotype of the cultured skin isolate.1 RFLP type (%) Type 1

Type 2

Type 3

(n=28)

(n = 44)

(n = 32)

P2

Blood culture positive

43

20

3

.001

Multiple EM lesions

29

23

6

.045

Blood culture positive or multiple EM lesions

57

34

9

.001

1 2

Adapted from Wormser et al. [15]. Likelihood ratio test..

Liveris et al. measured the B. burgdorferi burden in EM lesions by quantitative PCR analysis of 2 mm skin biopsy specimens [16]. The mean number of spirochetes varied from 10 to 11,000 per 2 mm biopsy. The number of B. burgdorferi organisms in EM lesions showed a significant association with B. burgdorferi genotype (P=.008).

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The average number of spirochetes in EM lesions of patients infected with RST1 strains was nearly three times higher that that in patients infected with either RST2 or RST3 strains [16]. This study suggested that the association of hematogenous dissemination with RST1 infection in skin may be caused by higher spirochete numbers in EM lesions of RST1-infected patients. To elucidate further the association between B. burgdorferi genotypic variation and disease pathogenesis, kinetics of spirochete dissemination and severity of resultant disease were evaluated in a murine model [17, 18]. C3H/HeJ mice were infected by needle inoculation with 10,000 organisms, blood and tissue specimens were collected at various times post-inoculation, specimens were cultured and spirochete loads were determined by quantitative PCR, and joint and cardiac histopathology were evaluated. The results were striking. B. burgdorferi were cultivated more frequently from specimens obtained from RST1-infected mice than those infected with RST3 isolates. This was true for all tissue types tested (Table 3). These studies revealed that there were two distinct groups within the RST3 genotype, those that yielded positive specimen cultures (RST3B) and those that did not (RST3A). It should be noted that the distinction between RST3A and RST3B is a functional one, i.e. it is based solely on whether an isolate disseminates in mice. This cannot be determined a priori by molecular typing, but rather only by mouse infection studies. RST3A isolates yielded no positive cultures from any animals, produced no ankle joint edema and resulted in virtually no evidence of disease in cardiac or joint tissue as measured by histopathology [18]. Significantly greater numbers of spirochetes were detected in blood, skin, heart and joint of RST1- and RST3B-infected mice than in those infected with RST3A isolates (Figure 1). Similarly, the mean number of spirochetes in skin, joint and heart were significantly greater in RST1-infected animals than in those infected with RST3B isolates. In total, 6 RST1 and 12 RST3 isolates have been tested for infectivity in C3H/HeJ mice. To date, 4 isolates have not yielded any evidence of dissemination and are designated as RST3A. By contrast, spirochetes were recovered from all animals infected by any RST1 isolate. RST3B infection resulted in intermediate rates of infection. These results strongly suggest that certain genotypes of B. burgdorferi are more invasive and pathogenic than others and are in agreement with the association of specific B. burgdorferi genotypes with hematogenous dissemination in humans [15].

Table 3. Culture of B. burgdorferi from tissues of C3H/HeJ mice inoculated with RST1, RST3A and RST3B isolates. No. positive/No. tested (%) Genotype (No. of isolates)

Blood1

Ear2

Heart3

Bladder3

Total

RST1 (6)

28/35 (80)

39/41 (95.1)

31/34 (91.2)

31/35 (88.6)

129/145 (89)4

RST3A (4)

0/19 (0)

0/27 (0)

0/24 (0)

0/24 (0)

0/94 (0)

RST3B (8)

5/15 (33.3)

33/40 (82.5)

30/40 (75)

25/39 (64.1)

93/134 (69.4)

1

Day 7. 2 Day 14. 3 Day 21. 4 P<.0001 for comparison to RST3B, chi-square.

I. Schwartz et al. / Genotypic Variation and Borrelia burgdorferi Pathogenesis

B

A

No. of spirochetes (log) per 200 ng of mouse DNA

3

No. of spirochetes (log) per ml plasma

6 5

**

4

*

3

**

2

*

1

2 1

0

0 RST1

RST3A

RST1

RST3B

RST3A

RST3B

Skin

Day 7 plasma

C

D

No. of spiorchetes (log) per 200 ng of mouse DNA

4

3

No. of spirochetes (log) per 200 ng of mouse DNA

4

3

**

**

2

**

1

**

**

2

**

1

0

Heart

ST 3A

ST 3B R

ST 1 R

ST 3B

ST 3A

R

R

RST3B

R

RST3A

Ear biopsy

ST 1

0

RST1

R

128

Joint

Figure 1. Spirochete burdens in tissues of mice infected with B. burgdorferi RST1 or RST3 isolates. Number of spirochetes in tissue were determined by quantitative PCR as described [18]. * P<.05 and ** P<.001 , vs. mice infected with RST3A isolates (Student’s t test). Adapted from Wang et al. [18], with permission.

3. B. burgdorferi Genomics and Pathogenesis What factors underlie the observed differences in pathogenicity between B. burgdorferi genotypes? To address this question more extensive genetic and genomic studies have been undertaken. A phylogram based on the 16S-23S rDNA spacer sequences of representative B. burgdorferi clinical isolates clearly differentiates RST1 and RST2 isolates as deeply branching clades; sequences for RST3 isolates are more heterogeneous (Figure 2). Furthermore, PFGE and plasmid content analysis of selected isolates revealed a significant association between RST type, PFGE pattern and plasmid content (P < 0.001) [19, 20]. These findings imply a clonal origin for these RST genotypes and are in agreement with earlier studies demonstrating little or no genetic exchange between chromosomal genes and little evidence for plasmid transfer between isolates [21, 22].

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129

B348

N40

B418

RST3 B356

B331

94

B500

297

99

84

B379

RST2

100 BL224

83 B31

100 B515

68

1 nt

RST1 B479

BL206

Figure 2. Phylogenetic tree based on 16S-23S rDNA spacer sequence of selected B. burgdorferi isolates. Tree was constructed by maximum parsimony in PAUP version 4.0b10. Statistical stability of each node was determined by bootstrap analysis with 1,000 replicates.

It should be noted that RST typing is based on a non-coding spacer region within the rRNA gene cluster of B. burgdorferi. It is unlikely that this region itself encodes any factors that have a direct role in dissemination or pathogenesis. It is clear, however, that this region is in linkage disequilibrium with genes that do encode functions involved in virulence. This is supported by the limited roles that recombination and plasmid exchange play in shaping B. burgdorferi genomic structure and the clonal nature of B. burgdorferi [22]. 3.1. B. burgdorferi Dissemination and OspC A number of studies have suggested that differential pathogenicity of B. burgdorferi isolates is determined by outer surface protein C (OspC) [23, 24]. Virtually all invasive isolates studied by Seinost, et al. were associated with only 4 of 21 major OspC groups [23]. OspC is required for mammalian infection by B. burgdorferi either because it facilitates migration of spirochetes from the tick midgut to the salivary glands or because it mediates transmission from tick to mammal [25, 26]. ospC expression is absent or minimal when spirochetes are in the tick midgut, but is rapidly upregulated during the bloodmeal [27, 28]. These findings have focused attention on OspC as a primary determinant of B. burgdorferi pathogenicity.

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In order to explore the relationship between RST types and OspC, OspC types for over 200 clinical isolates for which RST data were available were determined by either direct sequencing of the gene or reverse line blotting. The data presented in table 4 reveal a number of interesting observations. (1) There is a remarkable correlation between RST and OspC; each RST is associated with 2 or more unique OspC groups with no overlap between groups. (2) All RST1 isolates belong to either OspC group A or B. These latter groups were previously characterized as invasive clones [23]. This supports the earlier conclusions that RST1 isolates are highly invasive based on correlative patient data and mouse studies [15, 17, 18]. (3) Earlier studies suggested that only 4 OspC groups (A, B, I and K) could cause disseminated disease [23]. The data in table 4 indicate that at least 5 additional groups (C, F, H, M and N) are capable of dissemination in humans. Cultivation of OspC group C and N isolates from the blood of Lyme disease patients in Maryland has recently been reported by Earnhart et al. [29]. (4) All OspC types associated with RST1 and RST2 strains are capable of disseminating from skin to blood. In contrast, only 3 of 9 OspC types associated with RST3 isolates were isolated from blood.

Table 4. Correlation of RST and ospC types in B. burgdorferi clinical isolates. RST1 ospC type

RST2

Skin

Blood

A

26

14

B

17

5

Skin

RST3 Blood

C

Skin

Blood

2

2

D

4

E

14

F

5

2

G H

11 9

2

I

17

J

2

K

43

19

15

3

M N

6

O

1

U

10

7

1

At first glance, these findings would appear to strengthen the notion that OspC is the major determinant of B. burgdorferi invasiveness. However, analysis of four RST3A isolates (i.e., RST isolates that do not disseminate in mice) suggests that this is not the case. The isolates each fall into a distinct OspC group (E, G, I and M); three of these groups (E, I and M) also contain isolates that have been shown to either

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disseminate in mice or humans. Thus, the presence of a specific ospC is not the sole determinant of B. burgdorferi dissemination. Given the rarity of recombination in B. burgdorferi (see above) and the linkage disequilibrium between ospC and the rDNA spacer, it is possible that ospC is simply a linked marker for another locus (or loci) responsible for dissemination. Alternatively, expression of ospC, or its regulation, in RST3A strains may be reduced or impaired. 3.2. Comparative Transcriptional Profiling of B. burgdorferi We are currently taking a global genomic approach in an attempt to identify genes that may be responsible for the observed differences in pathogenicity. The gene expression profiles of a virulent RST1 strain (BL206) and an attenuated RST3A strain (B356) grown at 34˚C were determined using B. burgdorferi genome arrays and compared [30]. A total of 78 ORFs had significantly different expression levels in the two isolates; 35 ORFs were expressed at significantly higher levels in B356 and 43 ORFs had significantly higher expression in BL206. Twenty-one (27%) of the differentially expressed ORFs were chromosomally encoded, with the remainder located on plasmids. Forty-five of 78 these genes (58%) have no database match and are of unknown function. Interestingly, nearly 25% of the differentially expressed genes are predicted to have a cell surface localization. This implies that the two strains may have considerably different cell surface properties. Strain B356 does not disseminate in C3H/HeJ mice, whereas BL206 infection resulted in disseminated infection of all tested tissues with a high pathogen load [18]. In the case of B356, spirochetes could not be recovered from the inoculation site 24 hours after inoculation. Similar results were obtained with immunodeficient SCID mice [17, 18], suggesting that isolate B356 is rapidly cleared by components of the innate immune system. Given that pathogenassociated molecular patterns play a central role in the innate immune response to infection, the potentially distinctive nature of the BL206 and B356 cell surfaces may contribute to differential recognition of these isolates by the innate immune system, thereby yielding differences in spirochete dissemination. These studies demonstrate the utility of comparative transcriptional profiling for identifying expression differences among isolates with varying virulence potential. The 78 ORFs identified provide a reasonable starting point for investigation of factors involved in hematogenous dissemination of B. burgdorferi. 3.3. Comparative Genomics of B. burgdorferi Comparative genome hybridization may also reveal genomic differences between B. burgdorferi strains and provide clues to potential virulence genes. The genome content of 16 B. burgdorferi isolates was compared to that of the type strain B31MI by microarray analysis (Figure 3). RST1 isolates were the most homogeneous and showed the lowest variation relative to B31MI (this is not surprising since B31MI has an RST1 genotype). The RST2 and RST3 isolates are substantially more heterogeneous at the whole genome level. These findings are consistent with previous analyses based on comparison of either 16S-23S rDNA spacer or ospC sequence [18, 31]. This strain-tostrain variation is associated largely with differences in plasmid gene sequence or content. The B. burgdorferi chromosome, circular plasmids (cp26 and cp32) and linear plasmid lp54 are highly conserved. Ninety-three percent of chromosomal ORFs are

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nearly identical and where divergence does occur it is the result of single nucleotide variation; these SNPs, in general, result in synonymous or conservative codon changes. Similar levels of conservation were observed for cp26 and lp54 (with the notable exception of ospC [BBB19] and BBA70). By contrast, genetic content of the remaining linear plasmids is highly divergent. In this case, lack of hybridization to the array often resulted from complete absence of the ORF; a number of such deletions were confirmed by southern blotting of total genomic DNA separated by PFGE. Furthermore, these deletions were often clustered. There is a strong correlation between complete genome hybridization profiles and other typing methods, which also correlate to differences in pathogenicity. Comparison of genomic differences between the least and the most invasive isolates predict potential virulence-encoding elements. Further studies of these regions should facilitate identification of factors responsible, in part, for the pathogenicity B. burgdorferi. RST2 and RST3

RST1

chromosome

plasmid

Figure 3. Comparative genome hybridization of 17 B. burgdorferi isolates. Each column represents a different isolate and each row represents a different ORF. Black (or green) represents conserved ORFs and red represents divergent ORFs with respect to the B31MI reference sequence.

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4. Summary and Conclusions The studies described here demonstrate that there is substantial genotypic variation among B. burgdorferi clinical isolates. The risk of hematogenous dissemination in humans is associated with B. burgdorferi genotype and disease severity and spirochete burden is associated with this same genotype in a murine model of Lyme disease. Comparative transcriptional profiling identified several candidate genes that have altered expression in virulent and attenuated strains. Comparative genome hybridization demonstrated that genotypic variation is largely the result of differences in plasmid content and/or sequence. Furthermore, comparative genome hybridization reveals several plasmid-encoded candidate genes that are uniquely absent in attenuated strains. Several of the candidate genes identified by genomic analysis are currently under study for fulfillment of “molecular Koch’s postulates” as virulence determinants.

Acknowledgements This work was supported by grants AR41511 and AI45801 from the National Institutes of Health and the G. Harold and Leila Y. Mathers Charitable Foundation.

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Multilocus Sequence Analysis (MLSA) as an Alternative to Whole DNA/DNA Hybridization (WDDH) in Borrelia burgdorferi sensu lato Taxonomy Guy BARANTON 1 and Danièle POSTIC Laboratoire des Spirochètes, Institut Pasteur, Paris France Abstract. Thirty years after it was introduced, whole DNA/DNA hybridization (WDDH) is still considered as the gold standard for bacterial taxonomy. However, WDDH has serious limitations, as it requires large volumes of cultures, is a highly fastidious and time consuming procedure, lacks of reproducibility, and requires pairwise comparisons between unknown genome and genomes from previously described species. These limits have hampered progress in Borrelia taxonomy. With the development of sequencing, it is inviting to propose MLSA as an alternative to WDDH. We chose fragments from seven loci: rrs (16S rRNA), flaB, groEL, hbb, recA, ospA, and the internal transcribed spacer rrf-rrl (ITS). We sequenced these loci for 2–4 representative Borrelia burgdorferi sensu lato strains from each known species present on both the European and North American continents. Phylogenetic trees were built from sequences of each locus, confirming that lateral transfer was a rare event, and from the concatenated sequence of the seven loci from each strain. Genetic distances were calculated from concatenated sequences and compared to those obtained by WDDH (both published and unpublished). The two methods were strictly correlated. A precise cutoff of MLSA-based genetic distances (0.021) allowed us to delineate new species more safely and accurately than the usual 70% DNA relatedness cutoff. We have confirmed in this way a recently described sixth species in Europe, B. spielmanii, whose reservoirs are dormice. Three new nonpathogenic Borrelia species are delineated in California, genomospecies 1, 2, and 3. Finally, the phylogenetic tree obtained from the concatenated sequences from 42 isolates (from both Europe and North America) shows 12 clusters, each corresponding to a species. These clusters form two main groups. One group comprises only European species, whereas the other group comprises B. burgdorferi sensu stricto isolates from Europe and the USA, including both Borrelia spp. “border line” to B. burgdorferi sensu stricto. This clustering confirms our previous hypothesis of the American origin of B. burgdorferi sensu stricto and closely related isolates that have been imported recently into Europe from the USA. Keywords. Multilocus sequence analysis, whole DNA/DNA hybridization, taxonomy, Borrelia burgdorferi sensu lato

1 Corresponding author. Mailing address: Guy Baranton. Laboratoire des Spirochètes, Institut Pasteur, 28 rue du docteur Roux, 75724 Paris Cedex 15, France. Tel: 33 (1) 45 68 83 37. Fax: 33 (1) 40 61 30 01. E-mail: [email protected].

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Introduction The taxonomy of Borrelia burgdorferi sensu lato based upon whole DNA/DNA hybridization (WDDH) has been established by studies beginning in 1984 up to the present time. Twelve species have been identified this way: B. burgdorferi sensu stricto (s.s.) [1], B. garinii [2], B. afzelii [3], B. japonica [4], B. andersonii [5], B. turdi and B. tanukii [6], B. valaisiana [7], B. lusitaniae [8], B. bissettii [9], and B. sinica [10]. Recently, a potential new species, B. spielmanii, has been described but not validated due to the lack of WDDH [11]. The cumbersomeness of WDDH, considered the gold-standard method in bacterial taxonomy [12], has hampered taxonomic studies of the genus Borrelia. Indeed, the need for significant volumes of these fastidiously growing bacteria as well as the requirement to compare any new potential taxon to every species already identified on a given continent, have discouraged research in this field. In addition, experimental variations are common when WDDH is repeated and may lead to inconsistencies or discrepancies when applied to borderline isolates. Multilocus sequence typing (MLST) [13] was initially proposed to replace multilocus enzyme electrophoresis (MLEE) [14] in population genetic studies. MLST is used in a cluster analysis procedure. These conditions have led us to adopt some rules. These are to use housekeeping genes whose evolution is mainly chronometric and will lead to data very similar to those of MLEE, and to use fragments of about 400 bp that warrant an optimal number of alleles. Conversely, the use of multilocus sequence analysis (MLSA) [15] in taxonomy deals with whole genome evolution related to its suitability within a framework of an ecological niche. Therefore, the loci to be used should reflect the whole genome and not be restricted to a single category. To use only chronometric genes, for instance, could lead to ignoring recently emerged species or conversely, to overestimating the divergence within an ancient species. The size of the loci fragments involved in MLSA is not critical. What is important is to develop a significant “abstract” of the whole genome which constitutes about 10–3 of the actual genome size. The diversity of the loci under consideration is the most important point: conserved genes, variable genes, housekeeping genes, informational genes, adaptative genes, and even non-coding loci should be included. Another important point is to exclude as much as possible loci often subjected to lateral transfer. Finally, the concatenated sequences are analyzed by distance matrix methods. We propose to use MLSA to circumvent the difficulties entailed in using WDDH in Borrelia taxonomy studies. Our objectives will be pursued sequentially in three successive steps: 1.

2.

3.

Compare MLSA data obtained from seven distinct loci of isolates belonging to previously known species for which DNA relatedness values are available. A deduced MLSA specific cutoff has thus been applied to B. spielmanii to assess the potential alternative use of MLSA to delineate species. Use MLSA on a set of unclassified isolates (Borrelia spp.) from North America which in the past have led to borderline DNA relatedness values [9] as a step towards applying MLSA to actualize the taxonomy of B. burgdorferi sensu lato. Use MLSA to study a similar set of previously studied Borrelia spp. from Europe [11]. Since MLSA for taxonomic purposes is used in a phylogenetic way on concatenated sequences, it allows us to analyse the evolutionary

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relationships between different species on whichever the continent where they have been isolated, unlike WDDH, which cannot be used in this manner.

1. Materials and Methods 1.1. B. burgdorferi sensu lato Strains Forty-two strains used in this study (Table 1) belong to seven species endemic to Europe and North America: B. andersonii, B. bissettii, B. burgdorferi s.s., B. garinii, B. afzelii, B. valaisiana and B. lusitaniae. Information regarding DNA–DNA reassociation was available for each of these strains [2, 3, 8, 7, 17, 30], and sequences at several of the selected loci were accessible from literature or databases [7, 8, 24, 31-33]. For the delineation of B. spielmanii by MLSA, we used the first described isolate of this group, the strain A14S, isolated from a patient’s erythema migrans [28], and three strains, including the type strain PC-Eq17, derived from Ixodes ricinus ticks fed on garden dormice, Eliomys quercinus, live-trapped in Alsace, France [11], as well as two isolates obtained from acute skin lesions of a Danish and a Hungarian patient.

Table 1. Characteristics of the isolates used in this study.1 Isolate

Species

Geographic origin

Biologic origin

Reference

B31T

B. burgdorferi ss

USA

I. scapularis

[1]

IP1

B. burgdorferi ss

France

Human CSF

[16]

212

B. burgdorferi ss

France

I. ricinus

[17]

297

B. burgdorferi ss

USA

Human CSF

[18]

Sh2-82

B. burgdorferi ss

USA

I. scapularis

[19]

21123

B. andersonii

New York

I. dentatus

[17]

21133

B. andersonii

New York

I. dentatus

[17]

19952

B. andersonii

New York

I. dentatus

[17]

DN127

B. bissettii

California

I. pacificus

[20]

CA55

B. bissettii

California

I. neotomae

[17]

B. bissettii

California

I. neotomae

[21]

VS461

B. afzelii

Switzerland

I. ricinus

[16]

Pgau

B. afzelii

Germany

Human skin

[22]

B. afzelii

Sweden

Human skin

[17]

20047

B. garinii

France

I. ricinus

[23]

PBi

B. garinii

Germany

Human CSF

[24]

CA128 T

BO23 T

NT29

B. garinii

Japan

I. persulcatus

[17]

VS116T

B. valaisiana

Switzerland

I. ricinus

[7]

UK

B. valaisiana

United Kingdom

I. ricinus

[7]

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138

Table 1. Characteristics of the isolates used in this study.1 Isolate M7

Species B. valaisiana

Geographic origin Netherlands

Biologic origin

Reference

I. ricinus

[25]

NE231

B. valaisiana

Switzerland

I. ricinus

[26]

PotiB1

B. lusitaniae

Portugal

I. ricinus

[8]

T

PotiB2

B. lusitaniae

Portugal

I. ricinus

[8]

T

B. spielmanii

France

I. ricinus

[27]

PC-Eq2/1

B. spielmanii

France

I. ricinus

[27]

PC-Eq2r

B. spielmanii

France

I. ricinus

[27]

A14S

B. spielmanii

Netherlands

Human skin

[28]

DK35

B. spielmanii

Denmark

Human skin

[29]

PC-Eq17

PZ30802

B. spielmanii

Hungary

Human skin

Richter et al., in press

CA2

Borrelia spp.

California

I. neotomae

[21]

CA8

Borrelia spp.

California

I. pacificus

[21]

CA13

Borrelia spp.

California

I. neotomae

[19]

CA28

Borrelia spp.

California

I. pacificus

[21]

CA29

Borrelia spp.

California

I. pacificus

[21]

CA404

Borrelia spp.

California

D. californicus

[21]

CA443

Borrelia spp.

California

D. californicus

[21]

CA446

Borrelia spp.

California

D. californicus

[21]

NE49

Borrelia spp.

Switzerland

I. ricinus

[17]

5LM218

Borrelia spp.

France

I. ricinus

Unpublished

Z41493

Borrelia spp.

Germany

I. ricinus

[17]

Z41293

Borrelia spp.

Germany

I. ricinus

[17]

Z51094

Borrelia spp.

Germany

I. ricinus

[17]

1

Type strains indicated by T.

DNA of each of these strains was extracted from the centrifugation pellet of cultivated isolates, either by boiling at 100°C for 10 min. or with the QIAamp DNA Mini Kit. Most of the Borrelia spp. isolates have already been studied by molecular typing methods [9, 33] and on some occasions by WDDH, leading to borderline results [personal, unpublished]. 1.2. MLST Seven loci, rrs, hbb, groEL, recA, fla, ospA, and the rrf-rrl intergenic spacer, were selected for analysis. Characteristics of the amplified fragments and corresponding primer sequences are shown in Table 2. The numbering derives from B. burgdorferi s.s. strain B31T. On some occasions (CA13 for recA and CA2 for hbb), a second set of

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139

primers had to be used. All loci were amplified by a single PCR. Amplification reactions were performed in a final volume of 50 μl, comprising 0.2 PM of each primer of a primer pair, 200 PM of each deoxynucleoside triphosphate, 1.25 U or 1 U of Taq polymerase (Q. Bio gene or Qiagen, respectively), and 1x Taq buffer (1.5 mM MgCl2). The mixture was placed in a thermocycler, heated to 93°C for 1 min and subjected to 35 cycles of denaturation for 1 min at 93°C, annealing for 1 min at either 51°C for rrs and groEL loci, or at 59°C for the remaining loci, and extension for 1 min at 72°C, followed by a final extension step at 72°C for 5 min. Amplification products were sequenced either by Genome-Express (Meylan, France) or by the dideoxynucleotide chain termination method on a Licor DNA4200 sequencer using the same primers as for PCR.

Table 2. Characteristics of the amplified fragments and corresponding primer sequences. Locus

Fragment length

rrs

522

Primers F: AGAGTTTGATCCTGGCTTAG (10-29) R: CTTTACGCCCAATAATCCCGA (572-552)

fla

457

F: AACACACCAGCATCACTTTCAGG (475-497)

groEL

268

F: TACGATTTCTTATGTTGAGGG (552-572)

R: GATTWGCRTGCGCAATCATTGCC (976-954)

R: CATTGCTTTTCGTCTATCACC (861-841) hbb

327

F: GCGAAGAATTCATAAAAATAAGGCTGC (-79 -53) R: TATAAGAATTCACGATATTAACTGGC (End + 26) HbbF2: AGACACTGCTGCTAGAAAGCG (-97 -76) HbbR2: TGGAGTCTAGCTTACAATCCC (End + 29-50)

recA

156

F: GTGGATCTATTGTATTAGATGAAGCTCTTG (170-199) R: GCCAAAGTTCTGAAACATTAACTCCCAAAG (391-362) RecAF GAAGCTATTGAGCTTGCAAGAG (64-85) RecAR GCTACAGAATCAACTACAATCA (464-485)

ospA

261

F: AATAGGTCTAATATTAGCCTTAATAGC (21-47) R: TTGATACTAATGTTTTGCCATCTTCTT (334-308)

rrl-rrf spacer

197

F: CTGCGAGTTCGCGGGAGAG (3' rrf) R: AAGCTCCTAGGCATTCACCATA (5' rrl)

1.3. Sequence Analysis The Clustal W [34] algorithm was used for sequence alignments, and the Mega 3 [35] software for phylogenetic analyses of both individual and concatenated sequences. Distances were calculated by the Jukes and Cantor correction [36] in a pairwise deletion procedure. A similarity table (data not shown) was generated from a distance

140

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matrix (P-distance values) with Excel. Both the Unweighted Pair Group with Mathematical Average (UPGMA) [37] and the Neighbour Joining (NJ) [36] methods were used to build phylogenetic trees. Percentage support values were obtained in a bootstrap procedure.

2. Results and Discussion 2.1. Step 1: Assessment of MLSA in a Taxonomic Purpose by Comparaison with WDDH Data. Application to B. spielmanii Species Validation For this step, 13 isolates from European B. burgdorferi sensu lato pathogenic (n=9) or potentially pathogenic (n=4) species were used, including the type strains of each species. Sequences of the seven loci were either obtained from databases or sequenced for these 13 isolates and for six additional ones from six B. spielmanii isolates. Three of these latter isolates were from ticks collected on dormice trapped in France [11] including the type strain (Pc Eq17), and three strains were from human samples. Phylogenetic trees were drawn for each of the seven loci (data not shown), revealing a clustering in perfect agreement with the taxonomy. This point assesses that lateral transfer did not occur and that the seven loci are therefore adequate for resolving B. burgdorferi sensu lato species. Intraspecific and interspecific genetic distances deduced from sequence alignments were plotted on a diagram together with homologous distances deduced from WDDH [Baranton, published and unpublished] (Figure 1). There was a strong correlation between both types, phenetic versus genetic, distances (black triangles in Figure 1), and an interspecific cutoff of 0.022, corresponding to the maximal distance between the most divergent isolates from B. garinii (the most diverse species) was proposed as an alternative to the 70 % hybridization rate by WDDH.

Figure 1. Correlations of genetic distances by WDDH and MLSA of concatenated sequences.

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Figure 2. Rooted phylogenetic tree with B. lusitaniae as outgroup. Minimum evolution method with distances calculated as Jukes and Cantor. Numbers x 100 indicate bootstrap values on 1000 replications.

Phylogenetic trees have also been drawn for the concatenated sequences from the seven loci for the 19 isolates. Clearly, sequences of the representative isolates of known species do cluster on a branch deeply separated from other ones (Figure 2).

142

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Regarding the MLSA clustering of the six B. spielmanii isolates, obviously they do belong to a single and, up to now, not validated species (maximum intraspecific distance: 0.08; minimum interspecific distance: 0.62). In conclusion, we confirmed that B. spielmanii belongs to a new species associated with rare and narrow host spectrum (dormice and humans only up to now) and transmitted by I. ricinus in Europe [11]. This step led to an accepted publication [11]. 2.2. Step 2: Classification of B. spp. Isolates from California with Borderline WDDH Values Revisited by MLSA. Delineation of Genomospecies 1, 2, and 3 California isolates, which appeared atypical by molecular typing methods [9], were submitted to WDDH, which led to borderline values of DNA relatedness in WDDH experiments. Therefore, they were analyzed in an MLSA procedure. The topologies of the phylogenetic trees of the seven individual loci do not indicate any lateral transfer (data not shown). Distances deduced from MLSA concatenated sequences correlated well with those deduced from WDDH (Figure 1), in spite of some exceptions: grey squares indicate discrepancies between WDDH and MLSA values, but these last values were in complete agreement with 'Tm values, the round circle concerns strains 21123 and CA2, which by every molecular method have been shown to be too divergent to belong to the same species [9] (Figure 2). Moreover, MLSA genetic distances are more precise and therefore more significant than those for WDDH. The 0.022 cutoff value allows genomospecies 1, 2, and 3 to be delineated, in addition to known species B. burgdorferi sensu stricto, B. bissettii, and B. andersonii. Genomospecies 1 comprises CA404, CA443, and CA406 (intraspecific distance: 0.03 minimum interspecific: 0.35). Twentythree additional isolates, of which rrl-rrf intergenic spacer sequences were available in databanks (data not shown), likely belong to this new genomospecies. This genomospecies appears restricted to California and is associated with major hosts: Dipodomys californicus (n=18) and Odocoileus hemionus (n=2). The identified vectors of genomospecies 1 are I. jellisonii (n=4), I. spinipalpis (n=1), and I. pacificus (n=1). Genospecies 2 comprise only CA8 and CA29 (intraspecific distance: 0.01 minimum interspecific: 0.30). CA28 and CA2 isolates cluster together with a genetic distance of 0.015 and do constitute genomospecies 3. Isolates of CA13 do not cluster at distances below 0.21 and seem to continue to constitute an unclassified isolate in this step. 2.3. Step 3: MLSA Study of Atypical European Isolates and General Phylogeny of B. burgdorferi sensu lato Eight isolates from I. ricinus could not be identified by usual molecular methods [38]. In addition, several authors suggested that B. valaisiana exhibit in Europe two groups of isolates that constitute a non-monophyletic species [39]. All these strains, including a B. valaisiana isolate representative of the supposed second group (M07), were studied by MLSA. When looking at the individual trees, we noted that only one of them (rrs gene) showed a clear division into two groups (data not shown). In addition, when considering the tree built with concatenated sequences, we noted that B. valaisiana appeared to constitute a rather homogenous species. However, only one Japanese strain, Am501, representative of a potential 3rd (Asiatic) group [39], was included since we did not have the most divergent Asiatic strains in our collection of strains.

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As regards the European Borrelia spp. isolates, they constitute the single genomospecies 4, although they cluster in two groups separated by distances smaller than 0.022. Since MLSA allows phylogenetic studies, the 42 concatenated sequences representing maximal diversity in both North America and Europe (Figure 2) were studied altogether on a single phylogenetic tree in search of relationships among different lineages. The most striking feature is the high divergence exhibited by the B. lusitaniae species. Two (non-exclusive) hypotheses may be presented. Either B. lusitaniae indeed is an ancient species or this divergence is due to its particular ecological niches. The only reservoir identified for B. lusitaniae in Tunisia is a lizard [40], which, because it is a poïkilotherm, could well be associated with different mechanisms as the bacterium passed from host to tick. Apart from this divergent position of B. lusitaniae, which allows us to consider this species as an outgroup for phylogenetic studies, the main additional point is the division into 2 major clusters: one comprising all the American species and genomospecies, and the other comprising most of the European species. The exceptions are significant, since European isolates of B. burgdorferi sensu stricto cluster with American isolates, as well as all genomospecies and Borrelia spp. from Europe. This confirms that European B. burgdorferi have been imported from North America [41]. The same is true for genomospecies 4. In addition, indeed, genomospecies 4 comprises Californian isolate CA13 (Figure 2), suggesting that the common ancestor of this group of isolates was Californian. Moreover, among the North American cluster, with the exception of B. andersonii, all the remaining species and genomospecies are restricted to California or also present in California, which suggests that B. burgdorferi sensu stricto is a clone that emerged from the population of Borrelia endemic to California and expanded to the East Coast.

3. Conclusions We selected a panel of isolates representing the diversity of the whole B. burgdorferi sensu lato group. Among representative strains of previously delineated species, we include those for which DNA relatedness values were available. Seven loci were chosen for MLSA. Concatenated sequences lead to genetic distances that correlate with WDDH values. Sine these genetic distances are sequence based, they are precise and not subject to experimental variations as are hybridization rates. MLSA offers advantages such as no need to require an extensive strain collection, and results may be compared to a databank. Minute amounts of DNA that are easily transportable are used rather than large volumes of culture. Clustering of B. burgdorferi sensu lato isolates by MLSA led to delineate four new genomospecies, in addition to the eight already known species in both America and Europe. These genomospecies all are closely related to B. burgdorferi sensu stricto, and those from Europe have been imported from North America to Europe, as was the case for B. burgdorferi sensu stricto.

