Advances in Applied Microbiology, Volume 74

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Sui...

Author: Allen I. Laskin | Geoffrey M. Gadd | Sima Sariaslani

36 downloads 1201 Views 3MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!
Report copyright / DMCA form

DOWNLOAD PDF

Academic Press is an imprint of Elsevier 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 32, Jamestown Road, London NW1 7BY, UK First edition 2011 Copyright # 2011 Elsevier Inc. All rights reserved No part of this publication may be reproduced, stored in a retrieval system or transmitted in any form or by any means electronic, mechanical, photocopying, recording or otherwise without the prior written permission of the publisher Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone (+44) (0) 1865 843830; fax (+44) (0) 1865 853333; email: [email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://elsevier.com/locate/permissions, and selecting Obtaining permission to use Elsevier material Notice No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnoses and drug dosages should be made ISBN: 978-0-12-387022-3 ISSN: 0065-2164 For information on all Academic Press publications visit our website at www.elsevierdirect.com Printed and bound in the USA 11 12 13 14 10 9 8 7 6 5

4 3 2 1

CONTRIBUTORS

David E. Arnot Centre for Medical Parasitology, Department of International Health, Immunology and Microbiology, Faculty of Health Sciences, University of Copenhagen; Department of Infectious Diseases, Copenhagen University Hospital (Rigshospitalet), CSS Oester Farimagsgade 5, Copenhagen K, Denmark; and Institute of Immunology and Infection Research, School of Biology, University of Edinburgh, Edinburgh, Scotland, United Kingdom Adi Dror Department of Science Education—Biology, Faculty of Natural Sciences, University of Haifa-Oranim, Tivon, Israel Kevin W. George Field of Environmental Toxicology; Department of Microbiology, Wing Hall, Cornell University Ithaca, New York, USA Malka Halpern Department of Science Education—Biology, Faculty of Natural Sciences, University of Haifa-Oranim, Tivon, Israel Absarul Haque Department of Microbiology, Molecular Genetics and Immunology, University of Kansas, Medical Center, Kansas City, Kansas, USA Abdul Haseeb Department of Microbiology, Molecular Genetics and Immunology, University of Kansas, Medical Center, Kansas City, Kansas, USA Anthony G. Hay Field of Environmental Toxicology; Department of Microbiology, Wing Hall, Cornell University Ithaca, New York, USA Islam T. M. Hussein Department of Microbiology, Molecular Genetics and Immunology, University of Kansas, Medical Center, Kansas City, Kansas, USA

vii

viii

Contributors

Anja T. R. Jensen Centre for Medical Parasitology, Department of International Health, Immunology and Microbiology, Faculty of Health Sciences, University of Copenhagen; Department of Infectious Diseases, Copenhagen University Hospital (Rigshospitalet), CSS Oester Farimagsgade 5, Copenhagen K, Denmark Simcha Lev-Yadun Department of Science Education—Biology, Faculty of Natural Sciences, University of Haifa-Oranim, Tivon, Israel Mohammad A. Mir Department of Microbiology, Molecular Genetics and Immunology, University of Kansas, Medical Center, Kansas City, Kansas, USA Avivit Waissler Department of Science Education—Biology, Faculty of Natural Sciences, University of Haifa-Oranim, Tivon, Israel

CHAPTER

1 Bacterial Strategies for Growth on Aromatic Compounds Kevin W. George*,† and Anthony G. Hay*,†,1

Contents

Abstract

2 3 5 5

I. Introduction II. Overview of Aerobic Aromatic Degradation A. The upper pathway B. The lower pathway C. C23O inactivation and the production of toxic metabolites D. Genetic organization and regulation III. Bacterial Strategies for Growth on Aromatic Compounds A. Recruitment of catabolic genes B. Expression of repair enzymes C. Direct modulation of substrate range or kinetic parameters IV. Conclusion References

6 8 9 9 17 20 24 25

Although the biodegradation of aromatic compounds has been studied for over 40 years, there is still much to learn about the strategies bacteria employ for growth on novel substrates. Elucidation of these strategies is crucial for predicting the environmental fate of aromatic pollutants and will provide a framework for the development of engineered bacteria and degradation pathways. In this chapter, we provide an overview of studies that have advanced our knowledge of bacterial adaptation to aromatic compounds.

* Field of Environmental Toxicology, Cornell University Ithaca, New York, USA { 1

Department of Microbiology, Wing Hall, Cornell University Ithaca, New York, USA Corresponding author: e-mail address: [email protected]

Advances in Applied Microbiology, Volume 74 ISSN 0065-2164, DOI: 10.1016/B978-0-12-387022-3.00005-7

#

2011 Elsevier Inc. All rights reserved.

1

2

Kevin W. George and Anthony G. Hay

We have divided these strategies into three broad categories: (1) recruitment of catabolic genes, (2) expression of ‘‘repair’’ or detoxification proteins, and (3) direct alteration of enzymatic properties. Specific examples from the literature are discussed, with an eye toward the molecular mechanisms that underlie each strategy.