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[22] Wilske B., V. Preac-Mursic, G. Schierz, R. Kuhbeck, A. G. Barbour, and M. Kramer. 1988. Antigenic variability of Borrelia burgdorferi. Ann N Y Acad Sci. 539:126-143. [23] Valsangiacomo, C., T. Balmelli, and J. C. Piffaretti. 1997. A phylogenetic analysis of Borrelia burgdorferi sensu lato based on sequence information from the hbb gene, coding for a histone-like protein. Int J Syst Bacteriol. 47:1–10. [24] Preac-Mursic V., B. Wilske, and G. Schierz. 1986. European Borrelia burgdorferi isolated from humans and ticks: culture conditions and antibiotic susceptibility. Zentralbl Bakteriol Mikrobiol Hyg [A]. 263:112-118. [25] Wang G., A. P. van Dam, J. Dankert. 2000. Two distinct ospA genes among Borrelia valaisiana strains. Res Microbiol. 151:325-331. [26] Godfroid E., C. Min Hu, P. F. Humair, A. Bollen, and L. Gern. 2003. PCR-reverse line blot typing method underscores the genomic heterogeneity of Borrelia valaisiana species and suggests its potential involvement in Lyme disease. J Clin Microbiol. 41:3690-3698. [27] Richter, D., D. B. Schlee, R. Allgöwer, and F. R. Matuschka. 2004. Relationships of a novel Lyme disease spirochete, Borrelia spielmani sp. nov., with Its hosts in Central Europe. Appl Environ Microbiol. 70:6414–6419. [28] Wang, G. Q., A. P. van Dam, and J. Dankert. 1999. Phenotypic and genetic characterization of a novel Borrelia burgdorferi sensu lato isolate from a patient with Lyme borreliosis. J Clin Microbiol. 37:3025– 3028. [29] Theisen M., M. Borre, M. J. Mathiesen, B. Mikkelsen, A. M. Lebech, and K. Hansen. 1995. Evolution of the Borrelia burgdorferi outer surface protein OspC. J Bacteriol. 177:3036-3044. [30] Assous, M. V., D. Postic, G. Paul, P. Névot, and G. Baranton. 1994. Individualisation of two genomic groups among American Borrelia burgdorferi sensu lato strains. FEMS Microbiol Lett. 121:93–98. [31] Casati, S., M. V. Bernasconi, L. Gern, and J. C. Piffaretti. 2004. Diversity within Borrelia. burgdorferi sensu lato genospecies in Switzerland by recA gene sequence. FEMS Microbiol Lett. 238:115–123. [32] Park, H. S., J. H. Lee, E. J. Jeong, S. E. Koh, T. K. Park, W. J. Jang, K. H. Park, B. J. Kim, Y. H. Kook, and S. H. Lee. 2004. Evaluation of groEL gene analysis for identification of Borrelia burgdorferi sensu lato. J Clin Microbiol. 42:1270-1273. [33] Postic, D., C. Edlinger, C. Richaud, F. Grimont, Y. Dufresne, P. Pérolat, G. Baranton, and P. A. D. Grimont. 1990. Two genomic species in Borrelia burgdorferi. Res Microbiol. 141:465–475. [34] Thompson, D., T. J. Gibson, F. Plewniak, F. Jeanmougin, and D. G. Higgins. 1997. The CLUSTAL X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res. 25: 4876–4882. [35] Kumar, S., K. Tamura, and N. Masatoshi 1993. MEGA: Molecular evolutionary genetics analysis, version 1.01. The Pennsylvania State University, University Park, PA 16802. [36] Saitou, N., and M. Nei. 1987. The neighbor-joining method: A new method for reconstructing phylogenetic trees. Mol Biol Evol. 4:406–425. [37] Saitou, N., and M. Nei. 1987. The neighbor-joining method: A new method for reconstructing phylogenetic trees. Mol Biol Evol. 4:406–425. [38] Postic, D., N. Marti Ras, R. S. Lane, P.-F. Humair, M. M. Wittenbrink, and G. Baranton. 1999. Common ancestry of Borrelia burgdorferi sensu lato from North America and Europe. J Clin Microbiol. 37:3010–3012. [39] Ryffel, K., O. Péter, E. Dayer, A. Bretz, and E. Godfroid. 2003. OspA heterogeneity of Borrelia valaisiana confirmed by phenotypic and genotypic analyses. BMC Infect Dis. 3:14. [40] Dsouli, N., H. Younsi-Kabachii, D. Postic, S. Nouira, L. Gern, and A. Bouattour. 2006. Reservoir role of the lizard, Psammodromus algirus, in the transmission cycle of Borrelia burgdorferi sensu lato (Spirochaetacae) in Tunisia. J Med Entomol. In press. [41] Marti Ras, N., D. Postic, M. Foretz, and G. Baranton. 1997. Borrelia burgdorferi sensu stricto, a bacterial species “made in the U.S.A.”? Int J Syst Bacteriol. 47:1112–1117.

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

Diversity and Variability of ProteinEncoding Genes of Borrelia burgdorferi sensu lato and Implications for Pathogenesis and Diagnosis of Lyme Borreliosis in Europe Bettina WILSKE 1, Volker FINGERLE, and Ulrike SCHULTE-SPECHTEL Max von Pettenkofer-Institute, National Reference Center for Borreliae, Germany Abstract. Diversity of Lyme disease spirochetes and their protein encoding genes play a major role in pathogenesis and diagnosis of Lyme borreliosis. In a survey of 72 CSF isolates from Germany, we found 15 (20.5%) Borrelia afzelii, 39 (53.4%) B. garinii, and 18 (24.7%) B. burgdorferi sensu stricto ; among 160 skin isolates, B. afzelii was present in 107 (66.8%), B. garinii in 39 (24.3%), B. burgdorferi sensu stricto in 10 (6.3%), and the new species B. spielmanii in 4 (2.5%). A small number of isolates from synovial fluids (n=6) revealed heterogeneity of the causative strains (2 B. burgdorferi sensu stricto , 2 B. afzelii, and 2 B. garinii). Among 507 PCR-positive ticks from Southern Germany, 22% harboured B. burgdorferi sensu stricto , 25% B. afzelii, 34% B. garinii, 16% B. valaisiana, and 6% B. spielmanii. Sequence identities among major immunodominant proteins (DbpA, VlsE, OspC, OspA, BmpA, p83/100, p58, and flagellin) from the three main human pathogenic species increase from 40–44% to 96–97%. Comparison of dbpA sequences with ospC sequences from a panel of 59 strains revealed all kinds of cross-connections indicating processes of lateral gene transfer. The extent of sequence identities among the dbpA genes decreased from the DNA (67%) to the AA level (44%) about 23%, and ospC sequence identities differed about 10%, an indication that both proteins, but especially DbpA, play a role in immune escape. A combination of four DbpAs (including two different B. garinii DbpAs) was crucial for sensitive serodiagnosis. The use of primarily in vivo-expressed recombinant proteins like OspC, DbpA, and VlsE significantly improved antibody detection. VlsE and DbpA are the most sensitive antigens for IgG antibody detection. OspC is not only the immunodominant protein of the early (IgM) response in the infected host, but it is also crucial for host infectivity. It is controversial to assume that OspC is necessary for borrelial invasion of the tick salivary glands. We found that an OspC-positive B. afzelii wild type strain disseminated from the midgut to the salivary glands, while an OspC-negative mutant was only present in the tick midguts. Colonization of salivary glands by the mutant was restored by complementation of this strain with a plasmid construct that constitutively expresses the wild type ospC. Thus Borrelia strains might also differ in their potential to disseminate from the gut to the ticks’ salivary glands. Keywords. Borrelia burgdorferi, B. afzelii, B. garinii, B. spielmanii, molecular diversity, OspA, OspC, DbpA, VlsE, immunoblot, dissemination in ticks 1 Corresponding Author: Prof. Dr. Bettina Wilske, Max von Pettenkofer-Institute, University of Munich, Pettenkofer-Strasse 9a, D 80336 Munich, Germany; E-mail: [email protected].

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Introduction Diversity of Lyme disease spirochetes as well as diversity and variability of their protein encoding genes play a major role in pathogenesis and are crucial for diagnosis of Lyme borreliosis in Europe. Here we will discuss three major points: 1) the possible association of different Borrelia species with different clinical manifestations of Lyme borreliosis; 2) the implications of this diversity for microbiological diagnosis, especially serodiagnosis; and 3) the variability of expression of borrelial outer surface proteins, specifically OspC expression and its role for dissemination within Ixodes ricinus. In contrast to the United States where only one species, Borrelia burgdorferi sensu stricto, causes Lyme disease, the human pathogenic strains in Europe belong to at least four different species, B. burgdorferi sensu stricto, B. afzelii, B. garinii and the recently described species B. spielmanii (former genospecies A14S) [1, 2]. The different species have been associated with different clinical manifestations of the disease. B. afzelii has a high prevalence among isolates from human skin, especially those from patients with acrodermatitis chronica athrophicans (ACA) [3–6]. Isolates from cerebrospinal fluid most often belong to B. garinii [7–10]. A controversy exists between authors concerning the association of B. burgdorferi sensu stricto with Lyme arthritis [11–13]. In a study presented here, we provide the data of the molecular analysis of 238 human isolates from Germany cultivated in the Pettenkofer-Institute since 1984. We were interested (a) in the distribution of species according to the different human specimens and (b) in addition in the presence and frequency of the new species B. spielmanii among our isolates. Of importance for the conclusions that can be deduced from prevalence studies using human isolates is the distribution of the different species within the tick population. Therefore we investigated a large number of ticks (>2000) from southern Germany for species distribution. In addition, we were interested in the distribution of OspA-types among the respective borreliae from ticks and patients, an important issue for vaccine development. The diversity of target genes for PCR and the diversity of protein encoding genes have important implications for PCR detections of borreliae and serodiagnosis respectively. Here will be discussed strategies to improve serodiagnosis using recombinant proteins derived from different strains and especially those primarily expressed in vivo like OspC, DbpA and VlsE. Differential expression of borrelial proteins facilitate adaptation to and movement within different environments. One important point is which molecular changes are responsible for the dissemination within the tick (from the gut to the salivary glands). Two recent publications report different results concerning the role of OspC for the dissemination within the tick [14, 15]. In our studies we found that an OspC positive B. afzelii wild-type strain disseminated from the gut to the salivary glands, while an OspC negative mutant was only present in the tick guts [16]. Colonization of salivary glands by the mutant was restored by complementation of this strain with a plasmid construct that constitutively expresses the wild-type ospC.

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1. The Diversity of European Isolates from Humans and Ticks A total of 238 human isolates from Germany were analysed for species diversity and OspA-type. After PCR amplification, ospA amplicons were investigated by restriction fragment length polymorphism analysis or DNA sequencing as described [17]. Results for CSF and skin isolates are shown in Figure 1a. Skin

CSF n=72 B. bissetti

B. garinii

(OspA-types 3-8)

5

type 4

type 1

type 5

type 3

B. afzelii

B. afzelii type 3

e4 t yp

B. afzelii B. garinii B. burgd. s.s. B. bissetti

type 1

6

pe ty

B. burgdorferi s.s.

type 7

n=160 B. spielmanii B. burgdorferi s.s.

e typ

typ e6

ty pe 8 7 type

B. garinii

(OspA-types 3-8)

type 8

type 2

type 2

B. afzelii B. garinii B. burgd. s.s. B. spielmanii

15 (20.8 %) 38 (52.8%) 18 (25.0%) 1 (1.4%)

107 (66.8%) 39 (24.3%) 10 (6.3%) 4 (2.5%)

Fingerle, Schulte-Spechtel, Hoffmann, Weber, Wilske, unpublished

Figure 1a. Species and OspA-types in human isolates from Germany (n=232). Detection of B. spielmanii in 4 skin isolates (2.5%). B. garinii comprises the OspA-types 3-8.

B. spielmanii (6%)

B. valaisiana (13%) B. lusitaniae

B. burgdorferi s.s.

type 8

type 7

B. garinii (34%)

type 1 type 6 type 5

type 2 type 4

B. afzelii (25%)

type 3

Figure 1b. Prevalence of different species and OspA-types in ticks from Southern Germany (507 PCRpositive ticks). Detection of B. spielmanii in 28 ticks (6%).

Diversity of borreliae from ticks collected in southern Germany was investigated by PCR from tick material as described [17] (Figure 1b). Analysis of ospA amplicons allows species determination as well as determination of the OspA-type which is

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important for subtyping B. garinii [9]. We found 15 (20.5%) Borrelia afzelii, 39 (53.4%) B. garinii, and 18 (24.7%) B. burgdorferi sensu stricto; among 160 skin isolates, B. afzelii was present in 107 (66.8%), B. garinii in 39 (24.3%), B. burgdorferi sensu stricto in 10 (6.3%), and B. spielmanii in 4 (2.5%). A small number of isolates from synovial fluids (n=6) revealed heterogeneity of the causative strains (2 B. burgdorferi sensu stricto, 2 B. afzelii, and 2 B. garinii, OspA-type ). Comparison of the present data with data from previous studies is shown in Tables 1–4. For skin isolates a clear prevalence of B. afzelii was found in all studies as well as the prevalence of B. garinii in CSF isolates. Table 1. Distribution of Borrelia species among European skin isolates. Author

Total

B. burgdorferi s.s.

B. afzelii

B. garinii

Wilske et al. [18]

68

6%

84%

10%

Ruzic-Sabljic et al. [19]

87

1%

84.5%

14%

Fingerle et al. [unpublished]

160

6%

67%

24%

1

1

2.5% of the total were B. spielmanii.

Table 2. Distribution of Borrelia species among European CSF isolates. Author

Method

Total

B. burgdorferi s.s.

B. afzelii

B. garinii

PCR

12

0%

33%

67%

Ruzic-Sabljic et al. [7]

culture

40

3%

35%

67%

Wilske et al. [10]

culture

37

11%

24%

65%

Fingerle et al. [Unpublished]

culture

72

25%

21%

53%

Eiffert et al. [8]

It was suggested by Eiffert et al. [8] that the distribution of species among strains causing neuroborreliosis reflects the distribution of species among tick isolates from the same area. Table 3 shows the distribution of species and OspA-types among borreliae from ticks and CSF from northern [8] and southern Germany [Fingerle et al., unpublished], respectively. In both studies a high heterogeneity and a similar distribution of species and subtypes were found for borreliae from CSF and ticks, respectively. Notably, in our tick study from southern Germany, B. valaisiana was found in 13% of the ticks but has not been found in material from patients (Figure 1b). Another important finding was the presence of the new species B. spielmanii in 6% of the ticks, a species with pathogenic potential for skin manifestations in humans. With the exception of a study from France [11], borreliae detected by PCR from synovial fluids from European patients with Lyme arthritis are heterogeneous [12, 13] (Table 4). Isolation of B. burgdorferi s.l. from synovial fluid was reported only once in a case from Switzerland. This strain was B. burgdorferi sensu stricto [20]. In our study

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150

on human isolates we could confirm now also by culture that strains causing Lyme arthritis are heterogeneous in Europe (Fingerle et al., Table 4). Table 3. Analysis of culture isolates or PCR amplicons from CSF and ticks. Author

Source

Metho d

Tota l

B. burgdorferi s.s.

B. afzelii

B. garinii

CSF

PCR

12

0%

33%

67%

3, 4, 5, 8

CSF

Culture

72

25%

21%

53%

3, 4, 5, 6, 7, 8

Eiffert et al. [8]

Ticks

Fingerle et al. 1 1

Ticks

PCR

PCR

39

5%

507

28%

22%

25%

67%

34%

B. garinii OspA-type 3, 4, 6

3, 4, 5, 6, 7, 8

B. valaisiana was found in 13% and B. spielmanii in 6% of the ticks but not in CSF from the patients.

Table 4. Analysis of PCR amplicons or isolates from synovial fluid from Lyme arthritis patients. Author

Method

Total

B. burgdorferi s.s.

B. afzelii

B. garinii

Eiffert et al. [13]

PCR

7

3

1

3

Vasiliu et al. [12]

PCR

15

4

5

6

Jaulhac et al. [11]

PCR

10

9

0

1

Culture

6

2

2

2*

Fingerle et al. 1

1

B. garinii strains belonged to OspA-types 2 and 4.

2. Molecular Heterogeneity of Immunodominant Proteins and Development of Recombinant Immunoblots 2.1. MolecularHeterogeneity of Immunodominant Proteins Sequence identities among major immunodominant proteins (DbpA, VlsE, OspC, OspA, BmpA, p83/100, p58, and flagellin) from the three main human pathogenic species increase from 40-44% to 96-97% (Table 5). The most heterogeneous protein was DbpA (decorin binding protein A). Analysis of 25 dbpA sequences from the species B. burgdorferi sensu stricto , 16 from B. afzelii, 40 from B. garinii, and two from the recently described human-pathogenic species B. spielmanii (former genospecies A14S) revealed five distinct DbpA groups [21]. Group I comprises B. burgdorferi sensu stricto and group II B. afzelii, respectively. B. garinii is divided into groups III and IV, whereas the two B. spielmanii strains form a separate group (group V). OspA-types 3, 5, 6, and 7 form group III, and OspA-types 4 and 6 group IV. So OspA-type 6 is present in both B. garinii DbpA groups. In DbpA group III, dbpA

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sequences of OspA-types 6 and 3 are identical. The same is true for dbpA sequences of OspA-types 6 and 4 within DbpA group IV. Comparison of dbpA sequences as well as ospC sequences from a panel of the 59 strains revealed all kinds of cross-connections indicating processes of lateral gene transfer (Figure 2). Indications for lateral gene transfer among OspCs have been Table 5. Sequence identities among major immmunodominant proteins from B. burgdorferi s.s., B. afzelii and B. garinii (comparison of strains B31, PKo and PBi). 1 Protein

DNA Sequences Range (%)

Amino Acid Sequences Range (%)

DbpA*

51–63

40–44

VlsE

65–72

51–56

OspC*

61–77

54–68

OspA*

85–86

78–81

BmpA (p39)*

91–93

89–90

P58*

90–97

90–97

FlaB (flagellin)

94–95

96–97

FlaB int. fragment (aa 129-251)

92–93

92–96

P83/100*

81–87

87–89

1

Sequence identities were calculated without the leader sequence of the lipoproteins.

observed by several authors previously [22–24]. The extent of sequence identities among the dbpA genes decreased from the DNA (67%) to the AA level (44%) about 23%; in the case of the ospC sequence, identities differed about 10%. This is an indication that OspC and especially DbpA play an important role in immune escape. In contrast, flagellin (FlaB) is much more conserved with respect to DNA sequences as compared to AA sequences, which is in accordance with its role as a structural protein. The molecular heterogeneity of DbpA is reflected in antigenic heterogeneity. For groups I to IV several group-specific monoclonal antibodies have been produced (Figure 3a). The antigenic heterogeneity of the DbpAs is also documented by different reactivities with human sera (Figure 3b).

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OspC

100% 95% 90% 85% 80% 75%

1 ZS7 PBre 1 1 PMi PSpm 1 1 PSe 1 B31 1 IA 1 PAli PBoe 1 1 PHo 1 PTa PWag 1 PMu 1 1 JD1 HB19 1 1 297 1 PGl 1 N40 VS461 2 5 PLi 5 PBu PHei 5 6 Tn PWudII 6 PGau 2 2 PKo PWudI 2 2 PBo 2 PLe 2 A91 PLud 2 6 K48 PFiM 6 PHap PSigII 3 PFr 3 PBr 5 PBe 8 PLa 8 PKi PMit 6 6 B29 7 T25 PKuf 7 PBes 7 PRef 7 4 PEi PBar 4 4 PBn PBaeI 4 PMue 4 PScf 4 PWa 4 4 PVo 4 PBi PBet 6 PHez 6 PBol 6 PHC 6

DbpA

100% 90% 80% 70% 60% 50%

70%

100% 99% 100%

81%

Gr. I 99%

79% 77%

73%

79%

100%

78%

100% 100% 99%

Gr. II 81%

98%

79%

75%

72%

81%78%

100% 100%

76%

99% 100%

Gr. III

79%

99%

81%

100%

74%

Gr. V

100%

Gr. IV

99% 100%

92%

ZS7 1 PGl 1 PBre 1 PTa 1 PMi 1 PSpm 1 HB19 1 N40 1 B31 1 IA 1 PAli 1 PBoe 1 PHo 1 PMu 1 PSe 1 PWag 1 JD1 1 1 297 3 PFr PBr 3 6 PHc PHez 6 5 PBe 8 PLa 8 PKi 5 PLi PHei 5 6 B29 5 PBu 7 T25 PKuf 7 PRef 7 PBes 7 VS461 2 2 PBo PWudI 2 2 PKo PGau 2 2 PLe 2 A91 PLud 2 PSigII PHap PWa 4 4 PBn PBar 4 4 PBi 4 PEi PMue 4 PScf 4 4 PVo PBaeI 4 6 K48 PBet 6 PFiM 6 PMit 6 6 TN PBol 6 PWudII 6

40%

99%

98% 100%

99% 93% 100% 96%

100%

74%

97%

89% 59%

100% 94% 99% 100%

91%

99%

100% 94% 99% 100%

97%

44% 86%

89% 100% 93% 99%

59%

98%

100%

51%

99%

77% 62%

Figure 2. Comparison of the DbpA and OspC AA sequence identity trees. Figures behind the strains indicate the corresponding OspA-types. DbpAs are grouped into groups I—V.

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153

a B31 (group I)

PKo (group II)

PBr (group III)

PBi (group IV)

b

B31 (group I)

PKo (group II)

PBr (group III)

PBi (group IV) Figure 3. Reactivity of recombinant DbpAs from B. burgdorferi s.s. (B31), B. afzelii (PKo) and B. garinii (PBi, PBr) using the line immunoblot. a Exemplarily reactiveness of 10 sera from Lyme borreliosis patients and b monoclonal group-specific (groups I–IV) antibodies

2.2. Development of Recombinant Immunoblots Recently we have developed a recombinant immunoblot for detection of antibodies in patients with Lyme borreliosis [25]. We used the line blot technique. In contrast to the Western immunoblot, in this assay antibodies to antigens with identical molecular weight can be easily detected, i.e., homologues of a protein from different strains. The recombinant Borrelia line immunoblot has been based on 18 homologues out of seven different antigens, namely p100, p58, p41i, BmpA, VlsE, OspC, and DbpA (Figure 4). To verify sensitivity and specificity, 85 sera of patients with different manifestations of Lyme borreliosis and 110 control sera were tested for Borrelia specific IgG and IgM antibodies with a previous recombinant Western immunoblot [26]. In the line blot two additional VlsEs and two additional DbpAs were used. Sensitivity increased significantly from 70.6% (Western blot) to 84.7% (line blot) for IgG (p=0.042) and from 40.0% (Western blot) to 73.8% (line blot) for IgM (p<0.005). The increase of sensitivity for IgG detection basically depends on the new line blot technique, whereas for IgM mainly the additional antigens cause the higher sensitivity. VlsE and DbpA are the most sensitive antigens for IgG antibody detection (Table 6). Notably, VlsE of the B. garinii strain PBi displayed the highest sensitivity of all antigens tested for IgG detection. OspC is the immunodominant protein of the early (IgM) response in the infected host and displayed the best reactivity for IgM antibodies. VlsE was the second best antigen for IgM detection and VlsE of strain PBi was the most reactive (Figure 4). The combination of homologues from different strains increases sensititivity and was especially efficient in case of DbpA. Out of 50 patients with acute neuroborreliosis 39 (78.0%), were positive with at least one of the four DbpAs, but only sox (12%) with B. burgdorferi sensu stricto DbpA, 17 (34%) with B.

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154

EM

IgM

EM

NB p100

PKo

p58

PBi

BmpA

B31 PKo PBi

VlsE

PKa2 PKo PBi

OspC p41i DbpA

IgG

NB

ACA

B31 PKo PBi 20047 PKo PBi B31 PKo PBr PBi

Figure 4. Recombinant line immunoblot. Representative IgM blots and IgG blots of patients with Lyme borreliosis. Borrelia strains belong to the following species: B31 and PKa2 to B. burgdorferi s.s., PKo to B. afzelii, PBr to B. garinii OspA-type 3, PBi to B. garinii OspA-type 4, and 20047 to B. garinii unknown OspA-type. Sera were obtained from patients with erythema migrans (EM), early neuroborreliosis (NB), and Acrodermatitis chronica atrophicans (ACA).

afzelii DbpA and 32 (64.0%) with at least one of the two B. garinii DbpAs. Thus, a panel of DbpAs representing the four major DbpA groups of B. burgdorferi sensu stricto , B. afzelii, and B. garinii appears to provide optimal detection rates (Figures 3 and 4). Using the line blot technique, the recombinant IgG immunoblot became even significantly more sensitive than the conventional IgG sonicate immunoblot (i.e., 91.7% versus 68.8% in patients with early neuroborreliosis for the detection of IgG antibodies). Due to its excellent sensitivity and specificity combined with easy evaluation and judgement, this line immunoblot offers a useful improvement for serodiagnosis of Lyme borreliosis. As shown in Table 6, some correlations are seen between stage of the disease and antibody reactivities. For the IgG response VlsE was the immunodominant antigen in all stages (80–100%) whereas OspC is reactive only in 20 to 50%. Other proteins have low reactivities in early manifestations compared to those in late manifestations. This is especially apparent for p58 where reactivities increase from 7% (stage I) to 54% (stage II) and to 95% in stage III.

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Table 6. IgG reactivity of recombinant Borrelia proteins in the line immunoblot in early (EM, erythema migrans; NB, acute neuroborreliosis) and late manifestations (ACA, acrodermatitis chronica atrophicans; AT, arthritis). 1 Group

Sera tested

No. (%) of reactive sera with at least one homologue 2 DbpA

p41i

OspC

VlsE

p58

p100

EM

15

5 (33.3)

1 (6.7)

3 (20.0)

12 (80.0)

1 (6.7)

4 (26.7)

NB II

50

39 (78.0)

17 (34.0)

9 (18.0)

46 (92.0)

27 (54.0)

26 (52.0)

20

18 (90.0)

17 (85.0) 10 (50.0)

20 (100)

19 (95.0)

17 (85.0)

110

4 (3.6)

4 (3.6)

1 (0.9)

2 (1.8)

ACAor AT Negative controls

3

0 (0.0)

0 (0.0)

Based on data from reference [25]. 2 Multiple homologous proteins from different strains were used as antigens in the case of DbpA, p41i, OspC, and VlsE (same as in Figure 3). 3 None of the controls was reactive with more than one Borrelia protein. 1

DbpA reactivities are associated with the predominant causative species in the respective clinical manifestation. The prevalence of reactivity with B. garinii DbpA is in agreement with the fact that B. garinii is prevalent in patients with neuroborreliosis (Table 7). The low reactivity of DbpA derived from B. burgdorferi sensu stricto confirms the findings of Eiffert et al. [13] and Vasiliu et al. [12] that B. burgdorferi sensu stricto is not the prevalent causative agent in European patients with Lyme arthritis. Table 7. Reactivity of Lyme borreliosis sera with recombinant DbpA. Author

Diagnosis

Total

Positive

B. burgdorferi s.s.

B. afzelii

B. garinii

Heikkilä [27]

Neuroborreliosis

10

9

2

4

5

Göttner [25]

Neuroborreliosis

50

39

6

17

32

60

48 (80%)

8 (13%)

21 (35%)

37 (62%)

Heikkilä [28]

Both authors Neuroborreliosis Arthritis

52

51

19

36

38

Göttner [25]

Arthritis

10

10

2

8

8

Both authors Arthritis

62

61 (98%)

21 (34%)

44 (71%)

46 (74%)

3. The Variability of the Expression of Borrelial Outer Surface Protein OspC and Its Role for Dissemination within Ixodes ricinus The outer surface protein OspC is up-regulated in the midgut of the tick during tick feeding on mice or humans [29, 30]. The question is still unanswered whether OspC plays a crucial rule for the dissemination from the gut to the tick salivary glands. Two recent publications report different results concerning the role of OspC in this respect [14, 15]. Pal et al. found that OspC facilitates invasion of I. scapularis salivary glands

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Midgut

Salivary gland

600

500 400

number of borreliae

number of borreliae

500 400 300 200 100 0

0h

12h

24h

48h

Time after capillary feeding

96h

300 200 100 0

0h

12h

24h

48h

96h

Time after capillary feeding

PKo wild type

PKo 345 OspC-negative PKo 345 OspC-positive

Figure 5. Detection of B. afzelii in capillary-fed I. ricinus. Ticks were infected with OspC-positive borreliae (PKo wild type and cPKo345pBSV2/ospC+) and OspC-negative borreliae (cPKo345ospC-). Tick organs were dissected after different time intervals (two ticks per time point), and borreliae contained in the organs were counted after immunostaining with monoclonal antibodies against OspA and OspC.

with B. burgdorferi sensu stricto (strain B31 used for infection) [15]. In contrast, Grimm et al. described that OspC is strictly required to infect mice, but not for migration of B. burgdorferi sensu stricto (strain N40 used for infection) into the salivary glands (14). We have studied migration from tick gut to salivary glands using B. afzelii strain PKo (OspC-positive) and an OspC-negative clone cPKo345ospC(mutation with an insertion of guanine at position 200 within the ospC gene leading to a stop codon at position 220). The OspC-negative clone (cPKo345ospC-) was not able to disseminate into the salivary glands of I. ricinus after capillary feeding whereas the wild-type strain did [16]. In a recent experiment we investigated the ability of cPKo345 to migrate to the tick salivary glands after recomplementation with ospC [Goettner et al., unpublished]. The OspC-negative clone cPKo345ospC- was successfully transformed with the shuttle vector pBSV2 containing the wild-type ospC. Unfed Ixodes ricinus nymphs were artificially infected by capillary feeding either with B. afzelii wild type (strain PKo), the B. afzelii OspC-negative mutant (cPKo345ospC-), or the ospC-complemented OspC-positive clone (cPKo345pBSV2/ospC+). Tick midguts and salivary glands were investigated after different time intervals (Figure 5) for the presence of borreliae and for OspA and OspC by immunofluorescence staining with monoclonal antibodies. While the B. afzelii wild type strain, which exhibits abundant OspC on its surface, disseminated to the salivary glands, the OspC-negative mutant was only present in the tick midguts. The ospCcomplemented clone, which constitutively expresses the wild type ospC, was again able to colonize the salivary glands. The time course of dissemination to the salivary glands was similar between the wild type and the ospC-recomplemented borreliae. Possible explanations for the differences among the previous and present studies may include different dissemination behaviour of different Borrelia species and even

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strains belonging to the same species, the different modes of tick infection, and influence of different tick species. In a previous study using capillary fed I. ricinus nymphs, we showed that Osp composition patterns, as well as dissemination dynamics, may vary even among strains belonging to the same species [16]. Infection procedures may play also a role for the different outcomes of the three studies. Pal et al. [15] infected the ticks comparable to the present study by capillary injection of a B. burgdorferi sensu stricto strain 297 clone into the rectal sac, while in the study of Grimm et al. [14] I. scapularis ticks were infected by immersing them in a B. burgdorferi sensu stricto strain B31 clone culture. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14]

[15]

Wang, G., A. P. van Dam, I. Schwartz, and J. Dankert. 1999. Molecular typing of Borrelia burgdorferi sensu lato: taxonomic, epidemiological, and clinical implications. Clin.Microbiol.Rev. 12:633–653. Richter, D., D. B. Schlee, R. Allgower, and F. R. Matuschka. 2004. Relationships of a novel Lyme disease spirochete, Borrelia spielmani sp. nov., with its hosts in Central Europe. Appl.Environ.Microbiol. 70:6414–6419. Canica, M. M., F. Nato, M. L. du, J. C. Mazie, G. Baranton, and D. Postic. 1993. Monoclonal antibodies for identification of Borrelia afzelii sp. nov. associated with late cutaneous manifestations of Lyme borreliosis. Scand.J.Infect.Dis. 25:441–448. Ohlenbusch, A., F. R. Matuschka, D. Richter, H. J. Christen, R. Thomssen, A. Spielman, and H. Eiffert. 1996. Etiology of the acrodermatitis chronica atrophicans lesion in Lyme disease. J.Infect.Dis. 174:421–423. Busch, U., C. Hizo-Teufel, R. Böhmer, V. Fingerle, D. Rössler, B. Wilske, and V. Preac-Mursic. 1996. Borrelia burgdorferi sensu lato strains isolated from cutaneous Lyme borreliosis biopsies differentiated by pulsed-field gel electrophoresis. Scand.J.Infect.Dis. 28:583–589. Maraspin, V., E. Ruzic-Sabljic, J. Cimperman, S. Lotric-Furlan, T. Jurca, R. N. Picken, and F. Strle. 2001. Isolation of Borrelia burgdorferi sensu lato from blood of patients with erythema migrans. Infection 29:65–70. Ruzic-Sabljic, E., S. Lotric-Furlan, V. Maraspin, J. Cimperman, D. Pleterski-Rigler, and F. Strle. 2001. Analysis of Borrelia burgdorferi sensu lato isolated from cerebrospinal fluid. APMIS 109:707–713. Eiffert, H., A. Ohlenbusch, H. J. Christen, R. Thomssen, A. Spielman, and F. R. Matuschka. 1995. Nondifferentiation between Lyme disease spirochetes from vector ticks and human cerebrospinal fluid. J.Infect.Dis. 171:476–479. Wilske, B., V. Preac-Mursic, U. B. Göbel, B. Graf, S. Jauris, E. Soutschek, E. Schwab, and G. Zumstein. 1993. An OspA serotyping system for Borrelia burgdorferi based on reactivity with monoclonal antibodies and OspA sequence analysis. J.Clin.Microbiol. 31:340–350. Wilske, B., U. Busch, H. Eiffert, V. Fingerle, H. W. Pfister, D. Rössler, and V. Preac-Mursic. 1996. Diversity of OspA and OspC among cerebrospinal fluid isolates of Borrelia burgdorferi sensu lato from patients with neuroborreliosis in Germany. Med.Microbiol.Immunol.(Berl) 184:195–201. Jaulhac, B., R. Heller, F. X. Limbach, Y. Hansmann, D. Lipsker, H. Monteil, J. Sibilia, and Y. Piemont. 2000. Direct molecular typing of Borrelia burgdorferi sensu lato species in synovial samples from patients with lyme arthritis. J.Clin.Microbiol. 38:1895–1900. Vasiliu, V., P. Herzer, D. Rössler, G. Lehnert, and B. Wilske. 1998. Heterogeneity of Borrelia burgdorferi sensu lato demonstrated by an ospA-type-specific PCR in synovial fluid from patients with Lyme arthritis. Med.Microbiol.Immunol.(Berl) 187:97–102. Eiffert, H., A. Karsten, R. Thomssen, and H. J. Christen. 1998. Characterization of Borrelia burgdorferi strains in Lyme arthritis. Scand.J.Infect.Dis. 30:265–268. Grimm, D., K. Tilly, R. Byram, P. E. Stewart, J. G. Krum, D. M. Bueschel, T. G. Schwan, P. F. Policastro, A. F. Elias, and P. A. Rosa. 2004. Outer-surface protein C of the Lyme disease spirochete: a protein induced in ticks for infection of mammals. Proc.Natl.Acad.Sci.U.S.A 101:3142–3147. Pal, U., X. Yang, M. Chen, L. K. Bockenstedt, J. F. Anderson, R. A. Flavell, M. V. Norgard, and E. Fikrig. 2004. OspC facilitates Borrelia burgdorferi invasion of Ixodes scapularis salivary glands. J.Clin.Invest 113:220–230.