I. INTRODUCTION Microorganisms possess the extraordinary capacity to degrade a multitude of pollutants, many of which have only recently been introduced to the environment due to human activities. Of these pollutants, aromatic compounds are particularly problematic due to their resistance to degradation and deleterious effects on human and environmental health (ATSDR, 2007; Milennium Assessment, 2005). Given these concerns, it is critical to understand the strategies microorganisms employ to adapt to growth on aromatic substrates. The recent oil spill in the Gulf of Mexico is a topical reminder of the indispensible role microorganisms play in the degradation of petroleum hydrocarbons and the importance of microbial adaptation. Aromatic compounds comprise on average 15% of crude oils and as much as 70% of gasoline by volume (Hyne, 2001; Tessoro, 2003). A recent study of indigenous oil-degrading bacteria in oil-impacted areas of the Gulf of Mexico found that hydrocarbon biodegradation rates were faster than expected and revealed significant potential for intrinsic bioremediation of the oil (Hazen et al., 2010). Within the plume, the population of oildegrading bacteria was enriched as were genes associated with the degradation of key aromatic oil constituents such as isopropylbenzene and naphthalene. The bacterial capacity to metabolize a wide range of diverse aromatic substrates is likely related to the fact that the common unit of aromatic compounds—the benzene ring—is among the most ubiquitous chemical structures in nature (Harwood and Parales, 1996). This abundance has contributed to the selective pressure that has driven the evolution of a relatively conserved, yet genetically flexible, mechanism for degradation that relies on the addition of either one or two atoms of molecular oxygen to the ring (Harayama et al., 1992). Encoded by a significant amount of genetic diversity, the mixed function oxygenases that perform these reactions have been the subject of both basic and applied research. Much of this research has focused on their potential as catalysts for specialty chemical production (Cirino and Arnold, 2002; Gibson and Parales, 2000) and bioremediation (Gibson and Parales, 2000; Pieper and Reineke, 2000; Urgun-Demirtas et al., 2006).

Bacterial Strategies for Growth on Aromatics

3

Despite the vast metabolic potential of bacteria, aromatic biodegradation is not always effective, particularly in situ. There are numerous potential reasons for this, ranging from unfavorable growth conditions to limited pollutant bioavailability (van der Meer et al., 1992). In the case of structurally complex xenobiotics such as halogenated pesticides, a more obvious factor is the absence of preexisting bacterial enzymes capable of substrate degradation. Given that the chemical structure of certain xenobiotics may be quite different from the substrate specificity of existing enzymes, it is not surprising that mutations which expand substrate range or increase degradation efficiency may be required before a bacterium is capable of degrading a compound, let alone using it for growth. In some instances, alternative catabolic enzymes must be ‘‘recruited’’ either from within a bacterium’s genome or horizontally from other organisms. Aromatics that are toxic or produce toxic metabolites present additional challenges to bacteria and require their own unique adaptive responses and strategies. The need for these various mutations can often result in extended periods of adaptation prior to pollutant degradation (van der Meer, 2006; van der Meer et al., 1987, 1998). Understanding the mechanisms that underlie bacterial adaptation to novel aromatic substrates is likely to yield improved predictions of biodegradation potential and provide a framework for directed pathway evolution (Diaz, 2004). In this review, we describe some molecular mechanisms and bacterial strategies for growth substrate expansion and improved biodegradation of aromatic substrates. Before providing specific examples, we first review the conserved mechanisms of aerobic aromatic degradation. A brief overview of regulation in aromatic degradation pathways follows, with a concise discussion of genetic organization. The bulk of this review then focuses on genetic adaptations and strategies that allow bacteria to metabolize aromatic compounds. Broadly speaking, we discuss adaptations that fall into three main categories, resulting in either catabolic gene recruitment, expression of specialized ‘‘repair’’ proteins, or direct alteration of enzymatic properties. Though these strategies are discussed separately, it is important to note that they can and likely do occur simultaneously, working in concert to increase degradation capacity, decrease toxicity, and thereby expand growth substrate range.

II. OVERVIEW OF AEROBIC AROMATIC DEGRADATION Although both aerobic and anaerobic biodegradation contribute to the elimination of aromatic pollutants from the environment, the aerobic mechanism is much more prevalent in the biosphere (Cao et al., 2009) and will be the focus of this review. The aerobic degradation of both mono- and polycyclic aromatic hydrocarbons normally proceeds via

4


O

O

[O2] Upper pathway

OH

Ring hydroxylation

Catechol

OH

Lower pathway

[O2]

Metacleavage

Ring cleavage Funneling to TCA cycle

Orthocleavage

OH COOH COOH CHO

COOH

Dehydrogenase pathway

Hydrolytic pathway

O O

O

O HO

H3C

SCoA

H3C

COOH

H3C

OH

SCoA O

Acetyl-coA

Pyruvate

Acetyl-coA

Succinate

Tricarboxylic acid cycle FIGURE 1.1 General schematic of aerobic aromatic degradation. The dashed line divides upper and lower pathways.

two major steps, designated as the upper (or peripheral) and lower (or ring cleavage) pathways (Diaz, 2004) (Fig. 1.1). In the upper pathway, the critical step is the destabilization of the ring through mono- or dioxygenation, typically resulting in the addition of two hydroxyl groups (Mason and Cammack, 1992). The remainder of the upper pathway focuses on preparing the ring for cleavage, typically using dehydrogenation to form a catecholic intermediate. Following the formation of catechol or a closely related monocyclic compound such as gentisate, the lower pathway begins when the hydroxylated aromatic ring is cleaved by a second dioxygenase (Harayama and Rekik, 1989; Harayama et al., 1992; Vaillancourt et al., 2006). Once the ring is cleaved, further transformations usually funnel ring fission products into the Tricarboxylic acid cycle for energy production. Additional detail regarding the upper and lower pathways is provided below.

5


A. The upper pathway Due to the stability of both the aromatic ring and molecular oxygen (O2), the formation of a reactive oxygen intermediate is a prerequisite for ring oxidation. Ring-hydroxylating oxygenases are responsible for catalyzing the incorporation of oxygen atoms into the aromatic ring. Many of the best described oxygenases are Rieske, nonheme iron oxygenases comprising multicomponent complexes, and usually include electron transport proteins which transfer electrons from NAD(P)H to a terminal, iron–sulfur oxygenase (Butler and Mason, 1997; Gibson and Parales, 2000) (Fig. 1.2). The electron transport component often consists of an iron–sulfur ferredoxin along with a separate reductase or a combined ferredoxin–NADH-reductase (Gibson and Parales, 2000). The terminal oxygenase is typically a heteromultimer, composed of a large (a) and small (b) subunit although the arrangement of subunits varies (Furusawa et al., 2004). The a subunit contains a Rieske-type [2Fe–2S] cluster which is reduced by electron-transfer proteins and functions as an oxygen activation center (Butler and Mason, 1997). This subunit is also responsible for substrate binding and recognition (Furusawa et al., 2004; Gibson and Parales, 2000). Following the incorporation of oxygen, dehydrogenation usually takes place resulting in catechol or related structures such as gentisate, hydroquinone, or salicylate (Gibson and Subramanian, 1984).