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B. Wilske et al. / Diversity and Variability of Protein-Encoding Genes Fingerle, V., S. Rauser, B. Hammer, O. Kahl, C. Heimerl, U. Schulte-Spechtel, L. Gern, and B. Wilske. 2002. Dynamics of dissemination and outer surface protein expression of different European Borrelia burgdorferi sensu lato strains in artificially infected Ixodes ricinus nymphs. J.Clin.Microbiol. 40:1456–1463. Michel, H., B. Wilske, G. Hettche, G. Goettner, C. Heimerl, U. Reischl, U. Schulte-Spechtel, and V. Fingerle. 2003. An ospA-polymerase chain reaction/restriction fragment length polymorphism-based method for sensitive detection and reliable differentiation of all European Borrelia burgdorferi sensu lato species and OspA types. Med.Microbiol.Immunol.(Berl) 193:219–226. Wilske, B., U. Busch, V. Fingerle, S. Jauris-Heipke, M. Preac, V, D. Rössler, and G. Will. 1996. Immunological and molecular variability of OspA and OspC. Implications for Borrelia vaccine development. Infection 24:208–212. Ruzic-Sabljic, E., F. Strle, J. Cimperman, V. Maraspin, S. Lotric-Furlan, and D. Pleterski-Rigler. 2000. Characterisation of Borrelia burgdorferi sensu lato strains isolated from patients with skin manifestations of Lyme borreliosis residing in Slovenia. J.Med.Microbiol. 49:47–53. Schmidli, J., T. Hunziker, P. Moesli, and U. B. Schaad. 1988. Cultivation of Borrelia burgdorferi from joint fluid three months after treatment of facial palsy due to Lyme borreliosis. J.Infect.Dis. 158:905–906. Schulte-Spechtel, U., V. Fingerle, G. Goettner, S. Rogge, and B. Wilske. 2005. Molecular analysis of decorin binding protein A (DbpA) reveals five major groups among European Borrelia burgdorferi sensu lato strains with impact for the development of serological assays and indicates lateral gene transfer of the dbpA gene. Int J Med Microbiol (in press). Jauris-Heipke, S., G. Liegl, V. Preac-Mursic, D. Rössler, E. Schwab, E. Soutschek, G. Will, and B. Wilske. 1995. Molecular analysis of genes encoding outer surface protein C (OspC) of Borrelia burgdorferi sensu lato: relationship to ospA genotype and evidence of lateral gene exchange of ospC. J.Clin.Microbiol. 33:1860–1866. Livey, I., C. P. Gibbs, R. Schuster, and F. Dorner. 1995. Evidence for lateral transfer and recombination in OspC variation in Lyme disease Borrelia. Mol.Microbiol. 18:257-269. Dykhuizen, D. E. and G. Baranton. 2001. The implications of a low rate of horizontal transfer in Borrelia. Trends Microbiol. 9:344–350. Goettner, G., U. Schulte-Spechtel, R. Hillermann, G. Liegl, B. Wilske, and V. Fingerle. 2005. Improvement of Lyme borreliosis serodiagnosis by a newly developed recombinant immunoglobulin G (IgG) and IgM line immunoblot assay and addition of VlsE and DbpA homologues. J Clin.Microbiol 43:3602–3609. Schulte-Spechtel, U., G. Lehnert, G. Liegl, V. Fingerle, C. Heimerl, B. Johnson, and B. Wilske. 2004. Significant improvement of the recombinant Borrelia IgG immunoblot for serodiagnosis of early neuroborreliosis. Int.J.Med.Microbiol. 293 Suppl 37:158–160. Heikkila, T., I. Seppala, H. Saxen, J. Panelius, H. Yrjanainen, and P. Lahdenne. 2002. Speciesspecific serodiagnosis of Lyme arthritis and neuroborreliosis due to Borrelia burgdorferi sensu stricto, B. afzelii, and B. garinii by using decorin binding protein A. J.Clin.Microbiol. 40:453–460. Heikkila, T., H. I. Huppertz, I. Seppala, H. Sillanpaa, H. Saxen, and P. Lahdenne. 2003. Recombinant or peptide antigens in the serology of Lyme arthritis in children. J.Infect.Dis. 187:1888–1894. Fingerle, V., G. Liegl, U. Munderloh, and B. Wilske. 1998. Expression of outer surface proteins A and C of Borrelia burgdorferi in Ixodes ricinus ticks removed from humans. Med.Microbiol.Immunol.(Berl) 187:121–126. Schwan, T. G., J. Piesman, W. T. Golde, M. C. Dolan, and P. A. Rosa. 1995. Induction of an outer surface protein on Borrelia burgdorferi during tick feeding. Proc.Natl.Acad.Sci.U.S.A 92:2909– 2913.

Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Are Borrelia recurrentis and Borrelia duttonii the Same Spirochaete? Sally J. CUTLER a,1 , Julie C. SCOTT a and David J. M. WRIGHT b Veterinary Laboratories Agency, Addlestone, Surrey, KT15 3NB, UK b Imperial College of Science, Technology & Medicine, London, SW7 2AZ, UK a

Abstract. Relapsing fever Borrelia challenge microbiological typing as, unlike other microbes, they possess segmented genomes maintaining essential genes on large linear plasmids. Antigenic variation further complicates typing. Intragenic spacer (IGS, between 16S-23S genes), provides resolution among Lyme-associated and some relapsing fever spirochaetes. When applied to East African relapsing fever borreliae, two and four types were found respectively among Borrelia recurrentis and B. duttonii. However, IGS typing was unable to discriminate between the tick- and louse-borne forms of disease, raising the question as to whether these are indeed separate species. To address this question, further genes were sequenced to produce a multi-locus approach to resolve whether these are either a single or different species. Various housekeeping genes were selected from data deposited for B. hermsii (limited sequence information exists for either B. recurrentis or B. duttonii). Of selected targets, sufficient data was produced only from glpQ. Further genes analysed included flaB, rrs rDNA, and P66 outer membrane protein. Sequence comparison of multiple genes was undertaken, but restricted through the limited number of available isolates of these notoriously fastidious organisms that until recently were considered non-cultivable. Whereas the IGS typing was applied to a range of clinical isolates, patient blood samples and arthropod vectors, other genes were sequenced only from cultivable strains, potentially introducing a bias to the results. Our data highlights the remarkable similarity between these Borrelia with only minor differences at the nucleotide level. Collectively, this suggests a common ancestral lineage for these spirochaetes, with the limited differences revealed at the nucleotide level from these cultivable strains being able to divide both “species” into separate clades; however, it must be stressed that these differences ranged from 2 to 10 nucleotides depending on the gene used. It is more likely that these are clades of the same species, which have accumulated adaptive changes through time and pressures of different vector transmission. In contrast, the IGS sequence, being non-coding, is not under such selection and in consequence probably reflects changes accumulated over time alone, but without the constraints of producing functional gene products. Full genomic sequence analysis should reveal further insights into the taxonomic relationship between these microbes and elucidate the molecular basis of arthropod competence and pathogenicity among these spirochaetes. Keywords: Borrelia recurrentis; Borrelia duttonii; relapsing fever spirochaetes; louse-borne relapsing fever; tick-borne relapsing fever; phylogeny.

1 Corresponding Author: Veterinary Laboratories Agency, Addlestone, Surrey, KT15 3NB, UK; Phone: +44 1932 357807; Fax: +44 1932 357423; E-mail: [email protected].

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Introduction Though today it is believed that relapsing fever spirochaetes in the Old World occupy distinct geographical regions, this was not always the case. Prior to the 1960’s, major epidemics of the louse-borne form of the disease were responsible for significant worldwide morbidity and mortality. Since then, the disease persists only in Ethiopia, occasionally spilling into neighboring countries such as Sudan [1, 2]. The tick-borne varieties have also resulted in a significant public health burden. David Livingstone, while on his epic travels through Africa, noted the disease burden associated with ticks. In more recent times, it is now believed that Borrelia crocidurae persists in Western regions of Africa with countries such as Senegal considered endemic [3], while B. duttonii persists in East Africa, particularly in regions of Tanzania [4]. Early investigations questioned the host and vector specificities of relapsing fever spirochaetes, with several early studies allowing ticks to feed on patients with the louse-borne form of the disease and subsequently letting these ticks to feed on nonfebrile individuals. These studies were unable to establish conclusively whether these spirochaetes could be transmitted by alternative arthropod vectors. Similarly, different tick species were used to assess the vector specificity for the different tick-borne relapsing fever borreliae. Many of these early investigations were hampered by inability to reliably identify the organisms used, lacked cultivable strains, and had no means of assessing the immune status of recipient hosts. Speciation of the causative relapsing fever spirochaete has since been reliant on the geographical region where infection occurred and on the vector responsible for transmission. Only recently have molecular approaches been applied to identify these spirochaetes The relapsing fever spirochaetes present challenges to many molecular microbiological typing methods through their possession of segmented genomes carried on large linear plasmids. Inter- and intragenomic recombination associated with antigenic variation may result in size changes of plasmids and duplication of DNA, invalidating their use as typing tools [5]. In consequence, molecular typing of these spirochaetes has largely been reliant on gene amplification and sequencing. The rrs gene has highlighted the remarkable similarity among those relapsing fever species from the Old World, with only limited nucleotide differences discriminating between the known species. Others have utilized this approach but with the flagellin gene, flaB, and similarly found only minor differences between species. More recently, non-coding intragenic spacer DNA has been shown to be particularly valuable for characterizing relapsing fever spirochaetes [6, 7]. Despite being able to provide a means of subspecies grouping, a surprising finding was the failure of this approach to be able to differentiate B. recurrentis and B. duttonii [7]. To further investigate this apparent lack of differentiation between these two species, four additional gene targets were assessed for these isolates.

1. Materials and Methods We used various gene targets to analyse the phylogenetic relationships among isolates of B. recurrentis and B. duttonii to address the differences between these spirochaetes. These included targets previously shown to have utility when examining these spirochaetes (rrs, flaB); non-coding regions expected to show increased variability partial intragenic spacer (IGS) target residing between the 16S-23S rDNA genes (rrs-

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rrlA); surface exposed protein (p66); and other essential gene targets where sequence data existed for related spirochaetes that may enable amplification through sufficient conservation. 1.1. Isolates East African relapsing fever borreliae were isolated from patients in endemic areas for louse-borne relapsing fever (Addis Ababa, Ethiopia) and tick-borne relapsing fever (Mvumi, Tanzania) [4, 8, 9]. These organisms were cultivated using BSKII medium, stored at í700C for long-term storage, and revived prior to investigation [10]. 1.2. PCR The primer sequences used are shown in Table 1. All primers were supplied by Sigma Genosys (UK). PCR amplicons were resolved using 1% agarose gels. Products were cleaned and sequenced directly or bands excised, and DNA purified using a Wizard SV gel and PCR cleanup system (Promega).

Table 1. Primers used in study. Locus

Forward Primers (5' to 3')

Reverse Primers (5' to 3')

IGS Outer

GTATGTTTAGTGAGGGGGGTG

GGATCATAGCTCAGGTGGTTAG

IGS Inner

AGGGGGGTGAAGTCGTAACAAG

GTCTGATAAACCTGAGGTCGGA

16S rRNA (FD3-UniB)

AGAGTTTGATCCTGGCTTAG

T(AC)AAGGAGGTGATCCAGC

16S rRNA Fragment 1 (FD3-500R)

AGAGTTTGATCCTGGCTTAG

CTGCTGGCACGTAATTAGCC

16S rRNA Fragment 2 (400F1050R)

GGAGCGACACTGCGTG

CACGAGCTGACGACA

16S rRNA Fragment 3 (800F-rD1R)

ATTAGATACCCTGGTAG

AAGGAGGTGATCCAGCC

glpQ Fragment 1

TAATAATGTTTGCAATAAGTAC

CAATATTTTTCCCTGTGCTTTT

glpQ Fragment 2

CCAATATACCCTAACCGTTTTC

CTTTATTGATATATCAACAAAG

P66 (P6651P6631)

AGTGATTTTTCTATACTTGGACAC

GTTAATTTGATTAAGTTKTCTAGTTCT

flaB (Flf1-Flr1)

CGTGATGATCATAAATCATAATACG

CCAAGCTCTTCAGCTGTTCTTAC

flaB (Flf2-Flr2)

ACATATTCAGATGCAGACAGAGGT

CATATTGAGGTACTTGATTTGC

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1.3. IGS A Borrelia-specific nested PCR designed to amplify the 16S-23S intragenic spacer (IGS) region was used as described [11]. The outer primers are anchored in the 3' end of the rrs gene and the ileT genes, respectively, with internally nested inner primers. 1.4. rrs gene The16S rRNA rrs gene was amplified initially using primers FD3 and UniB using 94°C for one minute, 54°C for one minute and 72°C for one minute and 30 seconds for a total of 45 cycles. Overlapping contigs were amplified using further primers described by others [12]. 1.5. Flagellin Flagellin was amplified using two overlapping fragments. Annealing temperatures of 56°C and 52°C were used respectively with 35 cycles of one minute for denaturation at 94°C and annealing and extension at 72°C. 1.6. p66 Outer Membrane Protein Approximately a 750 bp region of the p66 outer surface protein was amplified using primers kindly described by Alan Barbour. These should include the loop region, hydrophobic flanking regions of this protein. Cycling conditions used were 94°C for one minute, 40°C for two minutes, and 72°C for two minutes for a total of 35 cycles. 1.7. Glycerophosphodiester Phosphodiesterase GlpQ The glycerophosphodiester phosphodiesterase GlpQ gene for four B. recurrentis strains had already been submitted (AF247152íAF247155). These sequences, together with those deposited for other relapsing fever spirochaetes, were aligned, and primers designed against homologous regions. The target gene was amplified using two overlapping fragments. Amplification was achieved using 35 cycles of one minute for denaturation at 94°C, annealing at 48°C, and extension at 72°C. 1.8. DNA Sequencing Purified DNA was either directly sequenced or cloned into pGEMT-easy (Promega) before being sequenced. Sequencing reactions were performed using BigDye terminator v3.1 cycle sequencing kit (Applied Biosystems) according to manufacturer’s recommendations and analysed on an Applied Biosystems Genetic Analyzer. 1.9. Data Analysis Nucleotide sequences were analysed using Chromas (version 1.45) and DNA Star software (Lasergene 6). Multiple alignments were performed using ClustalW. Results produced by IGS fragment typing were compared with those using the rrs gene using sequences held in GenBank.

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1.10. Phylogenetic Trees The phylogenetic relationships of sequences were compared using Mega software (version 3) and Neighbor-Joining methods for compilation of the trees. A bootstrap value of 250 was used to determine confidence in tree drawing parameters.

2. Results Only assays for flagellin, p66, 16S rRNA, IGS, and glpQ successfully generated amplicons from sufficient isolates to allow phylogenetic comparisons. Attempts to compare other gene targets through homology with sequences generated from B. hermsii were unsuccessful in amplifying targets from our isolates for recB; recC; bdrB; fruK; fruA2; and hexoV. A comparative summary of those genes successfully amplified is given in Table 2. Table 2. Summary of relationships within species and between species of African relapsing fever borreliae. Locus (fragment bp)

% Homology B. recurrentis1

% Homology B. duttonii1

% Homology between B. recurrentis & B. duttonii1

% Homology between B. recurrentis, B. duttonii & B. crocidurae1

flaB (981 bp)

100%

99.9%

99.8%

99.6%

IGS (587 bp)

99.66%

97.44%1

97.27%1

87.56%1

glpQ (932 bp)

99.79%

100%

99.79%

99.25%

16S rRNA (1525 bp)

99.9%

100%

99.74%

99.54%

Outer Membrane p66 (650 bp)

100%

100%

98.46%

98.46%

99.94%

100%

99.42

NT3

Overall Concatenated (4642 bp) 1 2 3

Deletions not scored as a difference. Includes uncultivated sequence types. Not tested as full concatenated sequence was not available on any single isolates.

2.1. Intragenic Spacer Amplifications Isolates of B. duttonii yielded a 587 bp portion of the IGS that when compared showed that these were all identical. The strain Ly was selected to represent these isolates and has been used by others as the representative for B. duttonii [13, 14]. Though further types have been described, these were detected only in direct amplifications from ticks described elsewhere [7] but were not represented among cultivable isolates.

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B.duttonii type II DQ000280 66 98 99 100

B.duttonii type III DQ000281 B.recurrentis type I DQ000277 B.recurrentis type II DQ000278 B.duttonii type I DQ000279

B.duttonii type IV DQ000282 B.crocidurae DQ000287 B.crocidurae-like DQ00028

0.01

Figure 1. Relationship of B. duttonii and B. recurrentis with their closest related species, B. crocidurae.

When applied to B. recurrentis, IGS fragment sequencing was able to group these into two types that differed by two nucleotides. Of the 18 isolates A1 through to A10, A17 and A18 comprised one group (type I), while isolates A11 to A16 gave a second profile (type II). The phylogeny based on IGS is shown in Figure 1. This figure includes sequences from the non-cultivable strains for reference, but it must be stressed that only those cultivable isolates (B. duttonii type I) were assessed using other gene targets. 2.2. Ribosomal 16S rDNA Comparison of a 1525 portion of the rrs gene sequences confirmed the difference between the two groups of B. recurrentis, however, using this target, only with a single nucleotide difference (Figure 2). Whereas the IGS fragment analysis produced different profiles within species, the rrs gene sequences, with the exception of the B. recurrentis types above, produced single clusters for each relapsing fever species (see Figure 2). Differences between these species were small, with just four nucleotide difference between B. recurrentis and B. duttonii; and six nucleotides differentiating between B. recurrentis and B. crocidurae; and only two nucleotides between B. duttonii and B. crocidurae. The sequence for the rrs gene of B. recurrentis isolate A2 generated in this study was given the accession number DQ346813. 2.3. Flagellin Partial flagellin flaB gene was sequenced from B. recurrentis isolates A1-A18 and B. duttonii isolates Ku; Ly; and Ma. When sequences were compared within species, total homology was demonstrated over almost the entire sequence. When both B. recurrentis and B. duttonii were compared over the full length of this 983/4 bp sequence, only a two adjacent nucleotide difference distinguished these species. At this position, these two adjacent nucleotides gave a diagnostic signature that may be of value for identification with B. recurrentis giving AA; B. duttonii GC; while B. crocidurae gave

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B.duttonii AB113315 B.hispanica U42294 B.hispanica DQ057988 B.recurrentis A11 AF107361 B.recurrentis A16 AF107362 B.recurrentis A4 AF107356 B.recurrentis A17 AF107360 B.recurrentis U42300 B.recurrentis A2 DQ346813 B.recurrentis A10 AF107359 B.recurrentis A5 AF107357 B.recurrentis A8 AF107358 B.recurrentis A1 AF107367 B.duttonii UESV 334RWA U42298 B.duttonii Ly AF107364 B.duttonii Ku AF107363 B.duttonii UR BD94MIT U42293 B.duttonii UESV 117DUTT U42288 B.duttonii Ma AF107366 B.duttonii La AF107365 B.crocidurae UESV MER U42291 B.crocidurae UESV 1096TEN U423 B.crocidurae DQ057989 B.crocidurae UESV 1045 U42290 B.crocidurae UESV 1043 U42295 B.crocidurae UESV 523SIS U4230 B.crocidurae UESV 1040DAK U422 B.crocidurae DQ057990 B.crocidurae UESV 917BAR U4228 B.crocidurae UESV 626BAN U4228 B.crocidurae U42286

0 .0 0 2

Figure 2. Phylogenetic relationship of B. duttonii and B. recurrentis based on 16S rDNA gene, rrs. Inclusion of other species enables a broader appreciation of the phylogenetic position of these spirochaetes.

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GA. Interestingly, B. duttonii strain 406K (D82859) gave a GA signature at this location, thus resembling B. crocidurae. Over the sequence length studied, one nucleotide was unique to B. recurrentis, one to B. duttonii (absent from D82859) and two specific for B. crocidurae. The phylogenetic relationship of flagellin sequences, together with available sequences from GenBank, are depicted in Figure 3. The two sequences for B. crocidurae compared from GenBank were divergent over 14 nucleotides, illustrated in Figure 3 where the sequence for U28496 falls away from clades containing other sequences types. This sequence was not included in the overall comparison of similarity given in Table 2. Sequences for B. recurrentis A1-A18 were given accession numbers DQ346814-DQ346831, while B. duttonii strains Ku, Ly, and Ma had the numbers DQ346837; DQ346833 and DQ346835 respectively. 2.4. Glycerophosphodiester Phosphodiesterase GlpQ Partial glpQ gene products were sequenced for eight isolates of B. recurrentis (A1; A8; A10; A11; A14; A15; A16; and A17 banked with numbers DQ346777-DQ346784 respectively) and for four B. duttonii isolates (Wi; La; Ly; and Ku banked with numbers DQ346785-DQ346788 respectively). These sequences were 932 bp and 938 bp long respectively. Alignment of these sequences revealed total homology among either B. recurrentis or B. duttonii isolates, and when each was compared with the other, only two nucleotide differences were found between these spirochaetes. When the B. recurrentis isolates were compared with those sequences already deposited in GenBank, only slight variation was found between strains at two positions (Figure 4). Thus the variation seen between different isolates of B. recurrentis was equal to that seen between B. recurrentis and B. duttonii. 2.5. P66 Outer Membrane Protein Although surface exposed, and thus subject to environmental pressures that could result in variability, the p66 membrane protein has been used to examine spirochaetes [15]. The primers successfully amplified a 683 bp portion of this gene. The fragment isolated from all isolates of B. duttonii showed sequence homology, as did those from B. recurrentis. When these sequences were compared with each other 10 bp’s differed between these two species. Similarly, when the other African relapsing fever spirochaete, B. crocidurae, was compared with either B. duttonii or B. recurrentis, each spirochaete differed from each other by 10 nucleotides. The equivalent level of differentiation between these spirochaetes can be seen in Figure 5. Validity of the sequence data produced in this study was gained through comparison with a sequence already deposited with GenBank for B. recurrentis. No data could be found for P66 for B. duttonii. Sequences generated in this study were given the accession numbers of DQ346789-DQ346806 for B. recurrentis isolates A1-A18 and DQ346807-DQ346812 for B. duttonii isolates Ly, Lw, La, Ku, Ma and Wi.

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B recurrentis A12 DQ346825 B recurrentis A7 DQ346820 B recurrentis A11 DQ346824 B recurrentis A18 DQ346831 B recurrentis A10 DQ346823 B recurrentis A5 DQ346818 B recurrentis A17 DQ346830 B recurrentis A6 DQ346819 B recurrentis A13 DQ346826 61

B recurrentis A3 DQ346816 B recurrentis A9 DQ346822 B recurrentis AY604984 B recurrentis A16 DQ346829 B recurrentis A8 DQ346821 B recurrentis A2 DQ346815

47

B recurrentis A1 DQ346814 B recurrentis A4 DQ346817 B recurrentis D86618 B recurrentis A14 DQ346827

89

B recurrentis A15 DQ346828 B duttonii Ku DQ346837 B duttonii Ly DQ346833 66

B duttonii Ma DQ346835 B duttonii D82859 B crocidurae X75204 B crocidurae U28496

0 .0 0 2

Figure 3. Neighbor Joining tree of flagellin flaB sequences.

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B. crocidurae AF247151 B. recurrentis A16 DQ346783 B. recurrentis A11 DQ346780 B. recurrentis A17 DQ346784 B. recurrentis A15 DQ346782 B. recurrentis AF247154 64

60

B. recurrentis AF247155

B. recurrentis A10 DQ346779 B. recurrentis A8 DQ346778 B. recurrentis AF247153 B. recurrentis A1 DQ346777 B. recurrentis AF247152 B. recurrentis A14 DQ346781 B. duttonii Wi DQ346785 B. duttonii Ku DQ346788 65

B. duttonii La DQ346786 B. duttonii Ly DQ346787

0 .0 0 0 5

Figure 4. Neighbor Joining tree for glpQ sequences for African relapsing fever borreliae.

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B recurrentis A17 DQ346805 B recurrentis A4 DQ346792 B recurrentis A16 DQ346804 B recurrentis A1 DQ346789 B recurrentis A15 DQ346803 B recurrentis A6 DQ346794 B recurrentis A13 DQ346801 B recurrentis A9 DQ346797 B recurrentis A2 DQ346790 B recurrentis A12 DQ346800 B recurrentis A8 DQ346796 B recurrentis A5 DQ346793 B recurrentis A3 DQ346791 B recurrentis A14 DQ346802 B recurrentis A11 DQ346799 B recurrentis A10 DQ346798 B recurrentis AF228024 B recurrentis A18 DQ346806 B recurrentis A7 DQ346795 B crocidurae AF125321 B duttonii Ku DQ346810 B duttonii La DQ346809 B duttonii Lw DQ346808 B duttonii Ma DQ346811 B duttonii Ly DQ346807 B duttonii Wi DQ346812

0 .0 0 2

Figure 5. Phylogeny of African relapsing fever borreliae using p66 outer membrane protein.

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84

B. recurrentis A11 B. recurrentis A16

B. recurrentis A1 B. recurrentis A10 61

B. recurrentis A8 B. recurrentis A17 B. duttonii Ku B. duttonii Ly 100

B. duttonii La

0 .0 0 0 5

Figure 6. Neighbor-joining phylogenetic tree of concatenated DNA sequences for isolates where all five loci were examined (bootstrap analysis using 250 replicates).

Taken together, assuming that polymorphisms have resulted from co-evolution reflecting the genomic background of these spirochetes and excluding environmental selective pressure, concatenation of sequences results in the phylogenetic relationship portrayed in Figure 6. The overall sequence homology between B. recurrentis and B. duttonii was 99.42% among those isolates with full concatenated sequences. These findings show the linkage disequilibrium of these different markers. However, these findings fail to conclusively demonstrate whether or not these two species should remain as such, or be considered as clones of a single species.

3. Discussion This study looked at the differences between various gene targets to elucidate whether B. recurrentis and B. duttonii were different species or should be better considered as clones of a single species. The targets chosen included surface exposed outer membrane protein, p66, likely to be under selective pressure and thus predicted to show greater variability; partial rrs-rrl intragenic spacer that as non-coding, would also be predicted to have greater diversity; flagellin flaB and the rrs genes, both used to address phylogeny of these spirochaetes in previous studies; and finally the glpQ gene. A similar approach, but using slightly different gene targets has been recently applied to assign a Borrelia species isolated from a dog in Florida to the B. turicatae species [16]. In this study, the 16S rRNA gene, flaB, gyrB, and glpQ were assessed to determine phylogenetic relationships. In summary, the outer membrane protein, p66, revealed a 10 nucleotide difference between B. recurrentis and B. duttonii. A similar level of divergence was also found

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between either of these spirochaetes and B. crocidurae, which would suggest that these spirochaetes may indeed represent different species. Using the rrs gene, which is not subject to such selective pressures, differences between these species were slight, with only a four nucleotide difference between B. recurrentis and B. duttonii; and six nucleotides differentiating between B. recurrentis and B. crocidurae; and only two nucleotides between B. duttonii and B. crocidurae. In contrast, the flagellin flaB gene showed only a two adjacent nucleotide difference between these spirochaetes, whereas B. crocidurae appeared to be more distantly related. Comparison of the isolates sequenced within this study again showed heterogeneity with those deposited in GenBank. These findings would support the view that B. recurrentis may indeed be a clone arising from a common ancestral strain or directly from the B. duttonii cluster. This is supported by the closer clustering of B. duttonii strain 406K with isolates of B. recurrentis, rather than the B. duttonii clade. However, the low numbers of isolates, coupled with the limited diversity at the nucleotide level, make it difficult to draw a definite conclusion. Given the similarity displayed among the B. recurrentis and B. duttonii isolates, it was a concern to see the divergence between the two B. crocidurae sequences deposited on GenBank (U28496 and x75204). These appeared to be more distantly related based upon their flagellin sequences. The sequence U28496 gave a two and three nucleotide difference from B. recurrentis and B. duttonii respectively, while the sequence X75204 gave 12 further differences above those observed for X75204. No information was available as to the source or geographical location of these isolates. The two adjacent nucleotide signature observed within this gene could prove of value for diagnostic approaches based on flagellin. This difference could be utilized for restriction endonuclease digest patterns or hybridization probe-based differentiation of these spirochaetes. Of concern is the possession of a B. crocidurae-like signature at this position for B. duttonii strain 406K. Although sequences indicative of B. crocidurae have been reported from areas endemic for B. duttonii, further investigation is needed before a general conclusion can be drawn [7]. Similarly, analysis of the glpQ gene revealed just a two base pair difference between B. recurrentis and B. duttonii. An equivalent divergence was found when the sequences generated within this study were compared with others deposited with GenBank. In an earlier study based on IGS sequence comparisons, greater diversity was found among B. duttonii when compared with B. recurrentis leading to speculation that this may be the ancestral species [7]. Many of these variants were, however, not represented among cultivable strains and were not available for investigation in this study. The existence of greater diversity among B. duttonii was not evident from the analyses presented in this study, which was reliant on cultivable organisms to produce sufficient biomas for this work. The limitations of this study must also be considered when attempting to draw conclusions. The quality of sequences deposited may be variable and thus lead to small differences among strains. Furthermore, the identity of these spirochaetes has been largely concluded based on sequence homology and epidemiological knowledge. This has recently been questioned as the characterization of B. crocidurae-like sequences present in spirochaetes derived from both patients and ticks in East Africa, an area endemic for B. duttonii, which has not previously been described as also having B. crocidurae [7]. Another potential cause of further variation is the use of different isolates for the depositions made to GenBank. Although the strains used in this study were largely the same, those used by others were different, sometimes multiply

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passaged either in vitro or in vivo. In contrast, the isolates we report were only from patients within a limited local area and thus may not truly represent the diversity among these spirochaetes. Given these limitations coupled with the linkage disequilibrium likely to exist between the different markers assessed, it is difficult to categorically conclude whether or not these are distinct species. Certainly, the rrs and p66 genes would argue against these being the same spirochaete, however, results from glpQ, IGS, and flaB tend to support the idea of B. recurrentis being a louse-adapted clone of either B. duttonii or a common ancestor. Again, the narrow ecological niche of B. recurrentis suggests that this spirochaete has recently evolved. Certainly these spirochaetes, unlike other relapsing fever spirochaetes, have no alternative animal reservoirs identified to date. Consequently, they have been refractory to in vivo investigation, requiring primate models. This is in stark contrast to other relapsing fever spirochaetes that have natural reservoirs in other mammalian species, typically small rodents. Whether reported differences in clinical presentation between B. recurrentis and B. duttonii are associated with their different vector transmission, host immunological susceptibility or clonally associated virulence factors remains to be determined. It is likely that this dilemma will not be fully resolved until full genomic sequencing has been completed.

References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14]

Cutler, S., Moss, J., Fukunaga, M., Wright, D., Fekade, D.Warrell, D., 1997. Borrelia recurrentis characterization and comparison with relapsing fever, Lyme-associated, and other Borrelia spp. Int. J. Syst. Bacteriol., 47: 958í968. Porcella, S.F., Raffel, S.J., Schrumpf, M.E., Schriefer, M.E., Dennis, D.T.Schwan, T.G., 2000. Serodiagnosis of louse-borne relapsing fever with glycerophosphodiester phosphodiesterase (GlpQ) from Borrelia recurrentis. J Clin Microbiol, 38: 3561í3571. Brahim, H., Perrier-Gros-Clau..., J., Postic, D., Baranton, G.Jambou, R., 2005. Identifying relapsing fever Borrelia, Senegal. Emerg Infect Dis, 11: 474í475. Cutler, S.J., Akintunde, C.O., Moss, J., Fukunaga, M., Kurtenbach, K., Talbert, A., Zhang, H., Wright, D.J.Warrell, D.A., 1999. Successful in vitro cultivation of Borrelia duttonii and its comparison with Borrelia recurrentis. Int J Syst Bacteriol, 49 Pt 4: 1793í1799. Penningon, P., Cadavid, D., Bunikis, J., Norris, S.Barbour, A., 1999. Extensive interplasmidic duplications change the virulence phenotype of the relapsing fever agent Borrelia turicatae. Mol Microbiol, 34: 1120í1132. Bunikis, J., Tsao, J., Garpmo, U., Berglund, J., Fish, D.Barbour, A., 2004. Typing of Borrelia relapsing fever group strains. Emerg Infect Dis, 10: 1661í1664. Scott, J.C., Wright, D.J.M.Cutler, S.J., 2005. Typing African relapsing fever spirochetes. Emerging Infectious Diseases, 11: 1722í1729. Cutler, S.J., Fekade, D., Hussein, K., Knox, K.A., Melka, A., Cann, K., Emilianus, A.R., Warrell, D.A.Wright, D.J., 1994. Successful in-vitro cultivation of Borrelia recurrentis. Lancet, 343: 242. Cutler, S.J., Moss, J., Fukunaga, M., Wright, D.J., Fekade, D.Warrell, D., 1997. Borrelia recurrentis characterization and comparison with relapsing-fever, Lyme-associated, and other Borrelia spp. Int J Syst Bacteriol, 47: 958í968. Barbour, A., 1984. Isolation and cultivation of Lyme disease spirochetes. Yale J Biol Med, 57: 521í525. Bunikis, J., Garpmo, U., Tsao, J., Berglund, J., Fish, D.Barbour, A.G., 2004. Sequence typing reveals extensive strain diversity of the Lyme borreliosis agents Borrelia burgdorferi in North America and Borrelia afzelii in Europe. Microbiology, 150: 1741í1755. Ras, N., Lascola, B., Postic, D., Cutler, S., Rodhain, F., Baranton, G.Raoult, D., 1996. Phylogenesis of relapsing fever Borrelia spp. Int. J. Syst. Bacteriol., 46: 859í865. Fukunaga, M., Ushijima, Y., Aoki, L.Talbert, A., 2001. Detection of Borrelia duttonii, a tick-borne relapsing fever agent in central Tanzania, within ticks by flagellin gene-based nested polymerase chain reaction. Vector Borne Zoonotic Dis, 1: 331í338. Brahim, H., 2005. Identifying relapsing fever Borrelia, Senegal. Emerg Infect Dis, 11: 474í475.

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[15] Bunikis, J., Luke, C.J., Bunikiene, E., Bergstrom, S.Barbour, A.G., 1998. A surface-exposed region of a novel outer membrane protein (P66) of Borrelia spp. is variable in size and sequence. J. Bacteriol., 180: 1618í1623. [16] Schwan, T.G., Raffel, S.J., Schrumpf, M.E., Policastro, P.F., Rawlings, J.A., Lane, R.S., Breitschwerdt, E.B., Porcella, S.F., 2005. Phylogenetic analysis of the spirochetes Borrelia parkeri and Borrelia turicatae and the potential for tick-borne relapsing fever in Florida. J. Clin. Microbiol., 43: 3851í3859.

Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Genotyping of Borrelia burgdorferi sensu lato in Russia Edward I. KORENBERG 1, Valentina V. NEFEDOVA, Irina A. FADEEVA, and Nataliya B. GORELOVA Gamaleya Research Institute of Epidemiology and Microbiology, Moscow Abstract. The Borrelia culture collection at the Vector Laboratory of the Gamaleya Research Institute of Epidemiology and Microbiology, Moscow dates from 1983. Today, this collection consists of almost 1,200 primary isolates from different sources from various geographic regions. A PCRRFLP analysis of isolates from Russia and neighboring countries has shown that they include primarily genospecies of B. garinii and B. afzelii, which are widespread and epidemiologically significant, along with B. burgdorferi sensu stricto, B. valaisiana, B. lusitaniae, and B. spielmanii (group A14S). Considerable genetic heterogeneity of B. afzelii has been revealed by comparing the sequences of the rrf (5S)-rrl (23S) ribosomal intergenic spacer in 139 primary isolates obtained from ixodid ticks and from several species of small mammals. All these isolates were previously identified as B. afzelii by PCRRFLP analysis of amplicons of the 5S23S rRNA spacer region. Two genomic subgroups have now been identified. The majority of the isolates have a high degree of homology to the most widespread genomic subgroup of B. afzelii, strain VS461 (98.899.6%). The remainder have a high degree of homology to the other genomic subgroup B. afzelii, strain NT28 (98.499.6%). Several genovariants have been identified within each subgroup with the degree of nucleotide homology within most of them reaching 100%. Seven of ten genovariants belong to subgroup VS461, three genovariants belong to subgroup NT28. Seven allelovariants have been detected by comparing the sequences of a p66 gene fragment (246377 bp) in 45 B. afzelii isolates. Three of seven B. afzelii VS461 genovariants are apparently common in Eurasian natural foci among different species of vectors and reservoir hosts, with one genovariant probably circulating mainly in foci where I. persulcatus is the main vector of Borrelia infection. The remaining four genovariants of this subgroup appear to be associated with I. ricinus ticks and circulate mainly in Europe. The distribution of the three genovariants belonging to B. afzelii NT28 subgroup needs further study. Nucleotide sequences of approximately 100 isolates of different Borrelia genospecies have been deposited in GenBank. Keywords. Borrelia, culture collection, primary isolates, genospecies, genomic subgroup, genovariants

________________________________________ 1 Corresponding author. Mailing address: Edward Korenberg, Gamaleya Research Institute for Epidemiology and Microbiology, Russian Academy of Medical Sciences, Moscow, Russia; E-mail: [email protected]

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Introduction Diseases caused by Borrelia burgdorferi sensu lato were first identified in Russia by serological methods in 1985 [1]. Within a short period of time, it was found that natural foci of Borrelia infections are distributed throughout the forest zone of Russia, from the Baltic region in the west to southern Sakhalin in the east. A major or even the greater part of the world range of Borrelia associated with ixodid ticks is therefore located in Russia [24], and a considerable diversity of these bacteria could be expected in such a vast and ecologically heterogeneous area. Indeed, subsequent studies have made a large contribution to the collection of Borrelia cultures at the Vector Infections Laboratory of the Gamaleya Institute of Epidemiology and Microbiology, Russian Academy of Medical Sciences (Moscow). This culture collection dates from 1983 in connection with research on relapsing fever in Central Asia [5]. The first Russian isolates of B. burgdorferi sensu lato (19871989) were identified using monoclonal antibodies [6, 7]. By 19951996, more than 400 primary isolates were obtained from different regions of Russia and neighboring countries. Their analysis by means of PCR–RFLP showed that they included several genospecies of B. garinii (subgroups 20047 and NT29), B. afzelii, B. lusitaniae, B. valaisiana, and B. burgdorferi sensu stricto [810]. It was also demonstrated that B. garinii and B. afzelii are especially widespread and have greater epidemiological significance in Russia, and the main vectors and reservoir hosts of these pathogens were identified [8, 1115]. These data not only confirmed our concept that different genospecies or other categories of Borrelia may simultaneously circulate in the same biocenosis, which was formulated in 1993 [16] and now appears self-evident, but also provided evidence for the widespread occurrence of vectors and reservoir hosts with mixed infections in natural foci [8]. Moreover, they indicated that the terms “Lyme disease” and “Lyme borreliosis” actually refer to a group of etiologically different diseases. For this reason, we proposed to name them ixodid tick-borne borrelioses (ITBB) in order to differentiate these diseases from argasid tick-borne borrelioses (ATBB) transmitted by soft ticks [4, 17]. In recent years, Borrelia isolates from northwestern [1824] and central [2528] regions of European Russia, western Siberia [2933], eastern Siberia [3436], and the Far East [3739] have been studied by means of PCR, PCR–RFLP, and, in some cases, sequencing of some genomic regions. The results of these studies unequivocally confirm the above data on the composition of Borrelia genospecies in Russia and the general pattern of their circulation in natural foci.

1. A Culture Collection of Borrelia as a Basis for the Study of These Pathogens The collection of borrelial cultures at the Gamaleva Institute currently includes almost 1,200 isolates attributed to eight B. burgdorferi sensu lato genospecies and five forms of relapsing fever Borrelia which now have binary Latin names (Table 1). Most of these isolates (97%) were obtained by our laboratory from different sources. These include three ixodid tick species, one argasid tick species, eight species of small mammals, and human patients. The latter are from 12 administrative regions of Russia (from the Kaliningrad Region in the west to South Sakhalin Region in the east) and eight neighboring countries.

176

Source I. persulcatus

B. afzelii

County

B. garinii

S461

NT28

Sp.

20047

NT29

Russia

54

6

136

136

190

Estonia

1

1

2

1

Kyrgyzstan Total isolates from I. persulcatus I. ricinus

Bbu ss

55

7

138

137

Russia

21

8

26

Belarus

2

3

6

Lithuania

1

Moldova

4

Ukraine

3

Czech Republic

1

Total isolates from I. ricinus I. pavlovskyi

Kazakhstan

I. trianuliceps

Russia

Total isolated from ixodid ticks

32

Blu

Bva

Bsp (Gr. A14S)

Bbi

Bja

Relapsing fever Borrelia

Not studied

Total

52

111

685

5

13

Mixed

2

1

1

2

193

55

116

701

2

3

73

1

2

8

2

3

1

12 1

1

12

1

2

4

14

2 1

5

48

1

16

35

144

205

228

1 1

1

2

3

6

2 1

1

13 4

4 11

1

2

5

7

132

7

1

65

67

124

902

4 5 92

19

29

4

3

6

11

1

2

E.I. Korenberg et al. / Genotyping of Borrelia burgdorferi sensu lato in Russia

Table 1. Borrelia isolates stored at the Vector Laboratory of the Gamaleya Institute of Epidemiology and Microbiology, Moscow.

Table 1. Borrelia isolates stored at the Vector Laboratory of the Gamaleya Institute of Epidemiology and Microbiology, Moscow. B. afzelii

County S461

Clethrionomys glareolus

Russia

NT28

11

Clethrionomys rufocanus

1

Clethrionomys rutilus

4

Microtus agrestis

1

Microtus oeconomus

6

Apodemus uralensis

2

B. garinii Sp.

20047

NT29

25

31

61

1

2

4

3

5

9

Total from small animals

1

5

2 26

Blu

Bva

Bsp (Gr. A14S)

Bbi

Bja

Mixed

17

Relapsing fever Borrelia

Not studied

Total

2

147

8

2

1

9

Sicista betulina Sorex araneus

Bbu ss

2

34

24 3

1

15

7

2

37

1

1

1

2

1

2

2

1

1

8

43

96

28

6

235

4 4

E.I. Korenberg et al. / Genotyping of Borrelia burgdorferi sensu lato in Russia

Source

177

178

Table 1. Borrelia isolates stored at the Vector Laboratory of the Gamaleya Institute of Epidemiology and Microbiology, Moscow. B. afzelii

County S461

Human patients

NT28

Blu

Bva

NT29

Bsp (Gr. A14S)

Bbi

Bja

Mixed 1

Not studied

Total

3

18 1

1

14

1

Uzbekistan

Total isolates in collection

Relapsing fever Borrelia

1 3 5

Received from other laboratories

1

20047

Bbu ss

14

Total from human patients

2

Sp.

Russia Lithuania

Onithodorus papillipes

B. garinii

118

21

8

2

1

7

187

250

339

10

1 6

11

1

2

1

96

19 5

4

8

31

9

141

1192

Abbreviations: Bbu ss, B. burgdorferi sensu stricto; Blu, B. lusitaniae; Bva, B. valaisiana; Bsp, B. spielmanii; Bbi, B. bissetti; Bja, B. japonica. B. recurrentis, B. persica, B. hermsii, B. turicatae, and B. parkerii.

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Source

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179

As early as 1997, the results of analysis of numerous isolates attracted our attention to differences in the distribution of two B. garinii subgroups (types), 20047 and NT29. The former has proved to be widespread in Eurasian natural foci where both Ixodes persulcatus and I. ricinus are the main vectors, with the latter subgroup (NT29

2. Genovariants and Allelovariants of Borrelia afzelii Borrelia afzelii is one of the three main Borrelia genospecies of epidemiological significance that circulate in the greater part of the Eurasian forest zone. Two genomic subgroups are known, B. afzelii VS461 [61] and NT28 [81]. Based on the sequencing ecosystems, and has a broad spectrum of reservoir hosts and vectors which indicates that this genospecies may be characterized by considerable genetic heterogeneity. However, it, like other ITBB pathogens, has not been sufficiently studied in this respect,. To analyze B. afzelii for genetic heterogeneity, we used 139 primary isolates from our collection They were obtained between 1987 and 2002 from I. persulcatus, I. ricinus, and I. trianguliceps ticks in different phases of development and from several small mammal species (reservoir hosts) captured in natural foci located in Russia, the Czech Republic, Lithuania, Estonia, Belarus, Ukraine, and Moldova (Table 2).

Table 2. Isolates used for analyzing genetic diversity of Borrelia afzelii. GenBank (accession no.) No. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21

Isolate Ip-3 Ip-21 Ir-148 Ir-171 Ip-269 Ir-335 Ir-359 Ip-374 Ip-384 Ir-469 Ir-474 Ip-550 Ip-589 Ip-637 Ip-797 Ip-847 Ip-867 Ip-901 Ipn-1023 Ip-1208 Ip-1272

Source Ixodes persulcatus I. persulcatus I. ricinus I. ricinus I. persulcatus I. ricinus I. ricinus I. persulcatus I. persulcatus I. ricinus I. ricinus I. persulcatus I. persulcatus I. persulcatus I.persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus nymph I. persulcatus I. persulcatus

Year 1987 1987 1988 1988 1989 1989 1989 1990 1990 1990 1990 1991 1991 1992 1993 1993 1993 1993 1993 1994 1994

Source location Russia, Leningrad region Russia, Leningrad region Lithuania Czech Republic Russia, Leningrad region Ukraine Moldova Estonia Estonia Russia, Leningrad region Russia, Leningrad region Russia, Kurgan region Russia, Krasnoyarsk region Russia, Leningrad region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region

5S-23S

p66

DQ020302 AY573192 DQ020297

DQ145764 DQ145770

DQ020311 AY772195 DQ020315 DQ020310

DQ020292 DQ020296 DQ020287

AY772044

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Table 2. Isolates used for analyzing genetic diversity of Borrelia afzelii. GenBank (accession no.) No.

Isolate

Source

Year

Source location 5S-23S

22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67

Ip-1286 Ipn-1377 Ipn-1382 Ip-1511 Ip-1567 Ip-1579 Ip-1599 Ip-1644 Ip-1705 Ip-1717 Ip-1779 Ip-1823 Ip-1828 Ip-1852 Ip-1882 Ip-1885 Ip-1901 Ip-1973 Ip-1997 Ir-2196 Ir-2204 Ir-2215 Ir-2221 Ir-2243 Ip-2250 Ip-2324 Ipn-2408 Ip-2499 Ip-2503 Ipn-2637 Ipn-2641 Ir-2771 Ir-2792 Ir-2829 Ir-2837 Ir-2842 Ir-2858 Ip-2922 Ip-2994 Ipn-3195 Ipl-3305 Ir-3452 Ir-3460 Ir-3463 Ir-3474 Ir-3478

I. persulcatus I. persulcatus nymph I. persulcatus nymph I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. persulcatus I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. persulcatus I. persulcatus I. persulcatus nymph I. persulcatus I. persulcatus I. persulcatus nymph I. persulcatus nymph I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. persulcatus I. persulcatus I. persulcatus nymph I. persulcatus larva I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus

1994 1994 1994 1995 1995 1995 1995 1995 1995 1995 1995 1995 1995 1995 1995 1995 1995 1995 1995 1996 1996 1996 1996 1996 1996 1996 1996 1996 1996 1996 1996 1996 1996 1997 1997 1997 1997 1997 1997 1997 1997 1997 1997 1997 1997 1997

Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Tyumen region Russia, Tyumen region Russia, Tyumen region Russia, Tyumen region Russia, Tyumen region Russia, Tyumen region Russia, Perm region Russia, Primorye Russia, Primorye Russia, Perm region Russia, Khabarovsk region Russia, Khabarovsk region Russia, Khabarovsk region Russia, Khabarovsk region Russia, southern Sakhalin Belarus Belarus Belarus Belarus Belarus Russia, Perm region Russia, Perm region Russia, Perm region Russia, Primorye Russia, Primorye Russia, Perm region Russia, Perm region Russia, Stavropol region Russia, Stavropol region Moldova Moldova Moldova Moldova Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region

p66

AY772052 AY772045 DQ145763 DQ020283 DQ020288

DQ020294

DQ145766

DQ020293

DQ145773

DQ020295 DQ020298

DQ145769

DQ020306 DQ060663 DQ066670 DQ020289 AY772053 DQ066666 DQ020308 DQ020303 DQ145772 DQ145768 AY772046

AY772048 DQ020316 DQ020286

DQ066667

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Table 2. Isolates used for analyzing genetic diversity of Borrelia afzelii. GenBank (accession no.) No.

Isolate

68 69 70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 100 101 102 103 104 105 106 107 108 109 110 111 112 113

Ir-3503 Ir-3507 Ir-3521 Ir-3527 Ir-3528 Ipn-4005 Ipl-4063 Ipl-4089 It-4139 It-4143 It-4144 Ir-4220 Irn-4263 Ir-4358 Ir-4408 Ir-4420 Ir-4422 Ir-4428 Ir-4429 Ir-4435 Ipn-4451 Ipl-4663 Ipn-4669 Ipn-4701 Ir-4731 Ir-4794 Ir-4801 Ipl-4846 Ipl-4849 Ipn-4859 Ipn-4882 Ipl-4885 Ipn-4889 Ipn-4895 Ipn-4987 Ir-5134 Ir-5160 Ir-5214 Ir-5215 Ir-5357 Ipn-5751 Ipl-5773 Itn-5777 Itn-5778 Cg-219 Ma-266

Source

Year

Source location 5S-23S

I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. persulcatus nymph I. persulcatus larva I. persulcatus larva I. trianguliceps I. trianguliceps I. trianguliceps I. ricinus I. ricinus nymph I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. persulcatus nymph I. persulcatus larva I. persulcatus nymph I. persulcatus nymph I. ricinus I. ricinus I. ricinus I. persulcatus larva I. persulcatus larva I. persulcatus nymph I. persulcatus nymph I. persulcatus larva I. persulcatus nymph I. persulcatus nymph I. persulcatus nymph I. ricinus I. ricinus I. ricinus I. ricinus I. ricinus I. persulcatus nymph I. persulcatus larva I. trianguliceps nymph I. trianguliceps nymph Cl. glareolus M. agrestis

1997 1997 1997 1997 1997 1998 1998 1998 1998 1998 1998 1999 1999 1999 1999 1999 1999 1999 1999 1999 1999 1999 1999 1999 2000 2000 2000 2000 2000 2000 2000 2000 2000 2000 2000 2001 2001 2001 2001 2001 2002 2002 2002 2002 1993 1994

Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Kaliningrad region Russia, Kaliningrad region Ukraine Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Stavropol region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Krasnodar region Russia, Krasnodar region Russia, Krasnodar region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Kaliningrad region Russia, Kaliningrad region Ukraine, Crimea Ukraine, Crimea Russia, Krasnodar region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region

p66

DQ020291 DQ020304

DQ020284 DQ066671 AY772049 AY772197 AY772198 AY772199 DQ020314 AY772196

DQ066664

DQ145771

DQ145767

DQ020305 DQ020301 DQ020290

DQ145774

AY772050 AY772054

DQ066662

DQ020309 DQ020285 DQ020307 AY772055 AY772051 AY772200 AY772201 AY772039 AY772056

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Table 2. Isolates used for analyzing genetic diversity of Borrelia afzelii. GenBank (accession no.) No.

Isolate

Source

Year

Source location 5S-23S

114 115 116 117 118 119 120 121 122 123 124 125 126 127 128 129 130 131 132 133 134 135 136 137 138 139

Mo-285 Crt-292 Cg-676 Cg-924 Cg-1260 Mo-1279 Au-1300 Au-1302 Cg-1304 Mo-2013 Mo-2037 Cg-2078 Mo-2115 Crt-2299 Cg-2350 Cg-2428 Crt-2431 Crf-2433 Mo-2487 Cg-2729 Cg-2783 Cg-2870 Mo-2953 Sa-3061 Sa-3078 Crt-3151

M. oeconomus Cl. Rutilus Cl. glareolus Cl. glareolus Cl. glareolus M. oeconomus A. uralensis A. uralensis Cl. glareolus M. oeconomus M. oeconomus Cl. glareolus M. oeconomus Cl. Rutilus Cl. glareolus Cl. glareolus Cl. Rutilus Cl. rufoɫanus M. oeconomus Cl. glareolus Cl. glareolus Cl. glareolus M. oeconomus S. araneus S. araneus Cl. Rutilus

1994 1994 1995 1995 1997 1997 1997 1997 1997 1998 1998 1998 1998 1998 1998 1998 1998 1998 1998 2000 2000 2000 1998 2001 2001 2002

Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region Russia, Perm region

p66

AY772058 AY772042

AY772038

DQ066665

AY772057 AY772040

AY772041

DQ066672

DQ066668 AY772047

AY772043

All these isolates were previously identified as B. afzelii by a PCRRFLP analysis of a fragment amplified from the 5S23S rRNA intergenic spacer [8, 15]. Heterogeneity of B. afzelii was inferred from the results of amplification and sequencing of the rrf (5S)-rrl (23S) variable spacer region (245247 bp) and a fragment of the ɪ66 gene (246337 bp). The methods and experimental procedure have been previously described in detail [45]. The dendrogram based on the results of sequencing (Figure 1) was constructed with the CLUSTAL X program package using information on 25 isolates/amplicons from Central Europe, the temperate zone of Russia, Turkey, China, and Korea stored in the EMBL/GenBank/DDBJ databases (the bootstrap indices for tree nodes in all dendrograms presented in this paper were determined with 1000 iterations). A comparative analysis of nucleotide sequences of the spacer region has shown that the isolates belong to two previously described genomic subgroups [61, 81]. A major part of the isolates has a high degree of homology to the most widespread genomic subgroup B. afzelii VS461 (98.899.6%), and the other part, to the genomic subgroup B. afzelii NT28 (98.499.6%). Ten genovariants have been identified in these

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subgroups, with the degree of nucleotide homology within most variants reaching 100%. Seven of ten genovariants belong to subgroup VS461. The first variant (Figure 1, No. 1) comprises 43 of 139 isolates. They have been obtained from I. persulcatus ticks (larvae, nymphs, and adults), the main vectors of Borrelia, I. trianguliceps ticks, which are accessory vectors, and several species of reservoir hosts, including voles of the genera Clethrionomys and Microtus (two species each), pygmy wood mice, and common shrews. All isolates of this genovariant originate from the part of the B. afzelii range where only I. persulcatus, one of the two main Borrelia vectors in Eurasia, is widespread, whereas the European species, I. ricinus, is absent. The pattern of their geographic distribution (Perm and Tyumen regions and Primorye) suggests that the corresponding genovariant of B. afzelii subgroup VS461 may occur in natural ITBB foci within the area extending at least from Eastern Europe to the Far East. The second variant of genomic subgroup VS461 (Figure 1, No. 2) is represented by 38 isolates from different regions of Eurasia, from southern Moravia in the west to southern Sakhalin in the east. Unlike the first variant, their sources included both main vectors, I. persulcatus and I. ricinus, along with I. trianguliceps ticks (parasites of small mammals) and three small mammal species that are common reservoir hosts of spirochetes. The nucleotide sequence of amplicon D-3 (AF497982) from the Czech Republic belongs to this genovariant of subgroup VS461, which appears to be most widespread and characteristic of the majority of ITBB foci in which B. afzelii circulates. The third genovariant (Figure 1, No. 3) is represented by only three isolates from I. ricinus ticks collected in the Kaliningrad and Leningrad regions (Russia) and Belarus. According to information from the EMBL/GenBank/DDBJ databases, the analogous intergenic spacer sequences have been found in isolates from the regions located much farther east, both in Europe (AB178342 and AB178343) and Asia (AB013913 and AB03914). In particular, such isolates were obtained in the Moscow region from I. ricinus and I. persulcatus ticks and in Korea from I. granulatus and I. nipponensis ticks. This is evidence that the corresponding genovariant is also widespread within the B. afzelii VS461 range, although occurs in natural foci less frequently than the previous genovariant. Isolates belonging to the fourth genovariant of this subgroup (Figure 1, No. 4) have been obtained so far only from I. ricinus ticks collected in southwestern Russia (the Stavropol and Krasnodar regions). The EMBL/GenBank/DDBJ database contains no data on the analogous intergenic spacer sequences (with the degree of homology exceeding 99.6%); likewise, we have not found such sequences in our material from other regions. It appears that this genovariant is confined to the southern part of the VS461 subgroup range. All 15 isolates of the fifth genovariant (Figure 1, No. 5) are from I. ricinus ticks collected in the western part of the pathogen range. The GenBank contains information on three isolates with analogous spacer sequences obtained from the same source in central Russia and Turkey. In addition, we included isolate Ir-335 in this genovariant, as it proved to have 100% homology to the GenBank sequence of an isolate obtained in the Moscow region from an I. ricinus tick. The sixth genovariant of subgroup VS461 (Figure 1, No. 6) is represented by only 1 of the 139 isolates, designated Ip-21. This well-known isolate, obtained from an I. persulcatus tick in the Leningrad region, was used to make the first description of B. afzelii as a genospecies [47]. In Russia, it has been employed for many years as the

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producer of the standard corpuscular antigen for indirect immunofluorescent tests. A comparative analysis of its spacer sequence and the sequences deposited in the GenBank provides evidence that this genovariant is widespread in both western and eastern parts of the B. afzelii range and has a broad range of vectors and reservoir hosts. No less extensive geographic distribution and range of hosts are characteristic of the seventh genovariant of B. afzelii subgroup VS461. In the dendrogram based on the results of sequencing, isolates belonging to this genovariant form a distinct and separate cluster (Figure 1, No. 7). Borrelia afzelii of genomic subgroup NT28 are represented by isolates from the Kaliningrad, Leningrad, Perm, Krasnodar, and Stavropol regions (Russia), Estonia, Moldova, and Belarus. In general, spirochetes of this subgroup are apparently widespread in many natural ITBB foci. Isolates from I. ricinus ticks form two distinct separate clusters in the dendrogram based on the results of sequencing (Figure 1, Nos. 8 and 9). In view of available data, these genovariants appear to be more “western.” The degree of their homology to other representatives of B. afzelii NT28 subgroup reaches 98.899.6%, compared to 97.198.0% in the case of comparison with B. afzelii VS461. However, isolates of one variant (Figure 1, No. 9) are highly homologous (up to 98.8%) to isolate 934U obtained in Korea [81], whose RFLP profile is similar to that of B. afzelii VS461. Hence, it may well be that subsequent research will show that this variant has a wider, Eurasian distribution. The third genovariant of B. afzelii NT28 subgroup is comprised of isolates from both “persulcatus” and “ricinus” types of natural ITBB foci (Figure 1, No. 10). With respect to the test sequence, their homology to the typical representatives of B. afzelii NT28 ranged from 98.4 to 99.2%, compared to 96.797.6% in the case of subgroup VS461. Recent data on the sequencing of conserved genomic regions have shown that some isolates previously classified as B. afzelii differ markedly from typical representatives of this genospecies in the corresponding sequences and, according to some authors [74, 78, 83], may be regarded as new, independent genospecies. Taking this into account, we performed repeated (control) identification of all the above genovariants by means of PCRRFLP analysis. One isolate from each genovariant was used for this purpose. In all cases, cleavage by MseI restriction enzyme yielded fragments 108, 68, 50, and 20 bp in size, which was specific for the restriction profile of B. afzelii [45]. An analysis of nucleotide sequences of the variable spacer region has revealed nucleotide substitutions characteristic of B. afzelii genovariants described above. Figure 2 shows the sequences of the isolates representing these genovariants (in the same order as in Figure 1) compared to the corresponding sequences of Borrelia subgroups VS461 and NT28. Identical substitutions characteristic of the genomic subgroups are in positions 9498; in addition, individual genovariants have substitutions in positions 56, 60, 76, 102, 161, 172, and 236 (Figure 2). Thus, the results of analysis of the rrf (5S)-rrl (23S) spacer sequences in a representative sample of isolates confirm significant intraspecific heterogeneity of B. afzelii. This genospecies, widespread in Eurasia, has two genotypic subgroups (VS461 and NT28), which comprise no less than ten genovariants. Three of seven genovariants of subgroup VS461 are apparently widespread in Eurasian natural ITBB foci among different species of vectors and reservoir hosts. The degree of sequence homology between isolates of these genovariants and the type B. afzelii VS461 strain is 98.899.6%. It may well be that one of these genovariants circulates mainly in natural

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ITBB foci of the “persulcatus” type. Available data indicate that the remaining four genovariants of this subgroup circulate mainly in Europe and are associated with I. ricinus ticks and, probably, their hosts. The distribution of the three genovariants of B. afzelii subgroup NT28 needs more detailed study. In addition, the genovariants described above were compared with respect to the nucleotide sequence of a 1446-bp fragment of the 16S rRNA gene, which was determined in one isolate per genovariant. In the dendrogram constructed on this basis, two clusters are distinct. One comprises seven rrf (5S)-rrl (23S) genovariants (suggesting their similarity), and the other comprises three such genovariants. However, isolates closely similar in the intergenic spacer sequence proved to fall into different clusters with respect to the 16S rRNA gene sequence (Figure 3). The sequences of a fragment of the chromosomal gene ɪ66 were determined in 45 primary B. afzelii isolates from three ixodid tick species and seven species of small mammals (reservoir hosts of Borrelia) inhabiting natural ITBB foci in Russia, Lithuania, Estonia, Belarus, Ukraine, and Moldova. A comparative analysis of these sequences made it possible to distinguish seven allelovariants of these isolates (Figure 4), with the degree of sequence homology within most variants reaching 99.9100%. Homology between isolates of different variants ranged from 98.9 to 99.7%. The first allelovariant (Figure 4, variant 1) comprises 15 out of 45 isolates obtained in Europe (from Moldova to the Cisural region) from the main Borrelia vectors, I. persulcatus and I. ricinus ticks (larvae, nymphs, and adults); accessory vectors, I. trianguliceps ticks; and reservoir hosts, bank voles (Clethrionomys glareolus). These isolates are 100% homologous to the nucleotide sequences of West European isolate ACAI deposited in the EMBL/GenBank/DDBJ databases (accession no. X87726). Their geographic distribution indicates that this allelovariant of B. afzelii occurs, at the lowest estimate, in natural ITBB foci located in Europe. Seven isolates belonging to the second allelovariant (Figure 4, variant 2) were obtained in different regions of Eurasia (from Estonia to southern Sakhalin) from nymphs and adult individuals of the main Borrelia vector, I. persulcatus. It may well be that spirochetes of this allelovariant will subsequently be found in other vectors and reservoir hosts. Apparently, this variant has a wide geographic distribution and occurs in many natural ITBB foci in which B. afzelii circulates. The third allelovariant (Figure4, variant 3) is represented in our material by only two isolates obtained from I. ricinus ticks in different regions of Europe, i.e., in Moldova and the Cisural region (Russia). The small sample size and the absence of information about isolates with analogous p66 gene sequences in the EMBL/GenBank/DDBJ databases do not allow any definite conclusions concerning the distribution of this allelovariant, but it probably occurs in natural foci less frequently than the previous variants. Isolates of the fourth allelovariant (Figure 4, variant 4) are from different vectors and reservoir hosts collected in natural ITBB foci in the European part of B. afzelii range, and the possibility of circulation of analogous spirochetes in Asian foci needs further verification. The fifth allelovariant is represented by only one isolate (Ip3) obtained from an I. persulcatus tick in the Leningrad region, which has the lowest degree of homology with the other variants (98.9%). The sixth variant is also represented by only one isolate (Ir-4429) from an I. ricinus tick collected in the Stavropol region. A wide distribution among different hosts in Eurasia is characteristic of the seventh allelovariant, with the corresponding isolates joining into a distinct cluster on the dendrogram (Figure 4, variant 7). Comparing the

186 E.I. Korenberg et al. / Genotyping of Borrelia burgdorferi sensu lato in Russia

Figure 1. Simplified dendrogram constructed on the basis of similarity in the rrf (5S)-rrl (23S) spacer sequence between 139 B. afzelii isolates [45].