B. The lower pathway Following the creation of a catecholic intermediate, ring fission occurs through ortho- or meta-cleavage. During ortho-cleavage, dioxygenation takes place at the 1,2-position of the catechol (within the hydroxyl groups), OH

NAD(P)H + H+

NAD(P)+

2H

OH

+

Oxidized

Oxidized

Oxidized

Reduced

Reduced

Reduced 2H+

Component Subunit composition Prosthetic group

ReductaseNAP

FerredoxinNAP

α

α

FAD, [2Fe–2S]

[2Fe–2S]

+ O2

OxygenaseNAP α3β3 3[2Fe–2S], 3Fe(II)

FIGURE 1.2 Organization of the components of the naphthalene dioxygenase system from Pseudomonas sp. strain NCIB 9816-4, a well-characterized ring-hydroxylating dioxygenase. The flow of electrons is illustrated in the top panel, while subunit composition and prosthetic groups are labeled below. Adapted from Gibson and Parales (2000).

6


Extradiol cleavage OH COOH CHO

metaCatechol-2,3-dioxygenase (C23O)

Intradiol cleavage OH

ortho-

Catechol-1,2-dioxygenase OH (C12O)

COOH COOH

FIGURE 1.3 Extradiol (meta-) versus intradiol (ortho-) cleavage. Extradiol cleavage takes place at the 2,3-position on the catechol (outside the hydroxyls), while intradiol cleavage takes place at the 1,2-position (inside the hydroxyls). The reactions are catalyzed by catechol-2,3-dioxygenases (C23Os) and catechol-1,2-dioxygenases (C12Os), respectively.

while during meta-cleavage fission takes place outside the hydroxyls and usually occurs at the 2,3-position (Harayama and Rekik, 1989) (Fig. 1.3). Distal ring fission in the 1,6-position of the catechol has also been reported (Horvath, 1970; Koh et al., 1997). Ortho- and meta-cleavages are catalyzed by intradiol and extradiol dioxygenases, which use Fe(III) and Fe(II) at the active site to catalyze ring cleavage, respectively (Harayama et al., 1992). Although these differences between the dioxygenases seem trivial, the enzymes have vastly different structures and dissimilar catalytic mechanisms (Vaillancourt et al., 2006). In general, extradiol dioxygenases (catechol-2,3-dioxygenases, C23Os) appear to be more versatile and occur in far more catabolic and biosynthetic pathways (Vaillancourt et al., 2006). Most products generated by meta-cleavage (meta-fission products, MFPs) take the form of a common structure, 2-hydroxy-6-oxohexa-2,4dienoate (HODA), with different substituents on the C6 carbon depending on the substrate (Khajamohiddin et al., 2008). Further degradation of MFPs proceeds via either a hydrolytic or an NAD-dependent dehydrogenase/4oxalocrotonate pathway (Sala-Trepat et al., 1972) (Fig. 1.4). These branches, and the specific reactions they catalyze, are key determinants of substrate range in aromatic degradation pathways: each branch has a limited substrate range that is constrained by the nature and position of the substituent(s). Para- or nonsubstituted catechols form aldehyde group containing MFPs and are acted upon by the dehydrogenase pathway, while cleavage of three-substituted catechols generates a ketone, which proceeds through the hydrolytic branch. Since most degradation pathways do not contain both branches, the nature of the lower pathway becomes a key determinant of substrate range. Within the hydrolytic branch, MFP hydrolases possess strict substrate specificities and appear to play crucial roles in determining substrate preference in bacteria (Khajamohiddin et al., 2008).

C. C23O inactivation and the production of toxic metabolites Many intermediates produced during aromatic metabolism are capable of causing cellular toxicity (Chavez et al., 2006; Gerischer and Ornston, 1995; Park et al., 2004; Perez-Pantoja et al., 2003; Pumphrey and Madsen, 2007)


7

R1 OH

R2

OH O2

C23O

Dehydrogenase branch

R1 R2

Hydrolytic branch

O COOH

R2

COOH COOH

Dehydrogenase OH

OH MFP hydrolase

Isomerase

R2

CO COOH O

R2

COOH COOH

Decarboxylase O

Hydratase

H2O

R2 COOH HO

O

Aldolase

Acetyl-coA + pyruvate

FIGURE 1.4 Meta-fission lower pathway for catechol and substituted catechols. The hydrolytic and dehydrogenase branches are denoted. Enzymes typically responsible for catalysis are labeled. Adapted from Khajamohiddin et al. (2008).

and require specific bacterial adaptations for degradation. Catecholic compounds, perhaps the most common intermediates in aromatic degradation, are particularly problematic. During the course of catalysis, C23Os may become inactivated by their catecholic substrates in a process known as mechanism-based or suicide inhibition (Bartels et al., 1984; Klecka and Gibson, 1981) which involves oxidation of active site Fe(II)–Fe(III) (Cerdan et al., 1994; Vaillancourt et al., 2002). This sort of C23O inactivation has been primarily observed with chlorocatechols, but some alkyl catechols can exert a similar effect (Vaillancourt et al., 2006). C23O inactivation and the subsequent accumulation of catechol have been shown to limit the substrate range of a variety of pathways (see Section III.B.1). In addition to suicide inhibition, catechols can initiate toxicity through a diverse range of molecular mechanisms, ranging from direct protein damage to the production of reactive oxygen species (ROS) (Schweigert