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ɪ66 gene sequences in isolates belonging to different allelovariants of B. afzelii, we have revealed characteristic nucleotide substitutions in positions 59, 94, 102, 246, 259, and 278 (Figure 5). Thus, the results of our study show that the structure of chromosomal gene p66 differs not only in different species of the group Borrelia burgdorferi sensu lato, as it was shown previously [71, 84, 85], but also within the same genospecies. Thus, B. afzelii has no less than seven corresponding allelovariants. Most of them are widespread in Eurasia among different species of ticks and small mammals inhabiting natural ITBB foci, but no connection between a certain allelovariant and a certain vector or reservoir host can be traced. As already noted, we found ten B. afzelii genovariants differing in the 5S23S rRNA intergenic spacer sequence [45]. Isolates representing nine genovariants were analyzed for variation in the p66 gene sequence. The results showed that most of them included several p66 allelovariants (Table 3), with all of the latter occurring in both known genomic subgroups of B. afzelii, VS461 and NT28. In general, no connection between a vector or reservoir host and a particular genovariant (allelovariant) of the pathogen can be traced: different species of ticks and small mammals may be the hosts of spirochetes belonging to different intraspecific variants (Table 4). However, genovariants 35 appear to tend toward the western part of the pathogen range, where I. ricinus is the main vector. Their geographic distribution indicates that this allelovariant of B. afzelii occurs, at the lowest estimate, in natural ITBB foci located in Europe.

Table 3. Occurrence of B. afzelii allelic variants (by gene ɪ66) amon0067genovariants revealedby sequencing the 5S23S rRNA intergenic spacer region. 5S23S

ɪ66 Number of cluster in Figure 4

Number of cluster in Figure 1 1

2

3

4

5

6

7

1

+

+

-

+

+

-

+

2

+

+

+

-

+

-

+

4

-

-

-

-

-

-

+

5

-

-

-

-

+

-

-

6

+

+

-

-

-

-

-

7

+

-

-

-

-

-

+

8

+

+

-

-

-

+

-

9

-

-

-

-

+

-

-

10

+

-

-

+

+

-

-

188 E.I. Korenberg et al. / Genotyping of Borrelia burgdorferi sensu lato in Russia

Figure 2. Nucleotide sequences of the rrf (5S)–rrl (23S) spacer region in isolates belonging to different genovariants and genomic subgroups NT28 and VS461 of B. afzelii.

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Fiure 3. Dendrogram of ten B. afzelii isolates representing different genovariants (with respect to the rrf (5S)–rrl (23S) spacer sequence) constructed on the basis of similarity in the sequence of a 1446-bp fragment of the 16S rRNA gene.

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Figure 4. Dendrogram of 45 B. afzelii isolates constructed on the basis of similarity in the nucleotide sequence of region 246337 of the p66 gene. Circles show the numbers of allelic variants.

E.I. Korenberg et al. / Genotyping of Borrelia burgdorferi sensu lato in Russia

Figure 5. A nucleotide sequence of region the size 220 bp of the p66 gene in isolates representing different allelic variants of B. afzelii.

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192

Table 4. Occurrence of B. afzelii genovariants and allelic variants in different vectors and reservoir hosts. p66

5S-23S Vectors and reservoir hosts

Number of cluster in Figure 1 1

2

3

4

5

6

7

8

Number of cluster in Figure 2

9

10

1

2

3

4

5

6

7

I. persulcatus Larvae

+

+

-

-

-

-

-

-

-

+

+

-

-

+

-

-

+

Nymphs

+

+

-

-

-

-

+

+

-

-

+

+

-

-

-

-

+

Adults

+

+

-

-

-

+

+

-

-

+

+

+

-

+

+

-

+

Larvae

-

-

-

-

-

-

-

+

-

-

-

-

-

+

-

-

-

Adults

-

+

+

+

+

-

+

+

+

+

+

-

+

+

-

+

-

Nymphs

+

-

-

-

-

-

-

-

-

-

-

-

-

-

-

-

+

Adults

+

+

-

-

-

-

-

-

-

-

+

-

-

+

-

-

-

C. glareolus

+

+

-

-

-

-

-

-

-

-

+

-

-

+

-

-

-

C. rutilus

+

+

-

-

-

-

-

-

-

-

C. rufocanus

-

-

-

-

-

-

-

-

-

+

-

-

-

-

-

-

+

M. oeconomus

+

-

-

-

-

-

+

-

-

+

-

-

-

+

-

-

-

M. agrestis

+

-

-

-

-

-

-

-

-

-

A. uralensis

+

+

-

-

-

-

-

-

-

-

-

-

-

S. arameus

+

-

-

-

-

-

+

-

-

-

I. ricinus

I. tranguliceps

No data

No data -

+

-

-

No data

3. Heterogeneity of B. afzelii in a Natural Focus Among isolates analyzed for the 5S23S rRNA spacer sequences, 72 isolates were obtained in the course of long-term studies in the same natural focus located in eastern Europe (the Perm region); in 22 out of these 72 isolates, the fragment of gene ɪ66 was sequenced. In addition to B. afzelii, B. garinii spirochetes circulate in this focus, with the taiga tick I. persulcatus being the main epidemiologically significant vector of both pathogens. The accessory vector I. trianguliceps is of purely epizootiological significance. The range of reservoir hosts includes several species of small mammals [13, 15]. Both tick species in different phases of development and seven species of small mammals (reservoir hosts) were the sources of the above isolates (Table 5). Analysis of the 5S23S rRNA spacer sequences has shown that B. afzelii of two genomic subgroups circulate in the focus. Most of the isolates (90.3%) have a high degree of sequence homology (98.899.6%) to the most widespread genomic group VS461; in the remaining isolates (9.7%), the degree of homology to subgroup NT28 is higher than to VS461: 98.499.2 vs. 96.797.6%.

E.I. Korenberg et al. / Genotyping of Borrelia burgdorferi sensu lato in Russia

193

Table 5. Occurrence of B. afzelii genovariants and allelovariants in vectors and reservoir hosts from a natural focus in eastern Europe.

Vectors

5S-23S

p66

Number of cluster

Number of cluster

in Figure 1

in Figure 4

and reservoir hosts 1

2

3

4

5

1

2

3

4

5

I. persulcatus Larvae

+

-

+

-

+

+

-

-

+

+

Nymphs

+

+

+

+

-

+

-

+

-

+

Adults

+

+

+

-

+

+

+

-

-

-

Nymphs

+

-

-

-

-

-

-

-

-

+

Adults

+

-

+

-

-

+

-

-

-

-

C. glareolus

+

-

+

-

-

+

-

-

+

-

C. rutilus

+

-

+

-

-

No data

C. rufocanus

-

-

-

-

+

-

-

-

-

+

M. oeconomus

+

+

-

-

+

-

-

-

+

-

M. agrestis

+

-

-

-

-

No data

A. uralensis

+

-

+

-

-

-

+

-

-

S. arameus

+

+

-

-

-

No data

I. tranguliceps

-

In each genomic subgroup, different genovariants of Borrelia are distinguished, with the degree of sequence homology between isolates within most genovariants reaching 100% (see above). On the whole, at least five genovariants differing in one to eight nucleotides circulate in the focus. Three genovariants (Figure 1, variants 13) belong to genomic subgroup VS461, and two genovariants (Figure 1, variants 4 and 5), to subgroup NT28. Genovariants of B. afzelii differing in the 5S23S rRNA spacer sequence were regularly revealed in the focus, provided the number of isolates obtained in a given year was sufficient. All possible hostsdifferent phases of I. persulcatus and I. trianguliceps ticks (the main and accessory vectors), as well as background species of small mammals (reservoir hosts)proved to be infected by spirochetes of several genovariants (Table 5). Among B. afzelii isolates from these natural foci, we distinguished five allelovariants differing in the p66 gene sequence. Most of them were revealed in different hosts every year, and no connection between a certain allelovariant and a tick or small mammal species could be revealed (Table 5). These data show that two genomic subgroups of B. afzelii (VS461 and NT28) can simultaneously circulate in the same focus among different vectors and reservoir hosts.

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194

These subgroups are represented by no less five allelovariants differing in the 5S23S rRNA spacer sequence and no less than five genovariants differing in the p66 gene sequence. None of these variants are confined to a certain host. On the other hand, a tick or small mammal species may be the host of several genovariants or allelovariants of B. afzelii, which contributes to the maintenance of genetic heterogeneity of the pathogen population in the natural focus.

4. Conclusion It has been shown that linked parasitic systems formed by different Borrelia genospecies can exist in the same area [16]. This phenomenon is a major prerequisite for the epidemic manifestation of mixed Borrelia infections [86]. Furthermore, PCRRFLP analysis has allowed us to reveal simultaneous circulation of B. afzelii and two subgroups of B. garinii (20047T and NT29) among their common vectors and reservoir hosts [8, 13, 15]. The sequencing of the 5S23S rRNA spacer and p66 gene region in B. afzelii isolates obtained in the same area in different years has shown that the parasitic system of this genospecies includes five genovariants and five allelovariants that belong to subgroups VS461 and NT28. Apparently, B. garinii is also represented by different genovariants in this linked parasitic system. This aspect, along with genetic heterogeneity of ITBB pathogen populations, will be analyzed in more detail in our subsequent studies. However, the data presented above provide sufficient evidence for considerable genetic heterogeneity of different Borrelia species, which agrees with observations made by other researchers [71, 87, 88]. This heterogeneity is probably accounted for by the clonal variation of Borrelia [9092] and subsequent selection and adaptation of the clones in the course of their circulation in a natural focus. The ecological significance of this phenomenon is evident: heterogeneity of microorganisms provides for the sustainable existence of the parasitic system they form and, in particular, for their survival upon interactions with various hosts and vectors [93]. However, its significance for infection pathology is as yet ambiguous and needs further analysis.

Acknowledgments This study was supported by the Russian Foundation for Basic Research (project ʋ 0404-48066).

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[70] A. Le Fleche, D. Postic, K. Girardet, et al., Characterization of Borrelia lusitaniae sp. nov. by 16S ribosomal DNA sequence analysis, Int. J. Syst. Bacteriol., 47 (1997), 921925. [71] D.E. Norris, B.J.B. Johnson, J. Piesman, et al., Culturing selects for specific genotypes of Borrelia burgdorferi in an enzootic cycle in Colorado, J. Clin. Microbiol, 35 (1997), 23592364. [72] C. Valsangiacomo, T. Balmelli, J.-C. Piffaretti, A phylogenetic analysis of Borrelia burgdorferi sensu lato, based on sequence information from the hbb gene coding for a histone-like protein, Int. J. Syst. Bacteriol., 47 (1997), 110. [73] J. Bunikis, C.J. Luke, E. Bunikis, et al, A surface-exposed region of a novel outer membrane protein (P66) of Borrelia sp. is variable in size and sequence, J. Bacteriol., 180 (1998), 16181623. [74] G. Wang, A.P. van Dam, L. Spanjaard, J. Dankert, Molecular typing of Borrelia burgdorferi sensu lato by randomly amplified polymorphic DNA fingerprinting analysis, J. Clin. Microbiol., 36 (1998), 768776. [75] D. Postic, N. Marti Ras, R.S. Lane, et al, Common ancestry of Borrelia burgdorferi sensu lato strains from America and Europe, J. Clin. Microbiol., 37 (1999), 30103012. [76] H.-S. Park, J.-H. Lee, E.-J. Jeong, et al., Evolution of groEl gene analysis for identification of Borrelia burgdorferi sensu lato, J. Clin. Microbiol., 42 (2004), 12701273. [77] C.-J. Fraenkel, U. Garpmo, and J. Berglund, Determination of novel Borrelia genospecies in Swedish Ixodes ricinus ticks, J. Clin. Microbiol., 40 (2002), 33083312. [78] M. Derdakova, L. Beati, B. Pet’ko. et al., Genetic variability within Borrelia burgdorferi sensu lato genospecies established by PCR-single-strand conformation polymorphism analysis of the rrfArrlB intergenic spacer in Ixodes ricinus ticks from Czech Republic, Appl. Environ. Microbiol., 69 (2003), 509516. [79] O.V. Morozova, N.V. Fomenko, V.A. Rar, et al., Molecular-genetic analysis of tick-borne infections in Novosibirsk region, V.I. Pokrovsky (Ed.), Genetic diagnosis of infectious diseases, vol. 2, Meditsina dlya Vsekh, Moscow (2004), 7981 (in Russian). [80] N.A. Rudnikova, L.S. Karan', G.S. Kislenko, et al., Molecular analysis in studies on the distribution of ITBB pathogens among I. persulcatus and I. ricinus ticks in Moscow region, V.I. Pokrovsky (Ed.), Genetic diagnosis of infectious diseases, vol. 2, Meditsina dlya Vsekh, Moscow (2004), 185185 (in Russian). [81] T. Masuzawa, T. Kamikado, A. Iwaki, et al., Characterization of Borrelia sp. isolated from Ixodes tanuki, I. turdus, and I. columnae in Japan by restriction fragment length polymorphism of rrf (5S)rrl(23S) intergenic spacer amplicons, FEMS Microbiol. Lett., 142 (1996), 7783. [82] J. Bunikis, U. Garpmo, J. Tsao, et al., Sequence typing reveals extensive strain diversity of the Lyme borreliosis agents Borrelia burgdorferi in North America and Borrelia afzelii in Europe, Microbiology, 150 (2004), 17411755. [83] G. Wang, A.P. van Dam, J. Dankert, Phenotypic and genetic characterization of a novel Borrelia burgdorferi sensu lato isolate from a patient with Lyme borreliosis, J. Clin. Microbiol., 37 (1999), 30253028. [84] A.G. Barbour, S.L. Tessier, and S.F. Hayes, Variation in a major surface protein of Lyme disease spirochetes, Infect. Immunity, 45 (1984), 94100. [85] J. Bunikis, L. Noppa, S. Bergstrom, Molecular analysis of a 66-kDa protein associated with the outer membrane of Lyme disease Borrelia, FEMS Microbiol. Lett., 131 (1995), 139145. [86] E.I. Korenberg, Mixed infections transmitted by ticks: Current state of the problem, Usp. Sovrem. Biol., 123 (1996), 475-486 (in Russian). [87] M. Derdakova, B. Dudioak, B. Brei, et al., Interaction and transmission of two Borrelia burgdorferi sensu stricto strains in a tickrodent maintenance system, Appl. Environ. Microbiol., 70 (2004), 67836788. [88] M.C. Dolan, J. Piesman, B.S. Schneider, et al., Comparison of disseminated and nondisseminated strains of Borrelia burgdorferi sensu stricto in mice naturally infected by tick bite, Infect. Immunity, 72 (2004), 52625266. [89] A.F. Elias, J. Schmutzhard, P.E. Stewart. et al., Population dynamics of a heterogeneous Borrelia burgdorferi B31 strain in an experimental mousetick infectious cycle, Wien Klin Wochenschr., 114 (2002), 557561. [90] A.F. Elias, P.E. Stewart, D. Grimm, et al., Clonal polymorphism of Borrelia burgdorferi strain B31 MI: Implications for mutagenesis in an infectious strain background, Infect. Immunity, 70 (2002), 21392150. [91] U. Pal, E. Fikrig, Adaptation of Borrelia burgdorferi in the vector and vertebrate host, Microbes Infect., 5 (2003), 659666.

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Ecological and Genetic Diversity within the Leptospiraceae Family: Implications for Epidemiology Yulia V. ANANYINA 1 , Anna P. SAMSONOVA, Evgeny M. PETROV, Igor A. SHAGINYAN, Marina Yu. CHERNUKHA, Marina S. ZEMSKAYA and Yulia S. ALYAPKINA Gamaleya Research Instititute for Epidemiology and Microbiology, Moscow, Russia Abstract. Spirochaetes within the Leptospiraceae family, encompassing the genera Leptospira, Leptonema and Turneria, are known to display substantial phenotypic and genetic variation resulting in their capacity to colonize diverse natural habitats worldwide. There are many lines of evidence to suggest that these microorganisms belong to at least three ecotypes. The first type consists of freeliving nonparasitic leptospires inhabiting water, including marine and soil environments (Leptospira biflexa sensu lato). Not infrequently, saprophytic leptospires contaminate nutrition media, their occasional alleged isolation from mammalian hosts or humans resulting not only in misdiagnosis, but also in taxonomic confusion. The second ecological type consists ofparasitic leptospires showing pathogenic potential for humans and various animal species as well as host and tissue specificity of a wide spectrum (L. interrogans sensu lato). The third group comprises leptospires with life styles intermediate between independent and symbiotic (also within L. interrogans sensu lato). The latter (for example, L. kirshneri serovar grippotyphosa) display features of so-called sapronotic agents: high survival and competitive potential in environmental ecosystems, widespread mammalian host range and pronounced antigenic and genetic heterogeneity. This paper addresses ecological and genetic diversity within the Leptospiraceae family as relevant to epidemiological and taxonomic issues. Key words: Leptospiracea family, species, ecology, antigenic and genetic diversity, taxonomy

Introduction Leptospirosis is known as a zoonosis affecting both human beings and a variety of wild and domestic animal species, and is therefore widespread over the world [1, 2]. Certain leptospiral pathogens (for example, Leptospira interrogans, L. copenhageni, L. icterohaemorrhagiae and L. canicola serovars) cause severe diseases with high fatality 1 Corresponding Author: Yu. V. Ananyina. Mailing address: 123098, Gamaleya str.18, Gamaleya Institute for Epidemiology and Microbiology,Moscow,Russia; E-mail: [email protected].

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rates or result in late clinical sequelae in affected patients. Meanwhile others, though less virulent (L. kirschneri, L. grippotyphosa serovars, et al.), are able to induce cluster or outbreak disease, mainly water-borne, with morbidity involving dozens and even hundreds of human beings [2, 3]. Spirochaetes of the Leptospiraceae family, encompassing the genera Leptospira, Leptonema and Turneria, display substantial phenotypic and genetic variation resulting in their capacity to colonize diverse natural habitats worldwide. From the phylogenetic perspective, they form the oldest, most deeply branching group among the spirochaetes [4]. In the pre-genomic era, leptospires had been subdivided into two major ecotypes, parasitic and free-living, differing with respect to their mode of existence and specific phenotypic patterns, first and foremost according to pathogenic potential. For decades a pet topic of long-term taxonomic discussions was discussion about whether they comprised one or two separate species or complexes of species [5]. Over time this initial differentiation was confirmed by various methods of phylogenetic analysis, inter alia, based on 16S rRNA and 16S rDNA sequences. Leptospires were shown to split into two monophyletic groups, one of which was formed by parasitic leptospires (L. interrogans sensu lato) and the other by nonparasitic free-living leptospires (L. biflexa sensu lato) [4, 6]. This paper addresses ecological and genetic diversity within the Leptospiraceae family as relevant to some epidemiological and taxonomic issues.

1. Free-living Nonpathogenic Leptospires (Leptospira biflexa sensu lato) Careful consideration has always been placed on ecological and genetic characterization of pathogenic leptospires causing disease in humans and animals of various species. The ecology and molecular genetics of free-living leptospires, lacking epidemiological significance,seems to be an underappreciated area. Meanwhile, the recent breakthrough in leptospira genetics was due to the isolation of leptophages from sewage water found to be specific for saprophytes [7] and the construction of the first Leptospira-Escherichia coli shuttle vector [8], followed by the first demonstration of gene knockout in Leptospira [9]. A system for random transposon mutagensis of L. biflexa, most important for elucidating mechanisms of leptospira physiology, metabolism and pathogenicity, has also been recently developed [10]. Nonetheless, at present not much is known about the lifestyle of these spirochaetes in natural microhabitats, except for data on their high ecological plasticity that permits survival over a broad range of environmental conditions. Saprophytic leptospires were shown to be highly tolerant to fluctuations of environmental factors (ultraviolet radiation, temperature, PH, osmotic pressure, moisture, nutrient deprivation) [5, 11]. They are also characterized by pronounced antigenic [12] and genetic diversity [13] that may reflect heterogeneity of their ecological niches. When co-cultured in vitro with pathogenic leptospires, their fast-growing, free-living “twins” show high competitive potential [14] that may be explained by the ecological mechanism of competitive exclusion. L. biflexa sensu lato are commonly found in diverse environments: moist soil; tap, fresh, and sewage water; and thermal and hydrosulfuric sources. In the mid 70s, the existence of sodium-dependent leptospires selectively adapted to survive in marine water was brought to light [15]. However, halophylic leptospires later fell out the field

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of vision of leptospirologists, although their position in the context of comparative phylogeny, genomics and taxonomy deserves special attention. Not infrequently, saprophytic leptospires contaminate nutritional media both commercial and home-made. Notwithstanding that this fact has been well known for decades [11, 16, 17], occasional alleged isolations of media contaminants from mammalian hosts or humans from time to time cause not only misdiagnosis, but also taxonomic confusion. Over recent decades a number of leptospira strains proposed for inclusion in classification schemes as new species or as higher taxons, e.g., genera, appear to be free-living contaminants of commercial media. Among them are the type strains of two out of three genera comprising the Leptospiraceae family. The first of these is the Leptonema genus with its single species L. illini [18]. The type strain 3055 was allegedly isolated from a bull’s urine in the USA [19], but was later shown to be a saprophyte. The second, strain Phoney parva (HT), representing the Turneria genus with its single species L. parva, was initially known to be isolated in England directly from commercially cultured medium [20]. L. parva has now been recommended to be transferred to the genus Turneriella as Turneriella parva, gen.nov.,comb.nov. [21]. The recent analysis of G+C content, DNA-DNA relatedness, and 16S rRNA sequence of L. turneria indicated that this species was not related to other Leptospira species. Within the Leptospira genus we also find species proven or suspected to be “pseudopathogenic.” Among them is L. inadai (lyme serovar), strain 10, that had been isolated from a skin biopsy of a Lyme-borreliosis patient and was subsequently delineated as belonging to a new pathogenic species, L. inadai [22]. Shortly thereafter, two leptospiral isolates (Enr86, Enr87) were obtained by us from one of the batches of the EMJH semi-synthetic medium enrichment (Difco, batch N744084). Interestingly, the concentration of contaminant-leptospires was as high as 7x105 cells per ml of medium enrichment, and their cells remained viable throughout two years of storage under 4°C. Several lines of phenotypic and genetic evidence testifying to the clonal origin of 10 (lyme), Enr86 and Enr87 strains were obtained. In ɫross-agglutinin absorbion studies, 10 (lyme), Enr86, and Enr87 strains were completely identical. Subcultures of these strains appeared to be nonpathogenic for Syrian golden hamsters weighing 25–30g, causing neither acute disease nor chronic renal carriership in challenged animals. Similar to saprophytes, they propagated at 13°C and in bicarbonate medium with NaHCO3- 1‰ [23]. Genomic fingerprinting of leptospiral DNA with M13phage DNA as a molecular probe also confirmed clonality of 10 (lyme), Enr86 and Enr87 isolates. In contrast to pathogenic leptospires, hypervariable repeats were detected in all representatives of L. biflexa tested, including L. inadai and L. phoney parva. Saprophytic strains showed marked genomic polymorphism, except for the 10 (lyme), Enr-86, and Enr-87 isolates which had identical profiles [13]. In agreement with these findings in the arbitrarily primed PCR, the three strains also were undistinguishable [24]. 1.1. Prevalence of the Gene for LipL32 in Various Leptospiral Taxa As has been recently shown, the gene encoding the major outer membrane protein LipL32 is the most abundant in the Leptospira surfaceome and is also highly conserved across pathogenic species. In contrast to LipL36, major outer membrane protein LipL32 is expressed during infection in the mammalian host [25, 26]. We have screened the prevalence of LipL32 gene in the DNA of 74 referenced leptospira strains and field isolates representing various leptospiral taxa, obtained from different sources

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and geographical locations in the period between 1915 and 2004. Primers (LEP21: 5' GTC TTG TGG TGC TTT CGG TGG - 3'/LEP22: 5' - GCG GGC TCA CAC CTG GAA TAC - 3'), flanking a 677 bp fragment of the LipL32 gene, were constructed using computer program OLIGO4.0. in an Applied Byosystems 381A DNA Synthesizer. The PCR assay conditions have been described elsewhere [27]. As would be expected, the LipL32 gene fragment was detected in the DNA of 59 leptospira strains belonging to pathogenic species (L. interrogans, L. borgpetersenii, L. kirschneri, L. noguchi, L. weilii). Some of these results are shown in Figure 1.

Figure 1. Amplification of a 677 bp fragment of the LipL32 gene by PCR assay using LEP21/ LEP22 primers from genomic DNA of various leptospiral species. Lane 1, O` Range Ruler 100 bp DNA Ladder; lane 2, L.interrogans M-20; lane 3. L. kirschneri Moskva V; lane 4, L. borgpetersenii Poi; lane 5, L. noguchii CZ 214K; lane 6, L. weilii Sarmin; lane 7, L. interrogans Akijami A. Lane 8, O` Range Ruler 100 bp DNA Ladder; lane 9. L. biflexa Patoc 1; lane 10, L. meyeri ICF; lane 11, Turneria parva HT; lane 12, Leptonema illini 3055; lane 13. L. wolbachii ɋDC; lane 14, L. inadai 10.

In all saprophytic strains and also in three L. inadai strains, Leptonema, Turneria, L. sanrarosai (shermani), and L. meyeri ICF–ranarum, the LipL32 gene fragment was not detected (Table 1), thus confirming their nonpathogenic character. Incidentally, there was a recent report on probable laboratory contamination of clinical specimens with L. meyeri [17]. L. fainei (serovar hurstbridge) has been recently described as an emerging pathogen in Australia [28, 29], the Seychelles [30], and Europe [31, 32]. However, the analysis of data published so far shows that the pathogenic status of this novel leptospira genomospecies is still open to question. Thus, the DNA-DNA relatedness or sequence-based classification scheme currently in use is overly biased in favor of nonpathogenic leptospires—media contaminants—with high taxonomic ranks, occupying two genera and quite a number of genospecies. As mentioned elsewhere, populations of free-living leptospires, similar to saprophytes of other taxa, are characterized by extremely high antigenic and genetic

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variability. There are therefore grounds for speculation that there exists in nature an infinite number of species and subspecies of leptospiral saprophytes. Their inclusion in the classification schemes is next to impossible and does not seem reasonable as in the long run such inclusion could result in taxonomic chaos.

Table 1. Leptospiral strains in which no 677 bp LipL32 gene fragment was amplifiedby PCR with LEP21/ LEP22 primers. Strain

Genomospecies

Serogroup

Serovar

H

Turneria parva

Turneria genus

parva

3055

Leptonema illini

Leptonema genus

illini

Patoc I

L. biflexa

Semaranga

patoc

CH-11

L. biflexa

Andamana

andamana

Parapatan

parapatan

T

Parapatan Bairam-Ali

-

Bairam ali

bairam-ali

10

L. inadai

Lyme

lyme

Enr-86

L. inadai

Lyme

lyme

Enr-87

L. inadai

Lyme

lyme

-

Li130

L. inadai

Manhao

lichuan

1342K

L. santarosai

Shermani

shermani

ICF

L. meyeri

Ranarum

ranarum

CDC

L. wolbachi

Codice

codice

Waz Holland

Genomospecies 3

Holland

holland

Sao Paulo

Genomospecies 5

Semaranga

saopaulo

Evidently, DNA-DNA relatedness or sequence-based data may not be sufficient for the description of a new species and, all the more, higher taxons. Basic phenotypic and ecological characteristics as well as the source of isolation should also be considered. Thus, the current taxonomy of leptospires needs to be revisited, also from this perspective. In the context under discussion here, the recent suggestion of creating, alongside sequence databases and international collections of bacteria, an Internetbased orphan bacterium repository intended to incorporate any relevant criterium for the accurate identification of novel bacteria [33] deserves special consideration.

2. Pathogenic Leptospires (L. interrogans sensu lato) At least 17 genomospecies have been recognized so far within the Leptospiraceae family, seven of these with proven pathogenic potential [34, 35]. Pathogenic leptospires are known to be highly divergent as to their virulence for human beings and a wide range of animal species. Severe diseases with high case fatality rates (up to 20%) or resulting in multiorgan complications and late clinical sequelae in affected humans are

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usually caused by the agents belonging to L. interrogans sensu stricto genomospecies (copenhageni, icterohaemorrhagiae, canicola, valbuzzi and other serovars) [1, 2, 3, 36, 37]. At the other extreme are leptospires with very low virulence, for example, L. borgpetersenii, serovar tarassovi [2, 39]. Pathogenic leptospires also differ as to their within-host survival patterns, in particular, tissue tropism. Thus, experimental evidence, obtained in an experimental model (Syrian golden hamsters) using culture and PCR techniques, testifies that lowvirulent L. borgpeterseni (serovars tarassovi and hanka), in contrast to serovars copenhageni and grippotyphosa, fail to display neurotropism [40]. Clinical data also confirm that neurological symptoms during acute phase of the disease and late neurological sequealae are usually observed in patients suffering from leptospirosis caused by the agents with selective tropism to nervous tissues [3]. Pathogenic leptospires display strict or relatively strict host specificity as with L. interrogans serovar copenhageni with Rattus norvegicus or serovar hardjo with cattle as preferred hosts. However, according to epidemiological and epizotological observations, leptospires not infrequently manage to breach the species barrier. For example, L interrogans, canicola serovar, re-emerging in Russia, occasionally causes leptospirosis infection not only in the principal host (dog) but also in cattle and swine. In some cases, it resulted in water-borne outbreaks of Stuttgart disease among humans [41]. Epizootological and experimental evidence was obtained showing that breaching the barrier of host specificity may occur when the mammalian host is co-infected with genetically and antigenically divergent leptospires [42, 43]. Clear differences may also be found among pathogenic leptospires in between host survival patterns. According to the results of earlier ecological experiments carried out under natural conditions with a focus on leptospirosis, the maximum period of survival for L. interrogans serovar grippotyphosa in soil was as many as 279 days, while for L. borgpetersenii serovar nero, the maximum period of survival was only three days [44]. Besides high survival potential in environmental ecosystems, grippotyphosa leptospires display other features of so-called sapronotic agents, namely, wide mammalian host range and genetic and antigenic heterogeneity on infrasubspecies level [24, 39].

3. Conclusions Within the Leptospira genus we may not be dealing with just two ecological types, parasitic and free-living. There is no clear line of delineation between them: there may be strains that occupy an intermediate position between independent and symbiotic. A growing body of evidence suggests that pathogenic leptospires display not only substantial antigenic and genomic heterogeneity at species, subspecies, and infrasubspecies levels, but also differ in within-host and between-host survival patterns. These findings support future studies of comparative genomics of leptospires differing in their mode of existence as well as revealing epidemiological and clinical implications of their biodiversity.

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[26] Haake D.A., C. Martinich, T.A. Summers, et al. 1998. Characterization of leptospiral outer membrane lipoprotein LipL36: downregulation associated with late- log- phase growth and mammalian infection. Infect. Immun. 66: 1579–1587. [27] Ananyina Yu.V., A.P. Samsonova, V.V. Lebedev, E.M. Petrov, E.M. Esipov. 2000. Genodiagnosis of acute and persistent leptospiral infection.2000. J.Microbiol. (Moscow) 4 (supplement):23–26. [28] Perolat P., R.J Chappel, B. Adler, G. Baranton, D.M. Bulach, M.L. Billinghurst ML, et al. 1998. Leptspira fainei sp.nov., isolated from pigs in Australia. Int. J. Syst. Bacteriol. 48:851–858. [29] Chappel RJ, D.A. Khalik, B. Adler, D.M. Bulach, S. Faine, P. Perolat. 1998. Serological titres to Leptospira fainei serovar hurstbridge in human sera in Australia. Epidemiol. Infect. 121:473–475. [30] Yersin C, P. Bovet, F. Merien, T. Wong, J. Panowsky, P. Perolat. 1998. Human Leptospirosis in the Seychelles (Indian Ocean): a population based study. 1998. Am. J. Trop. Med. Hyg. 59:9333–40. [31] Arzouni J.P., Ph. Parola, B. La Scola, D. Postic, Ph. Brouqui, and D. Raoult. 2002. Human infection caused by Leptospira fainei. Emerg. Inf. Diseases.8:865-868. [32] Petersen AV, K. Boyde, J. Blom, P. Schliting, and K.A. Krogfelt. 2001. First isolation of Leptospira fainei serovar hurstbridge from two human patients with Weil’s syndrome. J. Med. Microbiol. 50:96– 100. [33] Drancourt M., and D. Raoult. 2005. Sequence-based identification of new bacteria: a proposition for creation of an orphan bacterium repository. J. Clin. Microbiol. 43:4311–4315. [34] Yasuda P.H., A.G. Steigerwalt, K.R. Sulzer, A.F. Kaufmann, F. Rogers, and D.J. Brenner. 1987. Deoxyribonucleic acid relatedness between serogroups and serovars in the family Leptospiraceae with proposals for seven new Leptospira species. Int. J. Syst. Bacteriol. 37:407–415. [35] Brenner D.J., A.F. Kaufmann, K.R. Sulzer, A.G. Steigerwalt, F.C. Rogers, and R.S. Weyant. 1999. Further determination of DNA relatedness between serogroups and serovars in the familyLeptospiraceae with a proposal for Leptospira alexanderi sp.nov. and four new Leptospira genomospecies. Int. J. Syst. Bacteriol. 49:839–858. [36] Vijayachari P., S.C. Sehgal, M.G.A. Goris, W.J. Terpstra, and R.A. Hartskeerl. 2003. Leptospira interrogans serovar Valbuzzi: a cause of severe pulmonary haemorrhages in the Andaman Islands. J. Med. Microbiol. 52:913–918. [37] Covic A., D.J. Goldsmith, P. Gusbeth Tatomir, A. Seika, and M. Covic. 2003. A retrospective 5-year study in Moldova of acute renal failure due to leptospirosis: 58 cases and a review of the literature. Nephrol. Dial. Transplant. 18:1128–1134. [39] Ananyina Yu.V., and A.P. Samsonova. 2000. Interspecies and subspecies diversity among leptospires: phylogenetic, taxonomic and ecological aspects. Bull. Russ. Acad. Med. Sci. (Moscow) Medicine ed. 1:18–21. [40] Samsonova A.P., Yu.V. Ananyina, and M.Yu. Aksenov. 1994. Polymerase chain reaction in studying host persistence of leptospires. Molec. Genet. Microbiol. Virol. (Moscow) 1:19–23. [41] Ananyina Yu.V. Emerging and e-Emerging Infections in Russia: Current Trends. 2000. In: Protection against Microbial Threats. Conf. Proceed. SMI. Sweden, 31. [42] Ananyin V.V. 1964. Leptospirosis of humans and rodents caused simultaneously by leptospires of two serological types. J. Microbiol. (Moscow), 12:74–78. [43] Samsonova A.P., E.M. Petrov, N.V. Vishivkina, Yu.V. Ananyina. 2003. A new approach to studying leptospira host persistence during mixed leptospiral infection. J. Microbiol. (Moscow). 4 (supplement):37–39. [44] Zaitsev S.V., and Chernukha Yu.G. 1988. Survival of leptospires in the soil of a natural focus. Ecology of sapronotic agents. M., Moscow. 85–94.

Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Characterization of Borrelia burgdorferi sensu lato from Czech Patients and Ticks by Culture and PCR-Sequence Analysis a

Dagmar HULINSKA a,1, Martin BOJAR b, Václav HULINSKY c National Institute of Public Health, National Research Laboratory for Borreliosis, Prague, Czech Republic b Neurological Clinic of 2nd Faculty Hospital, Charles University, Prague, Czech Republic c Clinic of Internal Medicine Regional Hospital, Nymburk, Czech Republic Abstract. Lyme disease has continued to spread in the Czech Republic, and there have been increasing numbers of cases in central and eastern Bohemia and Moravia. We have shown that Borrelia burgdorferi sensu lato causes infection by migration through connective tissues of the skin, heart, liver, and synovium without inflammation and by adhesion to host cells. Twenty-three isolates of B. burgdorferi sensu lato originally cultured from cerebrospinal fluid, blood, and tissue samples were characterized phenotypically by immunosorbent electron microscopy and Western blots and genotypically by sequence analysis of the outer surface (Osp) A and C gene fragments. Comparison of genotyping of borreliae performed directly in patients and in Ixodes ricinus ticks collected in the same endemic regions with typing of cultures revealed that a large proportion of directly typed samples had more heterogenic Borrelia types or combined sequence variants. In addition, B. garinii OspA types 6 and 5 were in coinfection with B. valaisiana and rarely with B. burgdorferi, but OspA subtype 4 was not found in the mixure. More frequent and heterogenic direct proof of B. burgdorferi sensu lato in CSF and blood versus culture reflects either different behavior and exigencies of different genospecies or intergenospecies exchange caused by human and animal migration. Keywords. Electron microscopy, sequence analysis, Borrelia, patients, ticks

Introduction Lyme borreliosis (LB) is an emerging multisystem illness of humans and animals and is caused by tick-borne spirochetes currently assigned to four species pathogenic to humans, Borrelia burgdorferi sensu stricto, B. garinii, B. afzelii, and B. valaisiana [1, 2, 3]. The spirochetes were first isolated from the midgut of ticks and from human 1 Corresponding author: Dagmar Hulinska, National Institute of Public Health, National Research Laboratory for Borreliosis, Prague, Czech Republic; E-mail: [email protected].

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blood in modified Kelly medium [4]. Modification of this Barbour-Stoenner-Kelly (BSK-II) medium is routinely used to culture Borreliae in vitro from ticks and human tissue and body fluids [5, 6]. Culturing of skin biopsy specimens showed recovery of Borrelia in 57-86% of patients [7]. Recognizable changes have been found in the genetic and antigenic character of spirochetes after subcultivation in vitro [8]. Several genospecies and seven outer surface protein A (OspA) types have been described by Wilske et al. [9]. Increasing interest has also been focused on OspC because it is a protective antigen in animals, and because the OspC genes in B. burgdorferi sensu lato have been found to show diversity among strains [10-12]. The knowledge of the existence of variable strains and their OspA and OspC types in different regions of Europe are of importance for the selection of suitable antigens for diagnostic assays [13]. Genotyping and phenotyping of B. burgdorferi sensu lato spirochetes in Czech ticks and patients by cultivation versus direct sequence analysis from tissues, blood, and cerebrospinal fluid (CSF) has permitted identification of factors influencing borreliosis in endemic regions of central and eastern Bohemia and Moravia. It also resulted in the confirmation of a hypothesis of some authors as to whether different genospecies exert different pathognostic effects and whether clinical manifestations [14] can be suitably resolved with techniques [15] such as polymerase chain reaction (PCR) and sequence analysis [16] of partial gene fragments coding OspA and OspC proteins directly from the host specimens. Heterogeneity of B. burgdorferi sensu lato has been demonstrated by an OspA-type-specific PCR in synovial fluids [16, 17]. Heterogeneity among Borrelial strains might have important implications for understanding their epidemiology [18, 19, 21] and their correlation with different clinical manifestations and the course of self-limiting versus chronic disease [20].

1. Materials and Methods For the present study, 23 strains isolated from 100 patients were used. Isolation of 23 strains of B. burgdorferi sensu lato from tissue specimens, blood, and CSF was carried out in BSK II medium. Blood and skin strains were isolated from 45 early-stage Lyme disease patients at the time of appearance of erythema migrans (EM). CSF strains were isolated from 25 patients with EM and facial palsy, and from 21 patients with serologically confirmed late-stage borreliosis at the time of presentation with Bannwarth Syndrome and lymphocytic meningitis. Cardiac complications in patients with Lyme arthritis were present in nine patients, two of whom had cardiac biopsies. B. burgdorferi were grown at 34 qC in modified BSK-H medium (Sigma-Aldrich,Ltd., Germany) to a cell density of 106 cells per ml. A total of 1299 ticks collected in 2004 were used in this study; 700 adults (479 females) and 599 nymphs were examined individually by dark-field microscopy (DFM). The ticks were washed with alcohol and twice with sterile phosphate-buffered saline (PBS), and then transferred to sterile Eppendorf vials and partially homogenized with 1.5 ml of sterile PBS using sterile razors and pestles. One portion (2 x 15 μl) was observed with dark-field microscopy, and the other portion (0.5 ml) was cultured when dark-field microscopy (DFM) was positive. The remaining suspension was processed for PCR or Real-Time PCR. Cultivation of ticks was performed on 266 DFM positive samples with a density of spirochetes of >10 per field in BSK H medium, with 8% rabbit serum (Sigma) and 1.5% Bacto gelatin (Difco Laboratories, Germany).

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Harvested and washed borrelial strains cultivated from patients and ticks were resuspended in buffer containing 2.5% sodium dodecyl sulfate (SDS) and 2mercaptoethanol. After boiling for 5 min, the samples were subjected to electrophoresis in 10% polyacrylamide gel (PAGE). Proteins were stained with Comassie blue or electroblotted onto nitrocellulose sheet, blocked with 0.05% Tween 20, and probed with polyclonal or monoclonal antibodies against OspA and flagellin. The resulting immunocomplexes were labeled with protein A-horseradish peroxidase or with coloidal gold conjugates (Janssen Life Sciences, Redding, CA USA) and detected by a chromogenic assay or by immunosorbent (ISEM) negative staining using a Jeol 100CX electron microscope [21, 22]. Anti-Borrelia antibodies in serum and CSF samples were detected by separate IgG and IgM indirect commercial EIA (TEST-Line, Ltd., CZ) and immunoblots (BAG Med.AG, Germany and Biowestern Ltd., CZ). Paired undiluted serum and CSF (diluted as recommended to match CSF immunoglobulin concentrations) samples were evaluated. A CSF/serum ratio t 1.3 was taken as evidence of local antibody production. Evaluation of the serological assay was made by using confirmatory sera kindly donated by the United States Center of Disease Control and Preventation (CDC). Samples from these 100 patients with suspect clinical, serological, and anamnestic data of borreliosis from three endemic regions were used for direct sequence analysis. The control group included four CSF and six blood samples from patients with malignancies. DNA was purified from 23 patient strains, from 46 CSF samples, 45 blood and skin samples of patients with EM, 9 blood and heart samples, 233 DFM positive ticks, and 31 tick strains with the QIAamp DNA Micro Kit (Qiagen) and/or with High Pure PCR Template Preparation Kit (Roche). Nucleic acids of patient and tick samples, and the B31 control strain, were amplified with primer sets for OspC and OspA genes (24, 17); some samples were amplified with recA primers. PCR reagents from the Qiagen Taq PCR Master Mix Kit (Qiagen GmbH, Hilden) or from the Gene Amp kit and LightCycler FastStart DNA Master SYBR Green Kit (Roche Diagnostics, GmbH, Germany) were used as recommended by the manufacturers. PCR amplification was performed with a PTC 200 Peltier Thermal Cycler and with LightCycler (Roche). Nucleotide sequence homology searches were made through the NCBI BLAST network service. Sequence analysis of amplicons (392 bp) was performed from nested PCR of the OspA gene corresponding to the upstream primer 5' GCA AAA TGT TAG CAG CCT TGA T 3' and the downstream primer 5' CTG TGT ATT CAA GTC TGG TTC C 3' and products 295 bp of the OspC gene corresponding to the upstream primer 5' CGT CTT TCA GGT TTT TTT GGA ATT CCT AGG CGG CTG 3' and the downstream primer 5' GTG CGC GAC CAT ATG AAA AAG AAT ACA TTA AGT GCG 3'. PCR amplicons were purified using MultiScreen PCR Kit (Millipore, Bedford). PCR amplified DNA was prepared for sequencing on the MJ Research PTC thermocycler (96qC for 20 s, 50qC for 20 s, 60qC for 4 min, 30 cycles). Direct sequencing was carried out by the CEQ 2000 LX automatic sequencer. Sequences were aligned and analysed using CLUSTALW. The phylogenetic tree was generated with TRECON software, using the Kimura algorithm to calculate the genetic distances and the neighbor-joining method to infer the three topologies. The following GenBank accession numbers were used to retrieve the corresponding OspA sequences: AF 227319 for B. garinii, OspA serotype 3, X85440, AF227317 for B. garinii, OspA serotype 5, AF227319 for OspA serotype 4, U20360 for B. burgdorferi sensu stricto, and U20356 for B. afzelii. The collected sequences

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were analysed with the BLAST and DNAStar package. GenBank accession numbers used to retrieve the OspC sequences: AF501317 for B. garinii, L42880 for our strain KL11, AF416425 for B. garinii, X81526 for B. garinii (strain PWabsou), X84780 for B. garinii (strain SL10), X84781 (strain SL20), AJ236907 for B. garinii (strain Psh), X73624 for B. burgdorferi (strain DK26), and AF230185 for B. afzelii (strain Pgau).

2. Results Twenty-three strains of B. burgdorferi sensu lato originally cultured from skin, heart, placenta, CSF, and blood were characterized phenotypically with monoclonal antibodies using Western blots and ISEM. Positive cultures were obtained in 6 CSF samples from 25 patients with EM and facial palsy and from 3 of 21 patients with Bannwarth Syndrome. Cultures were positive from CSF from 6 to 22 days before therapy. Strain M76 was cultured from a patient with meningoencephalitis at the same time as borrelial flagella were demonstrated by ISEM (Figure 1). Four blood and nine skin strains were isolated from 45 early-stage Lyme disease patients at the time of appearance of erythema migrans (EM). The Borrelia isolated from CSF were slightly different in the first passage with regard to the proteins they expressed. Protein profiles of some blood and skin strains are shown in Figure 2A and of some CSF strains are shown in Figure 2B. All the analysed strains expressed OspA which differed in their molecular mass in SDS PAGE and in their reaction with antibodies. Expression of OspA, OspB, and OspC varied in different strains

Figure 1. Immunoelectron microscopy of borrelial flagella in CSF using monoclonal antibody H2754.

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Figure 2. Western blots show different reactions of IgG antibodies with antigens of different strains isolated from the skin and blood (A) and from CSF (B). OspA and OspC had different molecular weights. Lanes 3, 4, 7, 9 (A) and lanes 2, 5 (B) show reactions of B. afzelii strains, lanes 5, 6, 8 (A), reactions of B. valaisiana strains and lanes 1, 9 (A) and 1, 3, 6 (B) of B. garinii strains, lane 10 (A) shows B. burgdorferi strain and lanes 7, 8, 9 (B) show reactions of Neisseria meningitis and Leptospira sp. controls.

Figure 3. Electron microscopy of young growing borrelia in CSF form gamma-like structure (negative staining, x19 000).

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Strains classified as B. valaisiana (Figure 2A, lanes 5, 6) contained OspA proteins of size ranges 32-35 kDa. These did not react with IgG antibodies of patients with acrodermatitis chronicum atrophicans and also did not react with mAB H5423 but had OspC similar to B. garinii strains. The serum antibody index was positive in 29 patients out of 46 with neurologic symptoms. Sixteen met the criteria of a IgG CSF/serum ratio, and four had a CSF/Ab ratio t 1.5. Nine were positive by IgM criteria alone and had more reactive immunoblots. Pieces of antigen, membrane vesicles, and rarely, convoluted small Borrelia-like organisms, were detected with the ISEM method in 33% of 46 CSF shown in Figure 3. Granules were detected in some of 45 blood samples in reaction with monoclonal antibody and also in the skin tissue fibroblasts (Figure 4) Analysis of the partial OspA sequence of human isolates showed that 12 isolates (52 %) belonged to the species B. garinii, seven isolates (30.4%) to B. afzelii, three isolates (13%) to B. burgdorferi sensu stricto, and one isolate to B. valaisiana. B.garinii isolates M76, 31M, and Kc261 were OspA-type 4 . The only single nucleotide difference was in B. garinii strain M76. B. afzelii 51E, 78Kc, and strain Kc90 were isolated from the blood, and EM was evaluated as OspA-type 2 with an identity of OspC sequence to strain WABsou (X81526). Partial structures of OspA genes from different CSF isolates are shown in Table 1. The OspC amplicons from these B. garinii isolates were identical at the nucleotide level. Alignment of the sequences determined here with the OspA and OspC sequences available in the database revealed that they were nearly identical to strains Psh and PBi (accession No. AJ236907, X69595). A phylogenetic tree (Figure 5) shows relationships between partial OspC sequences of the B. garinii OspA-type 3 strains (61E, 144E, and K11) from EM similar to strain TIs1 (X81525) and differences from B. garinii, OspA-type 6 strains ( H13, M76), as well as from B. afzelii strains (51E, KC90, 78 KC).

Figure 4. Electron microscopy of a tangential section of granules inside outer membrane which formed cystlike structure with the rest of flagella (uranyl acetate-lead citrate stain; bar represents 10 μm).

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Table 1. Comparison of partial sequences of different OspA-types in CSF isolates of B. burgdorferi s. l. A (OspA-type 5)

ATTTGAAATCTTCAAAGAAGATGGCAAAACATTAGTATCAAAAAAAGTAA

C (OspA-type 4)

ACTTGAAGGTGAAAAAACTGACAAAAGTAAAGTAAAATTAACAATTGCTG

D (OspA-type 3)

ATTTGAAATTTTCAAAGAAGATGGCAAAACATTAGTGTCAAGAAAAGTAA

B (OspA-type 2)

ATTCGAACTTTTCAAAGAAGATGGCAAAACATTAGTGTCAAGAAAAGTAA

E (OspA-type 1)

ACTTGAAGTTTTCAAAGAAGATGGCAAAACACTAGTATCAAAAAAAGTAA

F (OspA-type 6)

ACTTGAAGGTGAAAAAACTGACAAAAGTAAAGTAAAATTAACAATTGCTG

1-144E-B 621 1000 1-61E-B 02-K11

782

03-K6 873 1000

963

2-BITS-B 13-BR14

4-261-B 1000 721

581

4-179KcB 14-M31 04-KC261

1000 1000

08-51E 07-78KC

1000 01-KC90 06-M76 05-H13

Figure 5. Phylogenetic tree shows of OspC sequencing of 13 human strains and 1 tick strain of B. burgdorferi. Each OspC sequence is identified by the strain number and the source it was extracted from (M for CSF, KC for citrated blood, E for EM, B for whole blood, H for heart, K for skin, Kl for tick). Horizontal bar represents 0. 1% sequence diversity.

B. afzelii strain 93M was one rarely isolated strain from CSF of seropositive patients with meningitis following EM. The similar OspC sequence had strain A10 from the skin and strain A541 from the blood of patients with ACA and two strains isolated from mice (A04-3B, 2B). Their phylogenetic relationships (Figure 6) are close as seen by the bootstrap numbers. They were similar to strain DK22 (X84769) and different from skin isolates 51E and K275 by the substitution of G for a C, and A for T at positions corresponding to nucleotides 230 and 340 in B. afzelii L42871.

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Figure 6. Phylogenetic tree generated from an alignment of the first 600 amino acids of OspC. The tree is labeled with bootstrap numbers at branch points which show relationships between direct sequencing results for 7 B. burgdorferi sensu stricto in the blood (designated A3) with strains 176, and 192 with OspC sequences of B. valaisiana and B. bissetti from ticks. Some heterogeneity was observed between OspC of B. afzelii from mice and patients and B. garinii from patients and tick.

Figure 7. Electron microcroscopy of ultrastructure of B. burgdorferi sensu stricto. in a heart biopsy shows borrelia cell in the intermysium covered with thin outer layer (uranyl acetate, x 29 000).

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B. burgdorferi sensu stricto was isolated from a biopsy of the heart in a patient with cardiovascular involvement after EM. These Borreliae were demonstrated in the intermysium by ISEM (Figure 7). Strains 192 and 176 had sequence similarity to B. burgdorferi 297 (U20360) with small phylogenetic distances constructed from directly sequenced B. burgdorferi DNA amplicons from isolates from ticks and patients (Figure 6) Using OspA-type specific PCR we identified also OspA types 1, 2, 3, 5, and 6 among 31 strains isolated from ticks. Analysis of Borrelia DNA directly isolated from Ixodes ricinus ticks showed higher positivity rates in adult females as compared to nymphs. All positive ticks in DFM (233) were analysed for the presence of B. burgdorferi sensu lato by specific PCR tests with OspA primers. B. garinii was most prevalent (48 %), followed by B. afzelii (30 %), B. burgdorferi sensu stricto (17 %), B. valaisiana (4 %), and B. bissettii-like (0.8 %). B. spielmanii A16S was identified in two ticks. The phylogenetic tree (Figure 8) constructed on the basis of the OspA gene sequences from ticks showed greater identity with B. afzelii strain VS461, B.garinii strain WaBsou, and B. valaisiana strain VS116. The neighbor-joining tree is resolved and shows 8 distinct sequence types among the tick-derived samples. Multiple infections with different genospecies or subtypes were detected in 18 of 233 PCR positive ticks. Double infection with B. garinii and B. afzelii was often detected.

Figure 8. Neighbor-joining tree based on comparison of OspA sequences isolated from ticks and patients showed heterogeneity between B. garinii, B. afzelii and B. burgdorferi strains. B. valaisiana 117E from skin did not share the same branch as the same species from tick. The numbers are the results of 1,000 bootstrap resamplings (values of less than 80 are not shown).

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B. valaisiana strains and/or specific DNA were identified from I. ricinus ticks and differed from EM isolate 117E. B. bissetii of the same sequence identity was found in EM (strain E50) and also in ticks (not shown). Direct PCR-sequence analysis of DNA isolated from skin, CSF, and blood samples from 100 patients with borreliosis and from PCR-positive ticks also showed deduced different amino acid sequences corresponding to B. garinii , OspA-types 5, 4, 3 and 6, respectively, and to B. afzelii and B. burgdorferi OspA-types 1 and 2 as has been detected between isolated strains. B. garinii, serotypes 4 and 5, were found most frequently in the CSF of the patients. Serotype 4 was not detected in the ticks. In 16 of the 49 samples (33%) of patients from the eastern region of the Czech Republic, only B. garinii sequences were identified. On the other hand, in the central region, 10 CSF (20%) samples contained sequences similar to B. burgdorferi sensu stricto and 8 CSF (16%) samples had sequences similar to B. garinii. Identical sequences in B. afzelii from skin isolates of six patients and two CSF samples (12%) from patients of the central region, and in four patients from Moravia where two CSF (6%) samples also contained B. burgdorferi sensu stricto. Ambiguous OspA sequences were verified by species-specific PCR targeting of OspA as type 5, 6 with B. valaisiana. OspA type 4 was not found in the mixure. In 20 patients from the eastern region, samples in which sequences are known were taken, as well as from 19 patients from the central region and 10 from Moravia were analysed with rec A primers using Real-Time PCR (Figure 9). Multiple infections with very close B. garinii serotypes were detected by Real-Time PCR in four (8 %) patient samples from the east and in two (4 %) patients from Moravia.

Figure 9. Melting curve analyses of amplification products of recA gene from DNA samples of patients with EM. The melting temperature (Tm) of the double-stranded fragment is visualized by plotting the negative derivative of the change of fluorescence (dF) divided by the change of temperature (dT) in relation to the absolute temperature. Melting curve results in a peaks which permited identification of the fragment-specific Tm 82-83 °C of B. garinii and B. valaisiana and Tm 84. 5 ºC of B. afzelii.

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3. Discussion Comparison of genetic heterogeneity in B. burgdorferi sensu lato as determined by culture versus direct PCR-sequence analysis performed in 23 strains and 49 PCR positive patient samples has implications for understanding the epidemiology and clinical diversity of borreliosis in the Czech regions. Several attempts have been made to classify Borrelia on the basis of phenotypic and genotypic traits [2, 9, 10, 15, 19]. Sequence analyses from skin, CSF, blood, heart, and placenta isolates cultured in BSKH medium were the ultimate approach for determining whether strains were similar or different and also how they differed from 49 directly sequenced skin, CSF, and blood isolates. Our culture success rate was 50% (5 of 9) in the skin. Karlsson et al. [25] reported that success in CSF culture was the highest (10%) in early disease and lower in replapsing chronic infections. Several publications regard positive PCR results as a proof of Borreliae responsible for clinical manifestations. Very high sensitivity rates of PCR have been published [20, 26, 27]. Patients with neuroborreliosis did not always remember tick bite or EM as were mentioned in studies concerning association between the early and late cutaneous manifestations and B. afzelii infection [7, 28, 30]. We have isolated B. garinii from skin, CSF, and blood, rather than B. burgdorferi sensu stricto or B. afzelii. Results acquired by direct PCR-sequence analysis of the OspA gene from 49 patient samples indicated a more genotypically heterogeneous group of Borrelia inclusive of B. burgorferi sensu stricto, B. valaisiana, and more predominantly, B. garinii and B. afzelii. Substitution of genospecies differed in geographically distinct regions. The variability of OspC is greater than OspA as Livey et al. [12] have shown. To date, only species in the B. burgdorferi sensu lato complex are known to cause neuroborreliosis in Europe. However, some atypical borreliae such as B. valaisiana or borreliae closely related to the previously designated genomic group DN 127 have been isolated from CSF. B. bissettii, described by Postic et al. [22], clustered separately from B. burgdorferi sensu stricto. B. bissettii strain E50 was isolated in the Czech Republic for the first time. In this study, the classification based on OspA serotyping [9,17] was done in accordance with recognized genotyping methods. We found OspA serotypes 4, 5, 6 primarily in CSF in patients from the eastern region of the Czech Republic. Each putative species would be expected to occur in its own geographic area. B. garinii was predominant in the eastern region where lakes, woods and pastures, sapling nesting sites and flight zones of migratory birds are present, and where B. valaisiana was also found. B. burgdorferi sensu stricto was found in samples from patients from the central region around Prague and in patients from Moravia around Brno where there are rodents and tourist areas. Mixtures of related B. garinii serotypes were demonstrated by direct PCR analysis and by real-time PCR of OspA, but not by initial cultures. Culture isolation of different spirochetes from the same patient has been reported [28, 29, 31]. Lebech et al. [32] found a mixture of B. garinii and B. afzelii by direct genotyping. Demaerschalch et al. [31] reported that 8 of 18 patients with neuroborreliosis were infected with more than one genospecies. Whereas only B. garinii has been isolated from the CSF of patients from Scandinavia [32], analysis of 11 CSF isolates from patients in Germany demonstrated the presence of two species [56]. Different OspA sequences in isolates from the CSF of children with neuroborreliosis have been described [33]. The prevalence of B. garinii-associated OspA types has been described [10] for strains from Europe. We obtained similar results using type-specific PCR [17].

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B. garinii serotype 5 was among the most prevalent of our isolated strains and in direct PCR seqencing performed on CSF samples.

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G. Baranton, Postic D, SaintGirons I, Boerlin P, Piffaretti JC, AssousM, Grimont PAD, Delineation of Borrelia burgdorferi sensu stricto, Borrelia garinii sp.now. and group VS461 associated with Lyme borreliosis. Int. J. Syst. Bacteriol. 42 (1992), 378–383. G. Wang , VanDam AP, Le Fleche A, Postic D, Peter O, Baranton G, de Boer R, Spanjaard L, Dankert J, Genetic and phenotypic analysis of Borrelia Valaisiana sp.now. Int. J. Syst. Bacteriol 47 (1997), 926–932. V. Preac-Mursic, Wilske B, Schierz G, European Borrelia burgdorferi isolated from humans and ticks. Culture conditions, and antibiotic susceptibility. Zbl. Bact. 263 (1986), 112–118. J.L.Benach, Bosler EM, Hanrahan JP, Coleman JL, Habicht GS, Bast TF, Cameron DJ, Ziegler JL, Barbour AG, Burgdorfer W, Edelman R, Kaslow RA: Spirochetes isolated from the blood of two patients with Lyme disease. N Engl. J. Med. 308 (1983), 740–742. B.W.Berger, Johnson RC, Kodner C, Coleman L, Cultivation of Borrelia burgdorferi from erythema migrans lesions and perilesional skin. J. Clin. Mocrobiol. 30 (1992), 359–361. T.G. Schwan, Burgdorfer W, Garon CF, Changes in infectivity and plasmid profile of the Lyme disease spirochete, Borrelia burgdorferi as a result of in vitro cultivation. Infect. Immun. 56 (1988), 1831– 1836. B. Wilske, Preac-Mursic V, Gobel UB, Graf B, Jauris-Heipke S, Soutscheck E, Zumstein G, An OspA serotyping systém for Borrelia burgdorferi based on reactivity with monoclonal antibodies and OspA sequence analysis. J. Clin. Microbiol. 31 (1993), 340–350. B. Wilske, Busch U, Eiffert H, Fingerle V, Pfister HW, Rossler D, Preac-Mursic V, Diversity of OspA and OspC among cerebrospinal fluid isolates of Borrelia burgdorferisensu lato from patients with neuroborreliosis in Germany. Med. Microbiol. Immunol. 84 (1999), 195–201. U. Busch, Hizo-Teufel C, Boehmer R, Fingerle V, Nitschko H, Wilske B, Preac-Mursic V, Three species of Borrelia burgdorferi sensu lato (B.burgdorferi sensu stricto, B.afzelii, and B.garinii) identified from cerebrospinal fluid isolates by pulse-field gel electrophoresis and PCR. J. Clin. Microbiol. 34 (1996), 1072–1078. I. Livey, Gibbs CP, Schuster R, Dorner F, Evidence for lateral transfer and recombination in OspC variation in Lyme disease Borrelia. Mol. Microbiol. 18 (1995), 257–269. U. Hauser, Krahl H, Peters H, Fingerle V, Wilske B, Impact of strain heterogeneity on Lyme disease serology in Europe, Comparison of enzyme-linked immunosorbent assays using different species of Borrelia burgdorferi sensu lato. J. Clin. Microbiol. 36 (1998), 427–436. A.P. Van Dam, Kuiper H, Vos K, Widjojokusumo A, De Jongh BM, Spanjaard L, Ramseloar AC, Kramer MD, Dankert J, Different genospecies of Borrelia burgdorferi are associated with distinct clinical manifestations of Lyme borreliosis. Clin. Infect. Dis. 17 (1993), 708–717. M.M. Picken, Picken RN, Han D, Cheng Y, Ruziþ SE, Cimperman J, Strle F, A two year prospective study to compare culture and polymerase chain reaction amplification for the detection and diagnosis of Lyme borreliosis. Mol. Pathol. 50 (1997), 186–193. S. Priem, Rittig MG, Kamradt T , Burmester GR, Krause A, An optimized PCR leads to rapid highly sensitive detection of Borrelia burgdorferi in patients with Lyme borreliosis. J. Clin. Microbiol. 35 (1997), 685–690. V. Vasiliu, Herzer P, Rossler D, Lehnert G, Wilske B, Heterogeneity of Borrelia burgdorferi sensu lato demonstrated by an OspA-type–specific PCR in synovial fluid from patients with Lyme arthritis. Med. Microbiol. Immunol. 187 (1998), 97–103. S. O´Connell, Granstrom M, Gray JS, Stanek G, Epidemiology of European Lyme borreliosis. Zbl. Bakteriol. 287 (1998), 229–240. G. Wang, Van Dam AP, Schwartz I, Dankert J, . Molecular typing of Borrelia burgdorferi sensu lato, Taxonomic, epidemiological and clinical implications. Clin. Microbiol. Rev. 12 (1999), 633–653. J. Oksi, Marjamaki M, Nikoskelainen J, Viljanen MK, Borrelia burgdorferi detected by culture and PCR in clinical relapse of diseminated Lyme borreliosis. Annal. Medicine 31 (1999), 225–232. D.W. Dorward, Schwan TG, Garon GF, Immune capture and detection of Borrelia burgdorferi antigens in urine, blood, or tissues from infected ticks, mice, dogs, and humans. J. Clin. Microbiol. 29 (1991), 1162–1170

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[21] D. Hulínska, Krausova M, Janovska D, Rohacova H, Hancil J, Mailer H, Electron microscopy and the polymerase chain reaction of spirochetes from the blood of patients with Lyme disease. Centr. Europ. J. Pub. Heal. 1 (1993), 81–85. [22] D. Postic, Ras N.M., Lane R.S., Hendson M., Baranton G., Expanded diversity among Californian borrelia isolates and description of Borrelia bissettii sp. nov. (formely Borrelia group DN127). J.Clin. Microbiol. 36 (1998), 3497–3504. [23] D. Hulinska, Votypka J, Valesova M, Persistence of Borrelia garinii and Borrelia afzelii in patients with Lyme arthritis. Zbl. Bakteriol. 289 (1999), 301–318. [24] M. Karlsson, Hovind Hougen K, Svenungsson B, Stiernstedt G, Cultivation and characterization of spirochetes from cerebrospinal fluid of patients with Lyme borreliosis. Journal of Clinical Microbiology 28 (1990), 473–479. [25] B.J. Luft, Steinman CR, Neimark HC, Muralidhar B, Rush T, Finkel MF, Invasion of the central nervous systém by Borrelia burgdorferi in acute disseminated infection. J. Am. Med. Assoc. 267 (1992), 1364–1367. [26] B.I. Schmidt, PCR in laboratory diagnosis of human Borrelia burgdorferi infections. Clin. Microbiol. Revi. 10 (1997), 185–201. [27] D. Liveris, Varde S, Iyer R, Koenig S, Bittker S, Cooper D, McKenna D, Nowakowski J, Nadelman RB, Womser GP, Schwartz I, Genetic diversity of Borrelia burgdorferiin Lyme disease patients as determined by culture versus direct PCR with clinical specimens. J. Clin. Microbiol. 37 (1999), 565– 569. [28] B. Jaulhac, Heller R, Limbach FX, Hansmann Y, Lipsker D, Monteil H, Sibilia J, Piémont Y, Direct molecular typing of Borrelia burgdorferi sensu lato species in synovial samples from patients with Lyme arthritis. J. Clin. Microbiol. 38 (2000), 1895–1900. [29] J.D. Luneman, Zarmas S, Priem S, Franz J, Zschenderlein R, Aberer E, Klein R, Schouls L, Burmester GR, Krause A, Rapid typing of Borrelia burgdorferi sensu lato species in specimens from patients with different manifestation of Lyme borreliosis. J. Clin. Microbiol. 39 (2001), 1130–1133. [30] I. Demaerschalk, Messaoud AB, De Kesel M, Hoyois B, Lobet Y, Hoet P, Bigaignon G, Bollen A, Godfroid E, Simultaneous presence of different Borrelia burgdorferi genospecies in biological fluids of Lyme disease patients. J. Clin. Microbiol. 33 (1995), 602–608 [31] A.M. Lebech, Hansen K, Rutledge Bj, Kolbert CP, Rys PN, Persing DH, Diagnostic detection of Borrelia burgdorferi DNA by PCR in cerebrospinal fluid in Lyme neuroborreliosis and direct genotyping of the causative agent. Molecular Diagnosis 3 (1998), 131–141. [32] H. Eiffert, Ohlenbusch A, Christen Hj, Thomssen R, Spileman A, Matuschka FR, Nondifferentiation between Lyme disease spirochetes from vector ticks and human cerebrospinal fluid. J. Infect. Dis. 171 (1995), 476–479.