8


et al., 2001a,b). Despite this marked toxicity, the production of catechols appears to be a ‘‘necessary evil’’ in aromatic degradation. Indeed, catechol-producing oxygenases are among the most conserved enzymes in aromatic degradation pathways, likely due to the requirement for ring destabilization prior to cleavage and the abundance of molecular oxygen. Since the production of catechol is not easily avoided, bacteria have developed multiple strategies to minimize C23O inactivation and alleviate deleterious effects. Previously described strategies include the expression of ‘‘repair’’ enzymes (Cerdan et al., 1994; Hugo et al., 1998, 2000; Park et al., 2002; Polissi and Harayama, 1993), the selection for, or recruitment of inactivation-resistant C23Os (Cerdan et al., 1994; Ramos et al., 1987; Rojo et al., 1987), and the selection of gene duplications which increase catechol consumption (Perez-Pantoja et al., 2003). Additionally, alterations in the kinetic properties of key enzymes including a recently reported ‘‘less is more’’ strategy (George, 2011) can also dramatically affect the range of aromatic substrates that individual bacteria can use for growth. Specific examples of these adaptations are discussed in the remainder of this review.

D. Genetic organization and regulation Genes encoding aromatic degradation pathways are typically arranged in operons or clusters. These clusters usually comprise catabolic genes which encode degradative enzymes, transport genes encoding proteins which permit uptake of the compound, and one or more regulatory genes which control total gene expression (Diaz, 2004; Khomenkov et al., 2008). The organization of a typical catabolic operon, encoding the toluene degradation (tod) pathway of Pseudomonas putida F1 (Zylstra and Gibson, 1989; Zylstra et al., 1988), is depicted in Fig. 1.5. Catabolic gene clusters are often present on plasmids which facilitate horizontal gene transfer between hosts. Even when encoded on chromosomal DNA, genes associated with aromatic degradation are frequently flanked by mobile genetic elements or located within large transposons (Top and Springael, 2003; van der Meer et al., 1992). These mobile elements can facilitate rearrangement not only within the genome but also between hosts through association with other mobile elements such as plasmids. Proper functioning of a pathway is dependent not only on the possession of appropriate catabolic enzymes but also on regulatory proteins that modulate gene expression in the presence of the suitable substrates. In catabolic pathways, regulatory systems generally fall into discrete families (Tropel and van der Meer, 2004). LysR-type regulators, for instance, comprise the largest family of bacterial regulatory proteins and function in the degradation of numerous aromatic compounds such as naphthalene and chlorobenzene. Other families include the IclR family,

9


CH3

todC1C2BA

CH3

CH3 OH

NADH H+ + O2

OH

TodD

TDO NAD+

OH

CH3

OH

NAD+ NADH H+

O COO–

TodE

OH

O2

TodF O

O

TodI

H3C C SCoA

H3C NADH CoASH + + H NAD

TodH

C H CH3

X

Transport

HO

OH

OH H2O

COO–

F

C1

COO–

TodG

C O

1 kb

R

COO–

H2O

C2 B A

D

Regulatory genes

E

G

l

H

S

T

Catabolic genes

FIGURE 1.5 Degradation pathway and genetic organization of the tod operon. The degradation of toluene is pictured. The todXFC1C2BADEGIH operon is transcribed from a single promoter, PtodX. Transcriptional activation is mediated by a two-component system (TodS and TodT), encoded by genes located downstream of the catabolic operon.

the XylR/NtrC family, and the AraC/XylS family, among others (Tropel and van der Meer, 2004). Extensive reviews of transcriptional regulators in aromatic degradation pathways have been published elsewhere and are beyond the scope of this review (Diaz and Prieto, 2000; Tropel and van der Meer, 2004). Intriguingly, closely related catabolic genes in distinct microorganisms are often regulated by different classes of regulators, suggesting that the construction of a catabolic operon and the acquisition of transcriptional control are independent events (Cases and de Lorenzo, 2001; Shingler, 2003).

III. BACTERIAL STRATEGIES FOR GROWTH ON AROMATIC COMPOUNDS A. Recruitment of catabolic genes The recruitment of alternative catabolic genes is perhaps the most widely reported bacterial strategy for adapting to growth on aromatic compounds. Gene recruitment occurs in two different ways: vertical, or recruitment of catabolic function(s) from within a bacterium’s genome, and horizontal, or recruitment involving gene transfer between bacteria.

10


1. Vertical recruitment The vertical recruitment of catabolic function utilizes a bacterium’s endogenous genome. While vertical recruitment is not capable of the sort of large-scale expansion of catabolic genes which characterize horizontal transfer, it can nevertheless have profound effects on bacterial adaptation and can even augment the effects of horizontal transfer. Strategies discussed below include changes in regulatory function, the spontaneous duplication of genes or clusters of genes, and the rearrangement or shuffling of preexisting genes.

a. Changes in regulation Mutations that result in changes in regulation are among the simplest adaptations which expand growth substrate range for aromatic compounds. In certain instances, aromatic substrates may not be degraded because transcriptional regulators do not recognize them as effectors. In these cases, even though cells possess the genetic capability to use the compounds for growth, the required catabolic proteins are not expressed. Mutations in regulatory proteins which alter effector specificity can cause pathway induction and result in growth substrate expansion. In pathways regulated by repressors, point mutations can prevent repressor binding, resulting in constitutive expression of requisite proteins. In rare occurrences, mutations can create promoter binding sites, resulting in increased transcription and expression of catabolic genes. Insertion sequence elements (IS elements) are abundant in bacterial genomes (Mahillon and Chandler, 1998; Siguier et al., 2006) and have also been found to modulate regulatory function by increasing transcription of downstream genes (van der Meer, 2002). Pseudomonas azelaica HBP1, for example, is an organism capable of growing on 2-hydroxybiphenyl (Kohler et al., 1988) and encodes enzymes which are capable of mineralizing substituted phenols such as isopropylphenol. Despite this catabolic potential, HBP1 cannot grow on substituted phenols as sole carbon sources because HbpR, the positive regulator of the hbp pathway, is unable to recognize them as effectors ( Jaspers et al., 2000). When exposed to isopropylphenol for long periods of time, however, HBP1 mutants were generated which could grow on substituted phenols. In these adapted strains, mutations in hbpR expanded the range of effectors recognized by the regulator, resulting in transcription and growth on substituted phenols ( Jaspers et al., 2000; Kohler et al., 1993). In Pseudomonas sp. CF600, mutations in the regulator dmpR were similarly capable of broadening effector specificity (Pavel et al., 1994) or conferring constitutive phenotypes (Shingler and Pavel, 1995). An abundance of work on transcriptional control within the TOL plasmid of P. putida has revealed analogous mutations in the regulators XylR and XylS (Dominguez-Cuevas et al., 2008; Ramos et al., 1997). In one study, it was


11

found that single amino acid substitutions in the signal receptor domain of XylR conferred constitutive expression of TOL genes in the absence of an effector (Delgado et al., 1995). Other reports have demonstrated that mutations in the positive regulator XylS can lead to either constitutive phenotypes (Zhou et al., 1990) or changes in effector specificity (Ramos et al., 1986) and thereby expand growth substrate range. Comamonas testosteroi TA441 contains a silent cluster of genes (aphKLMNOPQB) encoding the catabolic enzymes phenol hydroxylase and catechol 2,3-dioxygenase, along with aphR, a divergently transcribed regulatory gene (Arai et al., 1999b). In this pathway, it was found that mutations in aphS, a gene encoding a negative transcriptional regulator, derepressed transcription of the aph genes and conferred growth on phenol (Arai et al., 1999a). In certain degradation systems, multiple regulatory mutations are required to confer growth on new aromatic substrates. In P. putida F1 and related strains, transcription of the toluene degradation (tod) pathway (Zylstra and Gibson, 1989; Zylstra et al., 1988) is regulated by the twocomponent system TodST (Lau et al., 1997). TodS, the sensor kinase, recognizes a wide range of effectors (Busch et al., 2007; Lacal et al., 2006) and phosphorylates TodT (Lacal et al., 2008), the response regulator responsible for binding the tod promoter and activating transcription. Choi et al. (2003) demonstrated expansion of growth substrate range in F1 to include n-propylbenzene, n-butylbenzene, cumene, and biphenyl following two separate mutations in regulatory function. Mutations in todS expanded the range of effectors capable of activating transcription of the tod genes, while a mutation in cymR, a repressor of the cumate (cmt) operon (Eaton, 1996, 1997), allowed for constitutive expression and recruitment of the broad substrate MFP hydrolase CmtE (Choi et al., 2003). In wild-type F1, MFPs accumulated due to the inability of TodF, the endogenous MFP hydrolase of the tod pathway, to degrade them. Impaired CymR binding also allowed for expanded growth substrate range in P. putida CE2010, although this was achieved through a mutation in the cmt promoter rather than a mutation in the repressor itself (Ohta et al., 2001). In P. putida KL47, a strain capable of growing on diverse substrates such as biphenyl and cumate, mutation of TodS was identified as a crucial factor in catabolic adaptation. Although, once again, CymR binding was impaired, allowing constitutive expression of the cumate operon and the broad substrate MFP hydrolase CmtE as well (Lee et al., 2006). In rare instances, mutations can create consensus promoters upstream of catabolic genes. Using a promoterless phenol degradation operon (pheBA) in P. putida, it was shown that exposure to phenol generated mutants capable of growth (Pheþ). Further analysis of Pheþ mutants revealed that specific base substitutions were predominant in

12


phenol-exposed cells and resulted in the formation of a sequence similar to the consensus s70 promoter (Kasak et al., 1997). It was also found that the introduction of an IS elements could result in the creation of a new promoter and generate Pheþ mutants. Some IS elements, such as IS1411 (Kallastu et al., 1998), contain sequences near their termini which function as constitutive promoters. When these IS elements insert upstream of catabolic operons, constitutive transcription may take place. Working with promoterless pheBA genes expressed in P. putida PaW85 (Kasak et al., 1993), it was found that some spontaneous Pheþ mutants had IS1411 inserted upstream of pheB (Kallastu et al., 1998). Since IS1411 contains a strong promoter at its 30 end, its integration upstream of pheBA drove transcription and conferred the ability to grow on phenol. An earlier study also found that transposon integration upstream of pheBA, combined with point mutations in the promoter, permitted P. putida to grow on phenol (Nurk et al., 1993). In addition to creating new promoters, IS elements are also capable of modulating transcription through interruption of genes encoding repressors or their binding sites as discussed above.

b. Altering gene dosage Through the action of genetic engineering, homologous recombination, or transposition, the copy number of genes or clusters of genes may be increased. Duplications have long been considered an essential mechanism in the evolution of xenobiotic degradation pathways (van der Meer et al., 1992). Once a gene has been duplicated, the sequence of the redundant gene copy may diverge and accumulate mutations at an increased rate. As a result of mutation, substrate range or catalytic efficiency may be drastically altered (see Section III.C). In some cases, this may expand the range of aromatic substrates available for growth. Duplications of genes have been frequently observed in aromatic degradation pathways. In pathways which produce toxic intermediates such as chlorocatechols, specific catabolic gene duplications appear to serve novel roles in the mitigation of toxicity. Ralstonia eutropha JMP134 (pJP4) (formerly known as Alcaligenes eutrophus), originally isolated for its ability to metabolize 2,4-dichlorophenoxyacetic acid (2,4-D), is also capable of growing on 3-chlorobenzoate through the tfd pathway. Plasmid pJP4 includes two apparently isofunctional catabolic gene clusters, tfdCIDIEIFI and tfdDIICIIEIIFII (Laemmli et al., 2000; Trefault et al., 2004), which encode proteins involved in chlorocatechol metabolism. Of particular importance is chlorocatechol-1,2-dioxygenase (TfdC), the enzyme responsible for ortho-ring cleavage. Intriguingly, it was found that multiple copies of tfdC were required for R. eutropha JMP(pJP4) to grow on 3chlorobenzoate (Perez-Pantoja et al., 2003). In mutants which possessed only one tfdC gene, 3-chlorocatechol accumulated and caused marked