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Infection of Ixodid Ticks, Mosquitoes and Patients with Borrelia, Bartonella, Rickettsia, Anaplasma, Ehrlichia and Babesia in Western Siberia, Russia Olga MOROZOVA a,1 , Vera RAR a, Yana IGOLKINA a, Andrey DOBROTVORSKY b, Igor MOROZOV a and Felipe C. CABELLO c a Institute of Chemical Biology and Fundamental Medicine of Siberian Branch of the Russian Academy of Sciences, Novosibirsk 630090, Russia b Institute of Systematics and Ecology of Animals of Siberian Branch of the Russian Academy of Sciences, Novosibirsk 630091, Russia c Department of Microbiology and Immunology, New York Medical College, Valhalla, New York 10595, USA Abstract. Arthropod-borne bacteria can cause a variety of human infectious diseases and have a high diversity of geographic distributions, animal reservoirs, arthropod vectors, and pathogenic properties. Human-biting, questing adult Ixodes persulcatus and Dermacentor reticulatus ticks, as well as mosquitoes and midges from Western Siberia, Russia, were tested for infection with Borrelia, Bartonella, Rickettsia, Anaplasma/Ehrlichia, and Babesia using nested PCR assays with subsequent sequencing. I. persulcatus ticks were found to be infected with Borrelia spp. (39.5 ± 4.5%), Bartonella spp. (37.6 ± 4.3%), Rickettsia tarasevichiae (90.0 ± 4.8%), Anaplasma phagocytophilum (2.4 ± 1.4%), and Ehrlichia muris (8.8 ± 2.5%), whereas D. reticulatus ticks contained DNA of Borrelia spp. (3.6 ± 2.0%), Bartonella spp. (21.4 ± 4.5%), R. tarasevichiae (3.2 ± 3.2%), Rickettsia sp. RpA4 (51.6 ± 9.1%), and Babesia canis canis (3.6 ± 2.0%). Borrelia garinii, B. afzelii, and their mixed infections were observed among I. persulcatus, whereas B. garinii and B. spielmanii DNA were present in samples from D. reticulatus. Surprisingly, only two human pathogens—Bartonella henselae and B. quintana—were found in ixodid ticks in Siberia, despite long-term sample collection and phylogenetic analysis of all known Bartonella species. Moreover, currently both B. henselae and B. quintana, but none of other tick-borne infectious agents studied, were found in mosquitos of the genus Aedes. Bartonella DNA was detected in 1.9 ± 2.1% Aedes cantans mosquitoes but not in samples isolated from mosquitoes of other species including Ae. punctor, Ae. cinereus, and Ae. communis, or in Byssodon maculata midges. Ba. canis canis was the only subspecies found in D. reticulatus, but no Babesia species were observed in I. persulcatus. Thus, both bacterial infection rates and loads for I. persulcatus 1 Corresponding author: Olga Morozova, Institute of Chemical Biology and Fundamental Medicine of Siberian Branch of the Russian Academy of Sciences, Lavrentyev’s Avenue 8, 630090 Novosibirsk, Russia. Tel: +7(383)336 09 89. Fax: +7(383)333 36 77. E-mail: [email protected].

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exceeded those for D. reticulatus and Aedes spp. Considering the large number of ticks implicated in the transmission of bacteria, human exposure to these infectious agents may be more substantial than is currently believed. From 369 patients with clinical manifestations of generalized infection following tick bites, Borrelia DNA was found in 43 plasma samples, Bartonella DNA in 73 blood cell samples, and R. sibirica DNA in a single sample. Molecular typing indicated the prevalence of infection with B. garinii and B. quintana. Neither Ehrlichia/Anaplasma nor Babesia was found in human specimens in the Novosibirsk region. Keywords: tick-borne bacteria, protozoa, arthropod vectors, human blood, Asian part of Russia

Introduction Ticks are important vectors of bacteria, viruses, and protozoa worldwide [1]. In Western Siberia, Russia, prevalent tick species are taiga ticks Ixodes persulcatus Schulze and meadow ticks Dermacentor reticulatus (Acarina: Ixodidae) [1, 2, 3]. The life cycle of the ixodid ticks lasts from 2 to 5 years and includes the following stages: imago-egg-larvae-nymph-imago. Each stage (except for the egg stage) is dependent on the tick’s taking a single blood meal from a vertebrate host. Ticks parasitize many ground-foraging bird species and virtually all terrestrial mammals [3]. Besides, both tick species are capable of biting humans, thus transmitting different tick-borne infections [2, 3]. Persistence of tick-borne pathogens in natural populations might include transstadial transmission in arthropod vectors and vertical transmission in warm-blooded reservoir hosts, horizontal nonviremic transmission (NVT) between cofeeding ticks, sexual transmission between ticks and between vertebrates, and transfer of the infectious agents between ticks and mammals in both directions. Ticks can serve as both natural reservoirs and vectors for tick-borne viruses and rickettsia due to their effective transstadial and transovarial transmission [4]. However, for other intracellular arthropod-borne bacteria (Bartonella, Anaplasma/Ehrlichia), extracellular spirochetes (Borrelia), and intraerythrocytic protozoa (Babesia), the existence of transovarial transmission remains unknown, although transstadial transmission has been shown to occur [2, 4, 5]. The circulation of the infectious agents between arthropod vectors and warm-blooded reservoir hosts therefore seems to be necessary for their survival. Among tick-borne bacteria, extracellular spirochetes of the genus Borrelia are widespread and most studied. Some of these belonging to the Borrelia burgdorferi sensu lato (s.l.) complex, are causative agents of Lyme borreliosis [1, 2]. B. burgdorferi was originally characterized as a single species. However, later it has become clear that the complex is composed of a number of distinct species and genomic groups [6–8]. Recently, a new species, B. spielmanii (syn. A14S, I-77), has been described [6]. Other known pathogenic bacteria transmitted by ticks belong to the intracellular alpha Proteobacteria including the three families Bartonellaceae, Rickettsiaceae, and Anaplasmataceae [9]. Bacteria of the genus Bartonella infect erythrocytes and endothelial cells. Different species of Bartonella are the etiological agents of cat scratch disease, trench fever, and Carrion’s disease [5]. Rickettsia penetrate into endothelial cells from blood after arthropod bites and can also infect macrophages, hepatocytes and muscle cells. Infection of endothelial cells results in a higher level of permeability and clinical symptoms such as fever, headache, and muscle pain, followed by the appearance of rashes. The rickettsial diseases of man differ depending on the arthropod vector (fly-borne, louse-borne, flea-borne, mite- and tick-borne) and include

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North Asian tick typhus caused by Rickettsia sibirica, Rocky Mountain spotted fever (R. rickettsii), fièvre boutonneuse (R. conorii), Queensland tick typhus (R. australis), and spotted fever (R. rhipicephali) [4]. Members of the genera Ehrlichia and Anaplasma, from the family Anaplasmataceae, mainly infect monocytes and granulocytes, causing human and animal ehrlichioses and anaplasmoses, respectively [10]. The tick-borne protozoa of the genus Babesia reproduce in erythrocytes, resulting in babesiosis among humans and wild and domestic animals [11]. Until 1987 the taiga ticks were thought to be able to transmit only tick-borne encephalitis virus (TBEV), but extensive studies have shown their competence for transmission of pathogenic spirochetes (B. garinii and B. afzelii) [1, 2], and Ehrlichia/Anaplasma [1, 12, 13]. Infection of ixodid ticks with Bartonella spp. has recently been described in the USA [14], in Europe [15, 16] and in Western Siberia [17]. Infection of I. persulcatus with Babesia microti pathogenic for immunocompromised humans has been shown by PCR with genus- and speciesspecific primers [18], but nucleotide sequences of the specific PCR products remain unknown. The second tick species—D. reticulatus—inhabits meadows and pastures [2], as well as suburban areas from Europe to central Asia, but not taiga and dry steppes. D. reticulatus is well known as the vector of a canine pathogen B. canis canis; Rickettsia spp., Francisella tularensis, and Coxiella burnetii were also found in this tick species [1]. Borrelia spp. was detected in different Dermacentor species including D. reticulatus by means of PCR [19] and an indirect immunofluorescence assay [20]. Infection of D. reticulatus with Bartonella and Anaplasma/Ehrlichia species was previously unknown in spite of the detection of the bacterial DNA in other species of Dermacentor [14, 21]. Despite considerable epidemiological significance, little or nothing is known about the prevalence and the genetic variability of the tick-borne pathogens in ixodid ticks as well as their pathogenicity for humans and animals in Western Siberia, Russia [2].

1. Materials and Methods Unfed adult I. persulcatus ticks were collected by flagging of lower vegetation in different suburban places of mixed aspen-birch and pine forests of the Novosibirsk region (55qN, 83qE) and the Tomsk region (56°N, 85°E) in epidemiological seasons from May to August 2001–2005. Questing imagos of D. reticulatus ticks were collected by flagging in different locations of river valleys and forest-steppe zones of the Novosibirsk and Omsk regions (55qN, 73qE) during the spring–summer periods of 2003–2005. Questing adult mosquitoes and midges searching for blood donors were caught in mixed forests near the Ob River in the summer 2004 and 2005. Arthropod species were identified on the basis of morphological traits. Total nucleic acids were isolated from 181 individual I. persulcatus, 87 D. reticulatus ticks, 24 pools of mosquitoes, and 17 pools of midges (10 individuals in each), as well as from 369 blood samples of patients who had infectious symptoms after tick bites. DNA was prepared by lysis in guanidine isothiocyanate followed by phenol-chloroform deproteinization and isopropanol precipitation as described [22]. For detection of tick-borne infectious agents, nested PCR with specific primers was done as previously described [23]. For control of the suitability of DNA for PCR [23], Rickettsia-specific DNA was detected using primers specific for the citrate synthase (gltA) gene (outer primers: TATAGACGGTGATAAAGGAATC and

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TGCATTTCTTTCCATTGTGC; inner primers: ATGGCTATTATGCTTGCGGC and CAGAACTACCGATTTCTTTAAGC). Nucleotide sequences of the PCR products were determined using the BigDye Terminator Cycle Sequencing Kit and the ABI PRISMTM 310 or 3100-Avant Genetic Analyzers (Applied Biosystems, USA) at the DNA Sequencing Centre of the Siberian Branch of the Russian Academy of Sciences, Novosibirsk, Russia. Nucleotide sequences of PCR products determined in this study were analyzed by BLASTN and aligned using CLUSTALW [24] or DNA Star (DNASTAR Inc) software. Phylogenetic analysis was performed using MEGA 3.1 software [25]. UPMGA and neighbor-joining algorithms using the Kimura 2-parameter model for generation of the distance matrix as well as maximum parsimony and minimal evolution using a heuristic search. Bootstrap analysis was performed with 1000 replications. Nucleotide sequences determined in this study were deposited in GenBank under the following accession numbers. Borrelia: AF538265–AF538271, AY429014– AY429016, AY540051–AY540052, AY583237–AY583238, AY741543–AY741544 AY862885–AY862890, AY603350–AY603351. Bartonella: AY453166–AY453170; AY597524, AY612093, AY612094, AY885134–AY885146. Rickettsia: DQ124930, DQ127824, DQ131912, DQ176434, DQ191466. Anaplasma phagocytophilum: AY587607. Ehrlichia muris: AY587608. Babesia: AY527063 and AY649326.

2. Results and Discussion Arthropod vectors (ixodid ticks, mosquitoes and midges) and patients who had been bitten by ticks in Western Siberia, Russia were studied by nested PCR using genusspecific primers for five bacterial and one protozoan tick-borne pathogens. In order to control the suitability of DNA for PCR analysis, an 18S rRNA gene fragment was amplified. Negative samples were excluded from further analysis. Both tick species contained Borrelia, Bartonella, and Rickettsia DNA, whereas Ehrlichia or Anaplasma DNA were detected only in I. persulcatus and Babesia DNA— only in D. reticulatus ticks (Table 1). Table 1. Prevalence of tick-borne infectious agents in ixodid ticks in Western Siberia, Russia. Prevalence (%) Tick-borne pathogens Ixodes persulcatus

Dermacentor reticulatus

Borrelia spp.

39.5 ± 4.5

3.6 ± 2.0

Bartonella spp.

37.6 ± 4.3

21.4 ± 4.5

Rickettsia spp.

90.0 ± 4.8

54.8 ± 9.0

Ehrlichia spp.

8.8 ± 2.5

0

Anaplasma spp.

2.4 ± 1.4

0

0

3.6 ± 2.0

Babesia spp.

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57

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B. garinii NT29 (L30130) Japan HU (AY741544) Novosivirsk (AF538266) Novosibirsk 63 Ip 5 HB 35 (AY862886) Novosibirsk Ip strain P-2 (AY583238) Omsk 74 Ip 34 (AF538267) Novosibirsk Ip strain P-12 (AY583237) Omsk 48 5 Ip 29 (AY862886) Novosibirsk 2004 Ip 68 (AF538270) Novosibirsk 2002 6 HB 220 (AY862889) Novosibirsk 2004 52 Ip 15 (AY862887) Novosibirsk 2004 55 Ip 57 (AF538268) Novosibirsk 2002 44 B. garinii 20047 (L30119) France 4 Ip 59 (AF538269) Novosibirsk 2002 HB 82 (AY862888) Novosibirsk 2004 B. garinii ChY13p (AB003785) China Ip 3 (AF538265) Novosibirsk 2002 HB 2003 (AY741543) Novosibirsk

62 41

41

B. afzelii VS461 (AY032913) Taiwan Ip 63 (AF538271) Novosibirsk 2002 7 HB 117 (AY862890) Novosibirsk 2004 B. burgdorferi s.s. 297 (AJ006507) B. burgdorferi s.s. B31 (L30127) 68 B. burgdorferi s.s. 212 (L30121) B. valaisiana CKA1 (AB022127) China B. tanukii NPN1 (AB100436) Nepal B. spielmanii2102 (DQ286234) France B. spielmaniiC (AF497994) CzechRepublic 10 Dr 35 (AY540052) Omsk Dr 22 (AY540051) Omsk B. sinica TIIo (AB100438) Thailand B. sinica NNIo (AB100435) Nepal B. sinica CWO1 (AB022130) China

52 57 99

58

97 81

Figure 1. Phylogenetic tree based on Borrelia 5S-23S rRNA intergenic spacer sequences. The tree was constructed by the UPMGA method. B. sinica was used as outgroup. Numbers above the branch indicate bootstrap support. Abbreviations: HU, human urine; HB, human blood; Ip, I. persulcatus; Dr, D. reticulatus. Reference species are in bold. GenBank accession numbers are in parentheses.

In 39.5 ± 4.5% of samples isolated from I. persulcatus and in 3.6 ± 2.0% of samples from D. reticulatus, Borrelia DNA was found (Table 1). The nucleotide sequences of the PCR products corresponding to 5S–23S intergenic spacer determined in this study were compared to those of other Borrelia species (Figure 1). The sequences derived from ticks appeared to be distributed among three clades corresponding to B. garinii, B. afzelii, and B. spielmanii. However, no B. spielmanii DNA was detected in 43 PCR-positive blood samples from 369 patients examined

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(Figure 1). Although B. garinii and B. afzelii were separated, weak statistical support resulted from the known polymorphism of the non-coding region of the internal transcribed spacer (ITS) was shown (Figure 1). B. spielmanii and B. burgdorferi sensu stricto clades had excellent statistical support (bootstrap indexes of 100 and 99, respectively) (Figure 1). Data obtained revealed a very high prevalence of B. garinii both in vectors and patients following tick bites in the Novosibirsk region during a 5year period of observation. Although the presence of B. burgdorferi sensu stricto in Novosibirsk region was reported previously (Olga N. Grishaeva, “Vector-Best,” Novosibirsk, Russia, personal communication), the phylogenic analysis revealed no B. burgdorferi sensu stricto either in patient specimens or in ticks analyzed in our study. B. myomotoi DNA was also not found in patient specimens and ticks despite the previous discovery of this Borrelia species in Novosibirsk (Ludmila S. Karan, Central Research Institute of Epidemiology, Moscow, Russia, personal communication). Borrelia-specific DNA was detected in blood plasma and urine but not in synovial fluid from patients following tick bites. Some of the patients denied having had contact with a tick but confirmed their presence in the regions endemic to ticks during epidemiological seasons. B. quintana (NC005955) HB 055q (AY885142) HB 032 (AY885136) HB 4

(AY612093)

HB 044 (AY885138) 62

HB 139 (AY885134) HB 248 (AY885146) HB 045 (AY885139)

100

Ip 32 HB 4-2

(AY612094)

HB 179 (AY885143) HB 039 (AY885137) HB 223 (AY885145) 62 Ip 39

HB 207 (AY885144) HB 055h (AY885141) HB 048

99

(AY885140)

B. henselae (NC005956) 70

Ip 5

(AY453170)

HB 3-17 (AY597524) 59 Ip 3

(AY453168)

Ip 4

(AY453169)

HB 013

(AY885135)

B. bacilliformis (Z15160)

Figure 2. Phylogenetic tree based on Bartonella groEL gene fragment sequences. The tree was constructed by the UPMGA method. Bartonella bacilliformis was used as the outgroup. Abbreviations: HB, human blood; Ip, I. persulcatus. Reference species are in bold. GenBank accession numbers are in parenthesis.

O. Morozova et al. / Infection of Ixodid Ticks, Mosquitoes and Patients 90 30 24 52

227

Dm 23 (DQ176434) Ural R. slovaca (U59725) R. sibirica (U59734) Cr (DQ124930) HB-2 (DQ127824)

76

Rickettsia sp. DnS28 (AF120027) 99

75 64

Rickettsia sp. RpA4 (AF120029) Rickettsia sp. DnS14 (AF120028) R. rickettsii (U59729) Ip 4 (DQ131912)

100

R. helvetica (U59723) R. canadensis (U59713) Dr 18 (DQ191466)

100 100

R. tarasevichiae (AF503167)

Figure 3. Phylogenetic tree based on Rickettsia citrate synthase (gltA) gene fragment sequences. The overlapping bootstrap indexes in the upper part of the tree correspond to different cladistic groups: 1) group 1 (Dm23, R. slovaca, R. sibirica and Cr) with bootstrap index 24; and 2) group 2 = group 1 + HB-2 with bootstrap index 52. Abbreviations: Cr, Clethrionomys rutilus; HB, human blood; Ip, I. persulcatus; Dr, D. reticulatus; Dm, D. marginatus. Referenced species are in bold. GenBank accession numbers are in parentheses.

Bartonella DNA was detected by nested PCR with primers specific to the groEL gene in both I. persulcatus (37.6 ± 4.3) and D. reticulatus (21.4 ± 4.5) ticks (Table 1), as well as in mosquitoes and patients. Bartonella DNA was detected in 5 of 24 pools of mosquitoes that corresponded to the individual infection rate of 1.9 ± 2.1%. Aedes cantans, but not mosquitoes of other species including Ae. punctor, Ae. cinereus, and Ae. communis or midges Byssodon maculata, were shown to contain Bartonella DNA. PCR assay revealed Bartonella-specific DNA in 73 of 369 patient blood samples (19.8 ± 2.1%). Comparative analysis of the groEL gene fragment nucleotide sequences 190 bp long revealed two evidently separated clades with excellent bootstrap indexes (99 and 100, respectively) (Figure 2) corresponding to B. henselae and B. quintana. Only two human pathogens, B. henselae and B. quintana, were found in arthropod vectors and in blood cells from patients in Western Siberia during a 5-year period of observation despite comparative phylogenetic analysis with all known Bartonella species [23, 26]. Mixed infection with two different Bartonella species was found in one patient sample. It might have been due to the Bartonella high infection rate in Western Siberia. Rickettsia DNA was detected in the majority of ixodid ticks studied (Table 1). Phylogenetic analysis (Figure 3) showed the prevalence of Rickettsia with unknown pathogenicity: R. tarasevichiae in 90.0 ± 4.8% of I. persulcatus and in 3.2 ± 3.2% of D. reticulatus, Rickettsia spp. RpA4 in 51.6 ± 9.1%% of D. reticulatus. Pathogenic R. slovaca causing tick-borne lymphoadenopathy (TIBOLA) was not detected in ixodid ticks in the Novosibirsk region but only in a single sample Dm-23 from D. marginatus from the Urals (the Chelyabinsk region) (DQ176434). DNA of R. sibirica, the etiological agent of North Asian tick typhus, was not detected in any arthropod vector

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studied. As for DNA of other pathogenic species, R. helvetica was found in a single I. persulcatus tick (DQ131912) also from the Urals (the Chelyabinsk region) but not from any ticks from the Novosibirsk region. However, R. sibirica DNA was found in a single blood sample from a patient in the Novosibirsk region, as well as in blood and liver samples from warm-blooded reservoir hosts (small rodents and insectivores) from the Urals (data not shown) [27]. Consequently, in spite of the circulation of the pathogenic Rickettsia spp. in natural populations of the Western Siberia, their transmission cycles remain unclear, and the infection rate among patients was extremely low. Further studies will be necessary to reveal the pathogenic potential of other widespread Rickettsia, R. tarasevichiae, Rickettsia sp. RpA4, Rickettsia sp. DnS14, and Rickettsia sp. DnS28.

70 99 87

A.phagocytophilum (AF470701) A.phagocytophilum (U02521) Ip 4 (AY587607)

96

A.platys (M82801)

100

A.bovis (U03775) A.marginale (M60313) 100

E.muris (U15527) Ip 16 (AY587608) E.ruminantium (AF069758)

100

E.canis (M73221)

80 73 54

E.ewingii (M73227) E.chaffeensis (U60476) Wolbachia pipientis (AF179630)

Figure 4. Phylogenetic tree based on Anaplasma/Ehrlichia 16S rRNA gene fragment sequences. Abbreviations: Ip, I. persulcatus. Reference species are in bold. GenBank accession numbers are in parentheses. Wolbachia pipientis was used as the outgroup.

Ehrlichia and Anaplasma DNA were found in I. persulcatus but not in D. reticulatus ticks using nested PCR (Table 1). The nucleotide sequences of the A. phagocytophilum 16S rRNA gene fragment (629 bp) were identical to each other (GenBank accession number AY587607) and to the known A. phagocytophilum sequence (AF205140). Nucleotide sequences of E. muris were also identical to each other (GenBank accession number AY587608) and differed from the known E. muris DNA sequence (U15527) at only one position - 91 (CoT). In the phylogenetic tree, both A. phagocytophilum and E. muris sequences clearly formed distinctive clusters (Figure 4). Neither Ehrlichia, nor Anaplasma DNA were found in blood cells of patients following tick bites. Babesia DNA was found only in D. reticulatus ticks (Table 1). It was not detected in I. persulcatus or in mosquitoes, midges, and patient blood samples. All the nucleotide sequences of the Babesia 18S rRNA 1203 bp gene fragment determined in this study and previously [28] were similar to the known B. canis canis nucleotide

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sequence (GenBank AY072926) with three variable positions. In the phylogenetic tree B. canis canis sequences formed a distinctive cluster separated from other B. canis subspecies with excellent bootstrap support (Figure 5). B. canis canis is known to be transmitted by D. reticulatus in Europe and often causes severe disease characterized by fever, jaundice, splenomegaly, thrompocytopenia, hemolytic anemia, and haemoglobinemia among dogs. Molecular characterization of B. canis canis in canine blood is described elsewhere [28].

75 Dr 2 (AY649326) 100 99 96 100

B. canis canis (AY072926) Dr 5 (AY527063) B. canis vogeli (AY072925) B. canis rossi (L19079) B. divergens (U07885)

97

B. gibsoni (AB118032) B. microti (U09833) Plasmodium falciparum (M19172)

Figure 5. Phylogenetic tree based on Babesia 18S rRNA gene fragment sequences. Branch Dr2 has bootstrap support of 75. Abbreviations: Dr, D. reticulatus. Reference species are in bold. GenBank accession numbers are in parentheses. Plasmodium falciparum was used as the outgroup.

Phylogenetic trees constructed using UPGMA (Figures 1–5), neighbor-joining, maximal parsimony, and minimal evolution approaches (trees not shown) gave generally similar topologies.

3. Conclusions Detection of the tick-borne bacteria in unfed tick imagos proved the existence of transstadial transmission for five bacterial and one protozoan infectious agents. The high rate of Borrelia and Bartonella infection among unfed adult I. persulcatus suggested that immature stages of the ticks were naturally infected as the result of effective transstadial transmission or feeding on infected vertebrate hosts. The theoretically expected and experimentally observed values of the mixed infections of the ticks with Borrelia, Ehrlichia, and Bartonella and of patients with Borrelia and Bartonella were statistically similar and consistent with independent distribution of these pathogens as previously reported [13]. Thus, simultaneous coinfection with Borrelia, Ehrlichia/Anaplasma, and Bartonella found in 2.9% of I. persulcatus ticks even slightly exceeded the statistical probability of 1.8%. However, Ehrlichia/Anaplasma DNA, but no Babesia species, was detected in I. persulatus. And conversely, D. reticulatus appeared to be a vector for Babesia but not for Ehrlichia or

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Anaplasma. Therefore, the reciprocal inhibition between Babesia and Ehrlichia/Anaplasma or host-pathogen relationships cannot be excluded. Currently, pathogenic properties and clinical manifestations of tick-borne infections cannot always be assigned to a certain infectious agent since they depend on the general status of the patient and the possibility of mixed infection with known or yet unknown pathogens.

References [1] [2] [3] [4] [5] [6] [7] [8]

[9]

[10] [11] [12] [13]

[14] [15] [16] [17]

Parola, P. and Raoult, D. Ticks and tickborne bacterial diseases in humans: an emerging infectious threat. Clin Infect Dis 2001; 32:897–928. Korenberg, E.I., Kovalevsky, Y.V., Gorelova, N.B. Ecology of Borrelia burgorferi sensu lato in Russia. In: Gray, G, Kahl, O, Lane, RS, eds. Lyme Borreliosis Epidemiology and Control. Oxford: CAB International; 2002:175–200. Filippova, N.A. The taiga tick Ixodes persulcatus (Acarina, Ixodidae). Morphology, systematics, ecology, medical importance. Leningrad, Nauka, 1985. Balashov, Yu.S. and Daiter, A.B. Hematophage arthropods and rickettsia.Leningrad, Nauka, 1973. Jacomo, V., Kelly, P.J., Raoult, D. Natural history of Bartonella infections (an exception to Koch’s postulate) // Clin. Diagn. Lab. Immunol. 2002. V. 9. pp. 8–18. Richter, D., Schlee, D.B., Allgower, R., Matuschka, F.R. Relationships of a novel Lyme disease spirochete, Borrelia spielmanii sp. nov., with its hosts in Central Europe. Appl Environ Microbiol. 2004; 70:6414–6419. Ruzic-Sabljic, E., Arnez, M., Lotric-Furlan, S., Maraspin, V., Cimperman, J., Strle, F. Genotypic and phenotypic characterisation of Borrelia burgdorferi sensu lato strains isolated from human blood. J Med Microbiol. 2001; 50:896-901. Baranton, G. and Postic, D. Multi-locus sequence analysis (MLSA) as an alternative to whole DNA/DNA hybridization (WDDH) in Borrelia burgdorferi sensu lato taxonomy. NATO Advanced Research Workshop on the Molecular Biology of Spirochetes, 5–8 December 2005, Prague, Czech Republic (in press). Dumler, J.S., Barbet, A.F., Bekker, C.P., Dasch, G.A., Palmer, G.H., Ray, S.C. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: unification of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia, descriptions of six new species combinations and designation of Ehrlichia equi and 'HGE agent' as subjective synonyms of Ehrlichia phagocytophilum. Int J Syst Evol Microbiol 2001; 51:2145–2165. Chen, S.M., Dumler, J.S., Bakken, J.S., Walker, D.H. Identification of a granulocytotropic Ehrlichia species as the etiologic agent of human disease. J Clin Microbiol 1994; 32:589–95. Homer, M.J., Aguilar-Delfin, I., Telford, S.R., Krause, P.J., Persing, D.H. Babesiosis. Clin Microbiol Rev 2000; 13:451–469. Alekseev, A.N., Dubinina, H.V., Van De Pol, I., Schouls, L.M. Identification of Ehrlichia spp. and Borrelia burgdorferi in Ixodes ticks in the Baltic regions of Russia. J Clin Microbiol 2001; 39:2237– 2242. Morozova, O.V., Dobrotvorsky, A.K., Livanova, N.N., Tkachev, S.E., Bakhvalova, V.N., Beklemishev, A.B., Cabello, F.C. PCR Detection of Borrelia burgdorferi sensu lato, tick-borne encephalitis virus and human granulocytic erlichiosis agent in Ixodes persulcatus ticks from Western Siberia, Russia. J Clin Microbiol 2002; 40:3802–3804. Chang, C.C., Chomel, B.B., Kasten, R.W., Romano, V., Tietze, N. Molecular evidence of Bartonella spp. in questing adult Ixodes pacificus ticks in California. J Clin Microbiol 2001;39: 1121–1126. Schouls, L.M., Van De Pol, I., Rijpkema, S.G., Schot, C.S. Detection and identification of Ehrlichia, Borrelia burgdorferi sensu lato, and Bartonella species in Dutch Ixodes ricinus ticks. J Clin Microbiol 1999;37: 2215–2222. Sanogo, Y.O., Zeaiter, Z., Caruso, G., Merola, F., Shpynov, S., Brouqui, P. Bartonella henselae in Ixodes ricinus ticks (Acari: Ixodida) removed from humans, Belluno province, Italy. Emerg Infect Dis 2003;9:329-332. Morozova, O.V., Cabello, F.C., Dobrotvorsky, A.K. Semi-nested PCR detection of Bartonella henselae in Ixodes persulcatus ticks from Western Siberia, Russia. Vector Borne Zoonotic Dis. 2004; 4:306–309.

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[18] Alekseev, A.N., Semenov, A.V., Dubinina, H.V. Evidence of Babesia microti infection in multiinfected Ixodes persulcatus ticks in Russia. Exp Appl Acarol 2003; 29:345–353. [19] Hubbard, M.J., Baker, A.S., Cann, K.J. Distribution of Borrelia burgdorferi s.l. spirochaete DNA in British ticks (Argasidae and Ixodidae) since the 19th century, assessed by PCR. Med Vet Entomol 1998; 12:89–97. [20] Kahl, O., Janetzki, C., Gray, J.S., Stein, J., Bauch, R.J. Tick infection rates with Borrelia: Ixodes ricinus versus Haemaphysalis concinna and Dermacentor reticulatus in two locations in eastern Germany. Med Vet Entomol 1992; 6:363–366. [21] Holden, K., Boothby, J., Anand, S., Massung, R. Detection of Borrelia burgdorferi, Ehrlichia chaffeensis, and Anaplasma phagocytophilum in ticks (Acari: Ixodidae) from a coastal region of California. J Med Entomol 2003; 40:534–539. [22] Chomczynski, P., Sacchi, N. Single-step method of RNA isolation by acid guanidinium thiocyanatephenol-chloroform extraction. Anal Biochem. 1987; 162:156–159. [23] Rar, V.A., Fomenko, N.V., Dobrotvorsky, A.K., Livanova, N.N., Rudakova, S.A., Fedorov, E.G., Astanin, V.B., Morozova O.V. Tickborne Pathogen Detection, Western Siberia, Russia. Emerg Infect Dis. 2005; 11:1708–1715. [24] Thompson, J.D., Higgins, D.G., Gibson, T.J. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 1994;22: 4673–4680. [25] Kumar, S., Tamura, K., Masatoshi, N. MEGA3: Integrated software for Molecular Evolutionary Genetics Analysis and sequence alignment. Brief Bioinform. 2004; 5:150–163. [26] Fomenko, N.V., Rar, V.A., Romanova, E.V., Chernousova N.Ya., Gosteeva L.A., Fedorov E.G., Morozova O.V. Molecular-genetic analysis of the tick-borne infection in patients in Novosibirsk region. Molecular Medicine 2005, 4:48–52 (in Russian). [27] Igolkina, Ya.P., Fomenko, N.V., Livanova, N.N., Astanin, V.B., Gosteeva, L.A., Chernousova, N.Ya., Rar, V.A. Detection of different Rickettsia species from ixodid ticks, human blood, and blood of small mammals in the Southwestern Siberia and the Urals. Bulletin of Siberian Medicine 2005, ʋ1, P. 121125 (in Russian). [28] Rar, V.A., Maksimova, T.G., Zakharenko, L.P., Bolyakhina, S.A., Dobrotvorsky, A.K., Morozova O.V. Babesia DNA detection in canine blood and Dermacentor reticulatus ticks in Southwestern Siberia, Russia. Vector Borne Zoonotic Dis. 2005; 5:285–287.