13

toxicity. Thus, increasing the gene dosage of tfdC through duplication augmented the rate of chlorocatechol turnover, preventing toxicity and expanding growth substrate range. Duplications of chlorocatechol dioxygenase genes were also observed in the clc-element of Ralstonia sp. strain JS705 and might play similar roles (Muller et al., 2003; van der Meer and Sentchilo, 2003; van der Meer et al., 1998). In the laboratory, gene dosage can be increased through the expression of plasmid-borne catabolic genes. In P. putida F1, for instance, a construct which overexpressed TodE, the endogenous C23O of the tod operon, was capable of growth on styrene, while the wild-type strain was not (George et al., 2011). Expression of TodE in trans mimicked the effect of gene duplication and enhanced the rate of 3-vinylcatechol turnover, preventing toxicity in a similar fashion as reported for R. eutropha JMP(pJP4). Since duplications create homologous regions of DNA, they may direct catabolically relevant recombination mechanisms. If present on a plasmid, for instance, these duplications can cause integration of catabolic genes into the chromosome. This appears to be the case in the TOL plasmid, where two direct repeats have been suggested to be necessary for integration of this catabolic operon into the chromosome (Meulien et al., 1981; Sinclair et al., 1986). Thus, in addition to increasing gene dosage and the amount of critical enzymes, duplications can also play essential roles in the dissemination of catabolic genes through horizontal transfer (see Section III.A.2). Similar gene repeats have been found in numerous other catabolic plasmids and may similarly allow for recombination into or out of chromosomal DNA (van der Meer, 2002).

c. Genetic rearrangement and gene capture In nearly every described catabolic pathway, there is clear evidence that gene rearrangements have occurred (van der Meer, 2002). Transposons or IS elements are located within or in close proximity to many catabolic genes, suggesting that they have played a role in DNA shuffling and the movement of genes throughout the host’s genome. In Pseudomonas sp. strain P51, for instance, the tcbA and tcbB genes for chlorobenzene dioxygenase and chlorobenzene dihydrodiol dehydrogenase are located on a transposable element, Tn5280 (van der Meer et al., 1991). Tn5280 appeared to function as a composite transposon (Top and Springael, 2003), with nearly identical IS elements (IS1066 and IS1067) located at each end. The structure was mobile, and it was suggested that these IS elements had captured the tcaAB genes from another organism—likely with the aid of an additional mobile element (see Section III.A.2)—and subsequently grouped them with preexisting genes for chlorocatechol metabolism (van der Meer et al., 1991; Werlen et al., 1996). This interplay between horizontal transfer and vertical rearrangement is typical in aromatic degradation pathways. It is evident that similar events have occurred in numerous other

14


organisms and played crucial roles in pathway evolution. A number of additional examples involving gene rearrangement in catabolic pathways have been reviewed elsewhere (Khomenkov et al., 2008; Springael and Top, 2004; Top and Springael, 2003). As previously noted (see Section II.D), catabolic pathways tend to be organized into gene clusters which are under the control of one or more regulatory proteins (Tropel and van der Meer, 2004). Analysis of flanking regions makes it clear that IS elements and other mobile elements are responsible for the majority of this organization (Khomenkov et al., 2008). The clustering of catabolic genes into upper and lower pathways, along with the recruitment of regulatory control, is important for the assembly of a contiguous transcriptional unit and thus the energetically efficient degradation of aromatic compounds (van der Meer, 2002). Although pathway genes are usually grouped into one or two transcriptional units, this is not always the case. In the dibenzodioxin and dibenzofuran degrader Sphingomonas sp. strain RW1, for instance, catabolic genes are scattered throughout the genome (Armengaud et al., 1998). Intriguingly, while expression of upper pathway genes is modulated according to the presence of a suitable carbon source, expression of dbfB, the ring cleavage dioxygenase, is constitutive. This sort of differential expression is unusual and contrasts with many other well-studied catabolic pathways. In the course of analysis, numerous other catabolic genes were discovered dispersed throughout the genome (Armengaud et al., 1998). This scattering of diverse catabolic genes has been observed in other Sphingomonas strains such as Sphingomonas yanoikuyae B1 (Kim and Zylstra, 1999) and appears to be consistent with the catabolic diversity of this genus: although energetically expensive in some environments, a lack of coordinate regulation and constitutive expression of certain catabolic gene modules may aid in the utilization of varied aromatic substrates for growth.

2. Horizontal recruitment Horizontal recruitment of catabolic genes is mediated by a variety of mobile genetic elements which comprise a ‘‘horizontal gene pool’’ (Thomas, 2000). Elements capable of horizontal transfer include plasmids, transposons, and ‘‘genomic islands,’’ all of which are capable of transferring catabolic genes between disparate bacterial hosts. This transfer allows bacteria to rapidly adapt to, and degrade, aromatic compounds—even those compounds which are ‘‘newcomers’’ to the global environment (Top and Springael, 2003; van der Meer and Sentchilo, 2003). Although gene recruitment through plasmid transfer and transposition is discussed separately below, these elements often work in tandem in vivo. For instance, many plasmid-encoded catabolic pathways such as the TOL catabolic operon are flanked by ISs or occur within transposable elements.


15

a. Plasmid transfer In aromatic degradation, the importance of plasmidencoded catabolic genes cannot be overstated. The transfer of a plasmid is a one-step process which can result in the acquisition of up to 250 kb of DNA or more and can drastically increase catabolic capacity. Natural plasmid transfer—combined with the potential for mutation—creates enormous genetic potential in aromatic-degrading bacteria. The fact that some plasmids are capable of retrotransfer, or the capture of chromosomally encoded catabolic genes, adds to this tremendous potential (Ronchel et al., 2000; Trefault et al., 2004). Nearly every catabolic pathway described in the literature, even those that are not plasmid encoded, is associated with the presence of plasmids (van der Meer, 2002). Perhaps the most well-studied example is the TOL plasmid, encoding a pathway for the degradation of toluene and related compounds (Worsey and Williams, 1975). Plasmids involved in the degradation of naphthalene, xylenes, chlorobenzenes, and atrazine (among many others) have also been described (Cao et al., 2009; Dennis, 2005). The majority of these plasmids are transmissible and there is strong evidence for widespread natural plasmid transfer in situ (Dejonghe et al., 2000; Herrick et al., 1997; Peters et al., 1997; Springael and Top, 2004; Top et al., 2002). For example, phenoldegrading P. putida PaW85, containing plasmid-encoded pheAB genes, was deliberately released into a mine water-contaminated river. Several years following this release, it was found that approximately one-third of phenol-degrading bacteria isolated from the site contained the plasmidencoded pheAB genes. Intriguingly, these bacteria were all different species than the original donor strain, strongly suggesting horizontal gene transfer (Peters et al., 1997). More recently, a series of studies utilized 2,4D as a model compound and two plasmids which encoded 2,4-D degradation genes to study catabolic plasmid transfer in soil. A correlation between plasmid transfer to indigenous bacteria and augmented 2,4-D degradation was demonstrated in certain cases, emphasizing the efficacy of natural plasmid transfer, even when the original donor may not thrive in the new environment (Newby and Pepper, 2002; Newby et al., 2000a,b). The utilization of catabolic plasmid transfer to create hybrid strains capable of degrading an expanded range of aromatics has been reviewed extensively (Cao et al., 2009; Reineke, 1998; van der Meer et al., 1992). In many cases, plasmid-encoded genes are able to overcome metabolic blocks, allowing for complete degradation of a previously recalcitrant compound. This is generally known as the ‘‘patchwork’’ approach and combines enzymes from two or more existing pathways (Copley, 2000). The patchwork approach has been effectively utilized to produce strains capable of degrading a variety of recalcitrant aromatic compounds. To degrade monochlorobiphenyls, the ohb operon of Pseudomonas aeruginosa and the fcb operon of Arthrobacter globiformis was cloned and expressed in Comamonas testosterone sp. VP44 (Hrywna et al., 1999). Monti et al. (2005)

16


introduced genes involved in the degradation of 2,4-dinitrotoluene from Burkholderia sp. DNT into Pseudomonas fluorescens ATCC 17400, allowing for complete mineralization of the compound. Through transfer of the well-studied TOL plasmid pWWO, recombinant P. putida F38/D has been constructed which combines the tod and TOL pathways to degrade benzene, toluene, and p-xylene (Lee et al., 1994). A follow-up study created the hybrid strain P. putida TB105 which was also capable of degrading benzene, toluene, and p-xylene, this time through introduction of a plasmid which expressed tod pathway genes to augment the host-encoded TOL pathway (Lee et al., 1995). Analysis of natural isolates also makes it clear that a patchwork approach using plasmid-encoded genes is a broadly effective strategy for adaptation to aromatic compounds (Copley, 2000; Top and Springael, 2003).

b. Conjugative transposons and genomic islands Although class I and class II transposons (Springael and Top, 2004; Top and Springael, 2003) can move within a single host’s genome, they are typically incapable of horizontal transfer without the aid of a plasmid. Conjugative transposons are distinct from other classes in their ability to excise from host DNA, form a circular intermediate, and integrate into a new DNA target. This capability allows conjugative transposons to move laterally between hosts without the aid of plasmids (Tsuda et al., 1999). As with plasmid transfer, integration of conjugative transposons can rapidly expand the catabolic repertoire of the target bacterium. In the literature, there are several examples of catabolic genes associated with conjugative transposons. In Ralstonia oxalatica A5, a strain capable of metabolizing biphenyl and 4-chlorobiphenyl, the 13-kb bph region is located within a 55-kb transposon, Tn4371 (Merlin et al., 1997, 1999). Tn4371-like catabolic sequences have since been identified in other (chloro-)biphenyl-degrading bacteria and appear to encode expanded degradative capacity (Merlin et al., 1997; Springael et al., 2001). An additional example of a conjugative catabolic transposon is the bph-sal element of P. putida strain KF715. The bph-sal element encodes genes for biphenyl and salicylate metabolism and is located on a 90 kb chromosomal fragment. This element can be transferred to different P. putida strains—without the aid of a plasmid— at frequencies of approximately 1 10 6 per recipient cell (Nishi et al., 2000). Although they may be technically classified as conjugative transposons, large, self-transmissible regions of DNA such as Tn4371 contain specific features which allow them to be considered ‘‘genomic islands’’ (Ravatn et al., 1998; van der Meer and Sentchilo, 2003). In relation to aromatic degradation, the term ‘‘genomic island’’ has been proposed for