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Part 3 Gene Expression

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Genetic Studies of the Borrelia burgdorferi bmp Gene Family Felipe C. CABELLO a,1 , Lidiya DUBYTSKA a, Anton V. BRYKSIN a, Julia V. BUGRYSHEVA a and Henry P. GODFREY b a Department of Microbiology and Immunology, New York Medical College, Valhalla, NY USA b Department of Pathology, New York Medical College, Valhalla, NY 10595 USA Abstract. The development of genetic systems for B. burgdorferi has allowed identification of several regulatory and structural genes involved in the ability of this bacterium to colonize and disseminate in its arthropod and mammalian hosts. Progress has been slow as a result of the complex biology of this pathogen, and its genetic distance from Gram-positive and Gram-negative bacteria has impeded easy adaptation to B. burgdorferi of genetic tools developed for other bacteria. We have repeatedly encountered these issues in our attempts to use molecular genetic tools to analyze the role of the members of the paralogous bmp gene family in the biology and virulence of B. burgdorferi. For example, an attempt to characterize the role of bmpC in virulence using a bmpC deletion mutant of an infectious B. burgdorferi 297 strain and an extrachromosomally complemented derivative of this mutant was only partially successful. While deletion of bmpC did not decrease infectivity in mice, it significantly decreased B. burgdorferi DNA in joints and arthritis. Extrachromosomal complementation of this null mutation reduced infectivity and did not restore tissue pathogenicity. To overcome these problems, we have adapted the Tet system of controlled gene expression developed for other bacteria and eukaryotic cells to manipulate gene expression in B. burgdorferi. In a two plasmid Tet system, where one plasmid contains the TetR repressor fused to the flaB promoter and the other plasmid contains a hybrid borrelial promoter with a tet operator (Ptetl-1), expression of the gene fused to the Ptetl-1 promoter (truncated BmpA, green fluorescent protein) is regulated by anhydrotetracycline in a concentration dependent manner over a wide range of concentrations. Because the two plasmid Tet system has some inherent instability manifested by escape from TetR repression and potential plasmid loss requiring continuous antibiotic selection, we constructed a B. burgdorferi strain containing the flaB/tetR fusion inserted in a nonessential gene (luxS) and designed a new hybrid promoter containing two instead of one tet operators (Ptetl-2). The functionality of this Tet system was demonstrated by manipulating expression of the B. burgdorferi bmpA gene. These findings strongly suggest that the Tet system can be used in B. burgdorferi to manipulate gene expression and open the door to manipulating gene expression by antisense RNA technology and gene fusions that will permit isolation of conditional lethal mutants in B. burgdorferi.

Keywords: Borrelia burgdorferi, gene regulation, Tet system, lipoprotein, bmpA, bmpC, tissue pathogenicity. 1 Correponding author: Felipe C. Cabello, Department of Microbiology and Immunology, New York Medical College, Valhalla, NY 10595-1690 USA; E-mail: [email protected]; Phone: +1 (914) 594-4182.

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Introduction The paradigm developed by Koch’s postulates has been key to framing the search for factors involved in the biology and pathogenesis of bacterial infections [1, 2]. These postulates in their molecular form of expression permit establishment of cause-effect relationships between genes, phenotypes, functions and pathological events in vitro and in vivo [1, 2]. Crucial to this approach is the development of genetic tools that facilitate isolation of bacterial strains differing only in the phenotype of interest. This permits relating the presence (or lack) of a specific trait to expression (or failure of expression) of particular virulence properties in these strains [1-3]. This genetic and functional approach has been used extensively and productively in studies of the virulence of Gram-negative and Gram-positive bacteria [1-4], and in recent years, has been applied with increasing success to the study of the biology and virulence of B. burgdorferi [5, 6]. Many studies in vitro and in vivo have thus been able to identify and describe the role played by genes and gene products in the interactions of B. burgdorferi with its tick and mammalian hosts. The widespread application of genetic tools developed for Gram-positive and Gram-negative bacteria to the study B. burgdorferi has however encountered several limitations [5, 6], and there is still a great need to develop alternative genetic tools for this pathogen and for spirochetes in general. There are a number of factors that have impeded the application of general bacterial genetic tools to the study of B. burgdorferi (Table 1).

Table 1. Physiologic and genetic properties of B. burgdorferi that limit its genetic manipulation. Properties Undefined nutritional requirements [5, 6]

Limitations Inability to isolate auxotrophic mutants and conditional lethal mutants

Slow growth [5, 6]

Long wait for experimental results

Small colonies on agar [5, 6]

Limited ability to use colonial morphology as indicator of putative genetic change

Genetic distance from other bacteria [6]

Difficult adaptation of genetic systems developed for other bacteria

Segmented genome [5, 7]

Complex interactions between plasmid and chromosomal genes

Unstable genome [6, 7]

Loss of plasmids relevant for infectiousness

Lack of mechanisms of horizontal gene transfer [5, 6]

Inefficient electroporation is the only mechanism of gene transfer

Lack of transposons and integrons [6, 8, 9]

Difficult adaptation of these genetic tools from other bacterial systems

Paucity of genetic markers [5, 6]

Few selection markers for recombinants in vitro and in vivo

Illegitimate recombination [5, 6]

Insertion of electroporated DNA in nonhomologous regions of the genome

Increased rate of mutation to several antibiotics [10, 11]

Interferes with selection of recombinants

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1. Limitations in the Genetic Manipulation of B. burgdorferi The approach used to study genes and gene products that may be relevant for pathogenesis in B. burgdorferi has been to study the virulence of wild type, null mutants of a specific gene and their complemented derivatives. This approach has permitted identification and analysis of of genes and the regulation of their expression in vitro and in vivo in ticks and mammals [5, 6, 12, 13]. However, it has also encountered limitations resulting from the compact genome of B. burgdorferi where genes with overlapping transcriptional and termination signals can lead to destruction of common transcripts and polar effects in mutants isolated by classical insertional inactivation and allelic exchange (Table 2) [5, 6]. These constraints have also been accompanied by the frequent inability to complement these null mutants fully with an extrachromosomally located wild-type gene, most likely as a result of inbalances in gene expression generated by fluctuating copy numbers of cloning vectors through the growth cycle of B. burgdorferi [14, 15]. The null mutant-complementation approach is also hindered in B. burgdorferi by the presence of paralogous gene families where the lack of synthesis of one gene product produced by the null mutation may potentially be replaced by the synthesis of a paralogous gene product with similar or identical function(s) [16]. Moreover, we have frequently observed the occurrence of illegitimate recombination events following electroporation between the introduced DNA containing antibiotic resistance determinants and DNA segments of the B. burgdorferi genome (chromosome and plasmids) [data not shown]. These processes of illegitimate recombination undermine and delay isolation of null mutants and their complementation.

Table 2. Limitations to the use of insertion and deletion null mutations and their complementation in genetic manipulation of B. burgdorferi. Compact genome and overlapping genes increases the possibility of polar effects Paralogous gene families of proteins whose several members with similar function undermine the goal of obtaining null mutants Illegitimate recombination yields insertion of electroporated DNA in non-homologous regions Inability to generate conditional lethal mutants Failure of extrachromosomal complementation because of plasmid loss and disregulation

2. Paralogous Gene Family 36, the bmp Genes The above problems have been encountered by others in their studies of plasmidlocated lipoprotein genes, and by ourselves in our studies of the bmp genes, a family of lipoproteins located at nt339192-396506 on the complementary strand of the B. burgdorferi chromosome (Figure 1) [17-19]. In B. burgdorferi sensu stricto, the bmp genes (bmpA, bmpB, bmpC, bmpD) show 52-62% nucleotide sequence homology [16]. They are conserved in all B. burgdorferi sensu lato [17-21] but are partially duplicated in some genospecies ([22]). Even though BmpA is highly immunogenic in patients with

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F.C. Cabello et al. / Genetic Studies of the Borrelia burgdorferi bmp Gene Family Table 3. Characteristics of Bmp proteins [16-19].

Amino acids

BmpA

BmpB

BmpC

BmpD

339

341

353

341

Homology to BmpA (%)

100

49

37

46

MW (kDa)

36.9

37.5

39.8

37.1

Pi

5.03

4.86

9.9

5.32

Lyme disease [23] and, as mentioned below, the level of expression of bmpA is modulated by the host humoral immune system in mice [24, 25], the functions of BmpA and the other bmp gene products and their role in borrelial biology and virulence are still unknown. BmpA is the index member of the Bmp protein family (PFam 02608), a family with members in Treponema and in 76 other archeal and bacterial species. The Bmp proteins are also members of the Med orthologous protein group (COG 1744), an uncharacterized ABC-type transport system [26]. The lipoproteins encoded by the bmp genes have an amino-terminal type II signal sequence, a lipobox, 37-49% amino acid sequence homology and similar molecular weights (Table 3) [17-19]. The pI of BmpC is markedly more basic than the pI of BmpA, BmpB, or BmpC (Table 3). The conservation and chromosomal location of the bmp genes suggests that their study could provide important insights into B. burgdorferi biology and pathogenicity, and that examination of their expression under various environmental conditions could provide important information of their function in the various habitats B. burgdorferi encounters in its tick and mammalian hosts.

bmpD bmpA

rpsG

rpsL

bmpB bmpC

Transcription

bmpA

bmpA

bmpBT

bmpB

bmpC

bmpD

rpsG

rpsL

bmpD

bmpA

bmpA

Figure 1. Organization and complex transcription pattern of bmp and neighboring genes on the complementary strand of the B. burgdorferi chromosome [27, 28].

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The bmp genes are composed of two transcriptional units with complex transcriptional regulatory patterns (Figure 1) [27, 28]. One transcriptional unt contains bmpA, bmpB and bmpC, with bmpC being markedly less strongly expressed than bmpA or bmpB [27, 29]. The second contains bmpD and the genes for ribosomal proteins S7 and S12. Both units have a complex transcriptional regulatory pattern with polycistronic and overlapping mRNAs that appear to be coordinately regulated [27, 28]. We and others have observed condition-dependent differential gene expression of various bmp genes [21, 24, 27, 29-31], but the mechanisms of this differential expression are unknown. For example, we found that bmpD expression was sharply down modulated and the proportion of bmpD-rpsL message up regulated during coculture with tick cells compared to expression during culture in BSK-H; levels of other bmp genes did not vary under these conditions [29]. Revel et al. reported that expression of bmpC and bmpD increased significantly when B. burgdorferi were shifted from culture at 23°C, pH 7.5 (tick-like conditions) to culture at 37°C, pH 6.8 (mammalian host-like conditions), but showed no differences in expression levels when they were shifted from culture at 37°C, pH 6.8, to culture in peritoneal chambers or culture at 23°C, pH 7.5, in peritoneal chambers [31]. As noted above, Liang and coworkers found expression of bmpA, but not of bmpB, bmpC, or bmpD, to be down modulated in vivo in mice compared to levels seen during in vitro culture after 3-5 weeks as a result of selection pressure exerted by the host immune system [24, 25]. We have recently reported that patients with Lyme disease produce specific antibodies to BmpA (P39), BmpB and BmpD, implying that these proteins are expressed in infected human beings and are recognized by their immune systems [32]. No specific antibodies to BmpC were demonstrable in these patients. Whether this was a result of BmpC containing only epitopes cross-reactive with other Bmp proteins, or was due to the low expression of BmpC or to differences in localization in B. burgdorferi was not clear

3. Pathogenicity of a B. burgdorferi bmpC Null Mutant The different levels of expression of BmpA and BmpC could suggest that these proteins play different roles in B. burgdorferi virulence and pathogenicity. To answer this question, B. burgdorferi null mutants of BmpA and BmpC would be useful. They would also be generally useful to examine Bmp function in vitro and in vivo in mice. Unfortunately, repeated attempts over several years using several different protocols to isolate a BmpA null mutant were not successful. In the case of bmpC, we were able to isolate a null mutant in an infectious strain of B. burgdorferi 297 and have begun to use it to analyze the role of bmpC in virulence and pathogenicity in vivo. The bmpC gene was inactivated in low-passage, infectious B. burgdorferi 297 strain using PCR-based fusion [33] to construct a DNA segment containing bmpC interrupted by an inserted kanamycin resistance gene, much as was done to inactivate B. burgdorferi rel [34]. This PCR construction product was electroporated into low passage infectious B. burgdorferi 297 and mutants were selected by growing in kanamycin-containing medium. The presence of the interrupted bmpC gene was detected by PCR from B. burgdorferi genomic DNA (data not shown). The genetic structure of the deletion in these kanamycin-resistant B. burgdorferi was confirmed by sequencing to ensure that homologous DNA exchange had taken place between the wild-type bmpC gene in the chromosome and the interrupted bmpC allele. To complement the ǻbmpC mutant, a plasmid containing the bmpC wild-type allele under

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its own promoter (pAB77) was constructed from the B. burgdorferi spectinomycinresistant cloning vector, pKFSS1 [35], electroporated into the bmpC null mutant and selected by growing in kanamycin-containing medium. A control electroporation used only the pKFSS1 vector. PCR analysis of the partial diploid B. burgdorferi derivative showed that it contained both the disrupted gene on the chromosome and the wild-type bmpC gene. RT-PCR analysis using appropriate primers to amplify an internal bmpC region of this complemented derivative and its control electroporated with pKFSS1 demonstrated this inner fragment only in wild-type B. burgdorferi and in B. burgdorferi ǻbmpC pAB77, but not in B. burgdorferi ǻbmpC or in B. burgdorferi ǻbmpC pKFSS1. Reactions performed without reverse transcriptase showed no amplicons and confirmed the lack of DNA contamination in total RNA samples. PCR with appropriate primers showed that the 'bmpC derivatives contained the same plasmid complement as the wild-type parent.

Table 4. Infectivity of B. burgdorferi 297 and 'bmpC derivatives in C3H/HeJ mice.

Strain

No. cultures positive/No. Examined Skin

Blood

Heart

Bb 297 wild-type

10/11

10/11

0/6

Bb 297 'bmpC

11/12

9/12

0/6

Bb 297 'bmpC pKFSS1

0/5

0/5

0/5

Bb 297 'bmpC pAB77

0/5

0/5

0/5

Figure 2. B. burgdorferi 297 'bmpC and its derivatives exhibit decreased tissue pathogenicity. ***, P<0.001.

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Growth of the bmpC null mutant in complete BSK-H medium at 34qC was identical to that of the wild-type (data not shown). In preliminary experiments, its infectivity in needle-inoculated C3H/HeJ mice [34] (Table 4) and its ability to induce anti-Bmp antibodies were only slightly less than those of wild-type (data not shown). In contrast, neither the bmpC complemented or the pKFSS1-transformed B. burgdorferi 'bmpC strains were infective despite their containing all the plasmids of the wild-type parent by PCR analysis. The reason for this lack of infectivity in the face of a complete complement of plasmids of the wild-type parental strain is unknown. Although the bmpC null mutant did not display significantly decreased infectivity in mice as judged by culture (Table 4), it caused significantly less arthritis (Figure 2) and grew significantly less well in the tibiotarsal joints of infected mice as judged by PCR amplification of the flaB gene [36] than wild-type B. burgdorferi (Figure 2). This decreased tissue pathogenicity was completely unexpected in the face of the similar infectivity of the bmpC null mutant and the wild-type B. burgdorferi. In the case of the complemented and the vector-transformed 'bmpC derivatives, the lack of arthritis (tissue pathogenicity) and growth in the joint is consistent with their lack of infectivity. Despite our inability to complement this bmpC null mutant, these preliminary results suggest that this bmpC null mutant was attenuated, and that bmpC plays a role in B. burgdorferi joint pathogenicity. Since the function of bmpC and its gene product is not known, it is difficult to explain what the mechanism of this role is.

4. Manipulation of Gene Expression in B. burgdorferi Using the Tet System Although we were able to isolate a bmpC null mutant and use it to examine B. burgdorferi virulence and pathogenicity, our continuing inability to isolate a null mutation of bmpA despite repeated attempts suggested that we would need to develop alternative methods to null mutagenesis and complementation for manipulating expression of bmp and other genes in this organism. In many bacterial species, isolation of conditional lethal mutants has been an important tool in acquiring new knowledge regarding gene function and gene expression [3, 4]. Conditional lethal mutants are strains that contain a mutation in an essential gene where the mutated phenotype is expressed only under non-permissive conditions [3, 4]. This permits isolation of mutations whose defective phenotype is only expressed at high or low temperature, in a genetic background lacking suppressors or in media lacking a defined nutrient, with the defect usually resulting in the absence of bacterial growth [3, 4]. As an example of their use, conditional lethal mutants were crucial to dissecting DNA replication and DNA repair in bacteria and in identifying the enzymatic steps involved in many metabolic pathways [37]. Unfortunately, isolation of conditional lethal mutants in B. burgdorferi has not previously been possible because its slow growth, its strict temperature needs, while the lack of detailed knowledge of its nutritional requirements (Table 1) has impeded use of nutritional markers and non-permissive temperatures that have been used isolate conditional mutants in other bacteria. Other genetic methods used to obtain information regarding bacterial biology and virulence include manipulation of gene expression by altering their temporal and spatial regulation at the levels of transcription and translation [3, 38]. These methods have been used for complementation of null mutants with variable amounts of the wild-type gene product [3, 6], partial ablation of

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expression of a gene product of interest [3, 6], and isolation of conditional lethal mutants of essential genes [3, 6]. The ability to manipulate gene expression at will in bacteria has permitted use of antisense RNA technology and the preparation of antisense RNA libraries to identify virulence genes and to produce conditional lethal mutants in vitro and in vivo [38]. The ability to make gene fusions to promoters whose expression can be manipulated and controlled also provides an alternative means to isolating these conditional lethal mutants in genes of interest [39-42]. In light of this information and our inability to isolate null mutants of bmpA despite numerous attempts over several years, and considering the dearth of genetic tools in Borrelia that allow manipulation of its gene expression, we decided to try to develop genetic systems for B. burgdorferi that would permit this type of analysis.

5. Anhydrotetracycline (ATc)-regulated Expression of Green Fluorescent Protein (GFP) and Truncated rBmpA in B. burgdorferi We began to explore the possibility of regulating gene expression in B. burgdorferi using the Tet system developed by Bujard and his collaborators [42-45]. This system has been extensively used to modulate gene expression in eukaryotic cells [42-45] and in several bacterial systems [40, 41, 46]. The Tet system is based in the construction of hybrid promoters containing the tet operator DNA sequence [42-45]. This sequence tightly binds TetR; the bound TetR molecules then block initiation of transcription by RNA polymerase. In the presence of tetracycline or its analogues, TetR preferentially binds to these inducers and transcription from the hybrid promoter containing the tet operators is triggered. An important feature of the system is that the affinity of binding of tetracycline or its analogues to TetR is 1000-fold greater than their binding to the ribosome. Inactivation of the repressor can thus be carried out at inducer concentrations which have no antibacterial effect and which can freely diffuse through bacterial membranes [40, 41].

tetO

PbmpA

…TCCCTATCAGTGATAGAGATTGATGAAGTTATTGATCTAGAAAATAAAATAATAAGTGGAGAAA... -35 -10

Figure 3. Structure of Ptetl-1.

As a first step to applying the TetR system to B. burgdorferi, we created a hybrid B. burgdorferi promoter (Ptetl-1) containing a tet operator 5' to the bmpA promoter (Figure 3). To test the functionality of Ptetl-1 to ATc induction, we fused it to the GFP gene or to a truncated bmpA gene lacking 119 nucleotides (including those coding for the lipidation signal) using PCR-based fusion [33]. These GFP and rBmpA fusions were cloned into pKFSS1 to generate plasmids pLD2 and pLD3, respectively. Simultaneously, we constructed a plasmid containing the TetR gene under the B. burgdorferi flaB promoter using the B. burgdorferi cloning vector pED1 [14] to generate plasmid pLD1. We electroporated pLD1 into non-infectious B. burgdorferi B31 and constructed derivatives of this strain containing either pLD2 or pLD3. These

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B. burgdorferi derivatives containing tetR and either gfp or rbmpA genes under the control of the Ptetl-1 promoter were then exposed to the inducer tetracycline analogue, ATc [42-45].

Table 5. Fluorometric analysis (mean ± SD) of GFP expression after induction by 1.5 Pg/ml of ATc. ATc

Fluorescence intensity 1

B. burgdorferi pLD1

-

0.05 ± 0.1

B. burgdorferi pLD1

+

0.50 ± 0.7

Strain

B. burgdorferi pLD1 pLD2

-

1.7 ± 2.5

B. burgdorferi pLD1 pLD2

+

111 ± 11

1

Mean ± SD.

Figure 4. FACS analysis of regulated production of GFP in B. burgdorferi LD1/LD2 in the absence (-) and presence (+) of 1.5 Pg/ml of ATc.

Expression of GFP was dramatically increased after exposure of B. burgdorferi containing plasmids pLD1/pLD2 to 1.5 Pg/ml of ATc (Figure 4). Fluorometric analysis (Table 5) showed that there was a marked and significant increase in expression of GFP in B. burgdorferi pLD1/pLD2 after exposure to ATc. Table 5 also shows that there was detectable expression of GFP in the uninduced borrelia carrying both plasmids, suggesting that Ptetl-1 is not totally repressed by TetR and that there was some escape transcription from this promoter even in the absence of ATc. Lysates of B. burgdorferi pLD1/pLD3 induced with ATc contained immunoreactive truncated rBmpA demonstrable by immunoblotting (data not shown). The increase in expression of rBmpA was linear between 0 to 1.5 Pg/ml of ATc (data not shown). The Tet system in B. burgdorferi thus reproduces the characteristic linearity of gene expression in

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response to increasing concentrations of ATc that has been previously described in eukaryotic cells and in Gram-positive and Gram-negative bacteria [40-46]. These experiments confirm that the Tet system can be used in B. burgdorferi to manipulate gene expression at the transcriptional level. They show that varying the concentration of ATc provides the possibility of generating a wide range of levels of gene expression in B. burgdorferi. While the system described above readily allowed us to probe the feasibility of using the Tet system in B. burgdorferi, it had some clear shortcomings. These included the use of two potentially unstable plasmids that would require two antibiotics for their selection. Fluctuating copy number of the plasmids throughout the B. burgdorferi growth cycle could create regulatory imbalances between ATc and the concentration of TetR protein, and influence its binding to the Ptetl-1 promoter, interfere with regulation, and generate escape from repression. Moreover, the presence of two foreign plasmids in B. burgdorferi may increase the chances for illegitimate recombination between them and the B. burgdorferi genome. We therefore constructed an improved Tet system to forestall some of these problems.

6. Construction of an Improved Tet System for B. burgdorferi To overcome the challenges to the functionality of the Tet system produced by the genetic instability of its plasmid components and the potential variation in their copy numbers, tetR under the control of the B. burgdorferi flaB promoter was inserted into the luxS gene of low passage infectious B. burgdorferi 297. Inactivation of luxS does not appear to interfere with any of the biological properties of B. burgdorferi that have been examined, including its infectiousness [47].

tetO

tetO

…TCCCTATCAGTGATAGAGATTGATCCCTATCAGTGATAGAGAAATAAAATAATAAGTGGAGAAA... -35

-10

Figure 5. Structure of Ptetl-2.

We chose to insert a construct into luxS rather than into an intergenic region because B. burgdorferi has many overlapping genes, promoters and potential regulatory regions, and an insertion into an intergenic region could result in alteration of normal transcriptional patterns [16, 27]. B. burgdorferi 297 containing the flaB-tetR fusion in the luxS gene constitutively produced TetR demonstrable by immunoblotting (data not shown). Since the hybrid Ptetl-1 promoter containing only one tet operator might result in escape of TetR repression (Table 5), we constructed a second hybrid B. burgdorferi promoter responsive to TetR containing two tet operators in tandem, Ptetl-2 (Figure 5). This improved TetR system was used in the studies described below.

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7. Overproduction of BmpA by ATc-controlled Regulation of Expression Suggests that BmpA is an Essential Gene Product for B. burgdorferi As mentioned above, we have been interested in analyzing the function of BmpA and other Bmp proteins in B. burgdorferi biology for some time [27, 48]. We have repeatedly not been able to isolate null mutants that did not express BmpA despite using several genetic methods. These results suggested that BmpA is an essential gene product of B. burgdorferi, and that mutants unable to produce this protein were lethal. This hypothesis is also supported by our ability to produce null mutants of BmpC (see above) and to modulate expression of BmpD [Bryksin, Godfrey and Cabello, unpublished]. We therefore explored manipulating BmpA expression with the improved Tet system. For these experiments, we constructed a pKFSS1 derivative, pLD10, containing a copy of the native bmpA gene under the control of the Ptetl-2 promoter and electroporated it into a B. burgdorferi 297 derivative expressing TetR from the chromosome as described above. Treating these B. burgdorferi with 1.5 Pg/ml of ATc increased BmpA detected by immunobloting (data not shown), and this increase was associated with a morphological alteration and clumping (Figure 6) and a decrease in growth that was proportional to the concentration of ATc to which these cells had been exposed [46, 49]. Taken together, these experiments suggest that BmpA is in fact an essential gene product for B. burgdorferi since alterations in its carefully controlled expression are detrimental to B. burgdorferi biology. They also indicate that the Tet system will be able to provide a means of studying the effects of positively and negatively altered expression of BmpA on B. burgdorferi physiology that will allow us to perform experiments regarding the potential function of this protein in B. burgdorferi biology. These experiments also illustrate the usefulness of Tet-controlled regulation of gene expression to approach the study of other putatively essential genes in B. burgdorferi.

A

B

Figure 6. B. burgdorferi 297/tetR pLD10 incubated in the absence (A) or presence of 1.5 Pg/ml of ATc for 24 h (Acridine orange; magnification, 1,250u .

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8. Conclusions We have applied two molecular genetic tools to analyze the role(s) of bmpC and bmpA in B. burgdorferi biology and pathogenesis. In the case of bmpC, we found that a null mutation had no effect on borrelial growth in vitro and only a minimal effect on infectivity in mice. In contrast, the bmpC null mutant were unable to grow in the joints or cause arthritis. Our inability to restore this element of pathogenicity by extrachromosomal complementation (even in the presence of all parental B. burgdorferi plasmids) makes this conclusion somewhat uncertain, however. It suggests that other approaches (for example, intrachromosomal complementation) may be needed to confirm that bmpC is involved in B. burgdorferi virulence, particularly because the presence of the pKFSS1 vector reduced the infectivity of the organisms (Table 4). The lack of knowledge of the function of BmpC also makes it difficult to speculate meaningfully on the mechanism or mechanisms involved in mediating this difference between systemic and tissue infection. They might involve transport of ions or nutrients that were in limiting concentrations in the joints but were not limiting in the general circulation. However, the differences between systemic and local responses in mice infected with the bmpC null strain are consistent with previously reported differences in these two attributes in borrelialinfected mice. For example, mice whose innate immune responses to infection with B. burgdorferi were impaired by the absence of the adapter molecule, MyD88, did not exhibit any diminution of inflammatory joint responses to borrelial infection [50-52]. Our results with the B. burgdorferi bmpC null mutant and previous reports with mutant mice both suggest that the mechanisms involved in borrelial infection and in tissue pathogenicity to this infection differ. In the case of bmpA, it was not possible to isolate a null mutant despite repeated attempts. It was therefore necessary to develop another method for artificially modulating and manipulating bmpA gene expression. To this end, we applied the TetR system developed by Bujard et al. for manipulation of gene expression in eukaryotic cells [42-45] and used by other workers to study gene expression in bacteria [38, 40, 41, 46] to B. burgdorferi. As expected, expression of foreign and native genes with the Tet system in B. burgdorferi was linearly dependent on the concentration of the ATc inducer. Interestingly, increased levels of induction of truncated BmpA (lacking the lipidation signal) and native BmpA from these TetR-regulated hybrid borrelial promoters were both associated with decreases in B. burgdorferi growth in vitro [data not shown]. Hyperproduction of BmpA also generated abnormal cell morphology and decreased viability. These alterations suggest that BmpA may play an essential role in B. burgdorferi physiology consistent with our inability to isolate null mutants of this gene. The borrelial Tet system we have developed is based on constructing hybrid B. burgdorferi promoters made responsive to the TetR repressor by the addition of DNA sequences containing one or two tet operators able to bind this repressor tightly. The other component of this system was constructed by designing B. burgdorferi strains where the expression of the TetR molecule was achieved from extrachromosomal and chromosomal sites under the control of the constitutively expressed B. burgdorferi flaB promoter. Small amounts of escape of repression were detected with hybrid borrelial promoters containing one and two tet operator sequences (data not shown). This might suggest an imbalance of TetR and tetO concentrations in the cells [42-45], a possibility that could be dealt with by placing both components of the TetR system in the B.

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burgdorferi chromosome. Another possible solution might be to fuse the tetR gene to a stronger B. burgdorferi promoter than the flaB promoter. Although the TetR repressor, like the lacZ repressor, LacI, can be toxic when expressed in different bacterial cells [40, 41, 46], we did not observe any negative influence upon B. burgdorferi growth and viability in the present study when the TetR repressor alone was expressed extrachromosomally or chromosomally under flaB promoter control. In summary, we have used two molecular genetics approaches to gain an initial insight into the function of two members of the bmp gene family. For bmpC, we used a null mutant to reveal the importance of this gene for growth of B. burgdorferi in the joints and for production of arthritis. In contrast, isolation of null mutants was not a fruitful approach to analyzing the role of bmpA in B. burgdorferi biology. It was therefore necessary for us to apply the Tet system developed by Bujard and co-workers [42-45] to Borrelia before any progress could be made in understanding the function of bmpA and its gene product in B. burgdorferi growth and physiology. Our results suggest that the borrelial-adapted Tet system we have developed when combined with antisense RNA technology will permit future isolation of conditional lethal mutants in B. burgdorferi [53].

Acknowledgements We thank Ms. Harriett V. Harrison for assistance in manuscript preparation. This work was supported by United States Public Health Service grants R01 AI08856 and R01 AI43063 to F.C.C.

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Molecular Biology of Spirochetes F.C. Cabello et al. (Eds.) IOS Press, 2006 © 2006 IOS Press. All rights reserved.

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Porins of Borrelia Marija PINNE a, Yngve ÖSTBERG a , Roland BENZ b and Sven BERGSTRÖM a,1 a Department of Molecular Biology, Umeå University, SE-901 87 Umeå Sweden b Lehrstuhl für Biotechnologie, Biozentrum der Universitaet Würzburg, Am Hubland, D97074 Würzburg, Germany Abstract. The outer surface-exposed proteins play a critical role in survival and pathogenesis of Borrelia in different hosts and tissues, being involved in avoiding the host immune response, adhesion to different tissues, and nutrient acquisition. This review describes the characteristics of borrelial integral outer membrane proteins that play a suggested role in solute and nutrient uptake and provide a possible role in the environmental adaptation of Borrelia. Three Borrelia burgdorferi proteins, P13, BBA01, and P66, were shown to be porins and characterized structurally and functionally using a combination of biochemical, biophysical, and genetic methods. The channel-forming function of the 13 kDa protein, P13, was elucidated by a lipid bilayer assay. Furthermore, post-translational processing of P13 has been shown to occur at the Cterminus by C-terminal processing protease (CtpA)-dependent cleavage. The membrane-spanning architecture of P13 was determined by epitope mapping and computer-based structural predictions, which revealed that P13 is an unusual porin, not possessing the structural properties of conventional porins. Rather than forming Ebarrels, it is predicted to span the membrane with hydrophobic D-helices. p13 belongs to a paralogous gene family. The transcription of p13 and other gene family members during in vitro growth and in a mouse infection model was therefore investigated. The paralog BBA01, which has the highest sequence homology to P13, is expressed during in vitro growth in all three Lyme disease causing species, although at very low levels. Like P13, BBA01 is also processed by CtpA and exhibits very similar channel-forming activity. Furthermore, in the absence of P13, a proportion of total BBA01 protein is relocated to the bacterial surface with strong indications that BBA01 and P13 are functionally interchangeable. P66, found to be an integrin binding protein, was also determined to be a porin. Keywords. Channel forming proteins, outer membrane, integrin binding, spirochetes

Introduction The outer membrane of bacteria is a selective barrier between the cell and the environment that excludes certain molecules and simultaneously allows entry of appropriate nutrient substances. The outer membrane of conventional Gram-negative bacteria consists of an 1

Corresponding Author: Sven Bergström, Department of Molecular Biology, Umeå University, SE-901 87 Umeå, Sweden. E-mail: [email protected].

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asymmetrical bilayer with an inner leaflet composed of phospholipids and an outer leaflet containing lipopolysaccharides (LPS) [1]. Interestingly, LPS and phosphatidylethanolamine are notable omissions from the outer membrane of Borrelia [2, 3]. Moreover, this outer membrane also has a very low density of integral, membrane-spanning proteins [4, 5] but maintains an extraordinary high abundance of lipoproteins [6, 7]. In fact, 8% (105 in total) of all Borrelia burgdorferi ORFs are predicted to encode lipoproteins, the highest frequency among those bacterial genomes sequenced [7, 8]. This number could even be higher, since a new computer algorithm, SpLip, designed specifically to identify spirochetal lipoproteins, predicted 127 lipoprotein ORFs in B. burgdorferi [9]. Outer membrane proteins (OMPs) of Gram-negative bacteria are of great interest because of their location at the cell surface, which may facilitate interactions of bacterial pathogens with their host. In particular, OMPs may play a role in pathogenesis by acting as (i) adhesins, (ii) targets for bactericidal antibodies, (iii) receptors for various molecules, or (iv) porins. In addition, the selective permeability of the outer membrane is due to the physico-chemical properties of the membrane itself and to the action of specific binding and uptake molecules associated with the outer membrane. Substances can be taken into bacteria by diffusion through the outer membrane via porins [10] or by receptor-mediated uptake [11]. Clearly, OMPs allow Borrelia to adapt and respond to the different environments during its life cycle, in this way playing a critical role in survival, biology, and infectivity of Lyme disease spirochetes. Since the outer membrane also makes direct contact with the host, B. burgdorferi molecules exposed on the bacterial surface must mediate the intera