17

mobile regions of DNA, typically carrying genes for integration and conjugation, which encode catabolic proteins. As in the spread of pathogenic or antibiotic resistance ‘‘islands,’’ the transfer of catabolic genomic islands provides bacteria with a dynamic genetic resource for rapid adaptation and evolution ( Juhas et al., 2009). Perhaps the best known example is the clc-element which was originally identified in the 3-chlorobenzoate-degrading organism Pseudomonas sp. B13 (Gaillard et al., 2006; van der Meer et al., 2001). The clc-element comprises a 105 kb region of DNA which carries the clcRABD gene cluster. These genes are involved in the mineralization of chlorobenzoate and chlorocatechols via ortho-cleavage. In a variety of bacteria, the clc-element integrates site specifically into a tRNA gene, mediated by the P4-type integrase IntB13. Similar elements which carry identical catabolic genes exist in other chlorobenzoate-degrading bacteria which were independently isolated from geographically distinct locations (Gaillard et al., 2006). Interestingly, the clc-element has also been shown to undergo duplication events which were necessary for its benzene-degrading host to grow on chlorinated benzenes (van der Meer et al., 1998). This example further demonstrates the intertwining of both vertical and horizontal strategies for enhancing growth substrate range. Genomic islands related to the clc-element, but carrying different catabolic information such as that for biphenyl degradation (bph), have also been described (van der Meer and Sentchilo, 2003).

B. Expression of repair enzymes The expression of proper catabolic genes does not always ensure the efficient degradation of aromatic compounds. Indeed, aromatic compounds and their degradation byproducts can cause enzyme inactivation and, in some cases, lead to more generalized cellular toxicity. To deal with these deleterious effects, bacteria may express proteins which alleviate toxicity and allow for proficient aromatic metabolism. Below, we discuss two classes of proteins: those involved in enzyme reactivation and those which alleviate general cellular stress.

1. Enzyme reactivation As mentioned above (see Section II.C), metabolites formed during aromatic degradation have the capacity to inactivate catabolic enzymes. In the case of catecholic intermediates, inactivation of C23Os may occur directly during ring cleavage, in a process known as suicide inhibition (Bartels et al., 1984; Klecka and Gibson, 1981), or indirectly through the action of oxidative species (Vaillancourt et al., 2006). Suicide inactivation has been shown to limit the range of toluates degraded by the TOL pathway, polychlorinated biphenyls degraded by the bph pathway, and

18


chloro- and alkyl-benzenes degraded by the tod pathway (Cerdan et al., 1994; Klecka and Gibson, 1981; Ramos et al., 1987; Rojo et al., 1987; Vaillancourt et al., 2002; Ward et al., 2004). Given the crucial role of C23O inactivation in limiting substrate range, some bacteria have developed enzymatic ‘‘repair’’ systems to reverse enzyme inactivation. Collectively, these proteins are known as [2Fe–2S] plant-like ferredoxins. Plant-like ferredoxins catalyze a redox-dependent reaction, reducing oxidized Fe(III) to the catalytically active Fe(II) (Hugo et al., 1998). Perhaps the most well-studied [2Fe–2S] ferredoxin is XylT, which promotes reactivation of XylE, a C23O in the TOL plasmidencoded pathway of P. putida mt2. Mutants deprived of a functional xylT gene lost the ability to grow on both p-xylene and p-toluate as sole carbon sources (Polissi and Harayama, 1993). The impeded growth of xylT mutants, coupled with the observation that the intermediate 4methylcatechol irreversibly inactivated XylE only in mutant strains, led to the supposition that XylT functioned in C23O reactivation (Polissi and Harayama, 1993). Subsequent purification of XylT revealed that it was a [2Fe–2S] ferredoxin which reactivated XylE through reduction of the iron atom in the enzyme active site (Hugo et al., 1998). Further studies have revealed the existence of XylT-like [2Fe–2S] ferredoxins in other catabolic pathways including those for the degradation of naphthalene, cresols, nitrobenzene, aniline, and chlorobenzene (Herrmann et al., 1995; Hugo et al., 2000; Parales et al., 1997; Powlowski and Shingler, 1994; Tropel et al., 2002). The abundance of [2Fe–2S] ferredoxins in such a diverse array of pathways and bacteria underscores the efficacy of this particular strategy of substrate expansion. Through this reactivation strategy, bacteria are able to increase catechol turnover and permit growth on previously inaccessible substrates. Given catechols’ noted toxicity (see Section II.C), efficient catechol turnover is essential for the viability of cells producing it. Recently, it was shown that expression of the C23O DmpB, along with its endogenous ferredoxin DmpQ, reduced C23O inactivation and allowed P. putida F1 to acquire the ability to grow on styrene (George et al., 2011). Intriguingly, expression of DmpQ alone also protected against cell death, preventing marked declines in cell viability observed in wild-type F1 exposed to styrene, even though its expression was not sufficient to confer growth on styrene.

2. Reduction of cellular stress In addition to inactivating the enzymes responsible for their degradation, catechols can also exert toxicity directly through a variety of mechanisms such as membrane stress and redox cycling (Schweigert et al., 2001a,b). Catechols, however, are not the only intermediates which may be acutely toxic. Various hydroxylated intermediates and products of ring fission may also be toxic to bacterial cells. In addition to direct metabolite


19

toxicity, evidence has accumulated which suggests bacteria experience considerable oxidative stress during the degradation of many aromatic compounds (Imlay, 2003). During the degradation of polychlorinated biphenyls, for example, metabolism is accompanied by the production of ROS and the induction of cellular stress (Chavez et al., 2006). In some instances, the parent aromatic compound is also capable of exerting toxicity. This is the case for many hydrophobic polycyclic aromatic hydrocarbons as well as solvents such as toluene and styrene (Ramos et al., 2002). Growth in the presence of high concentrations of toluene and similar aromatic solvents is enhanced, for example, by the expression of efflux pumps or by alterations in membrane fluidity (Ramos et al., 2002). Importantly, one of the efflux pumps (TtgGHI) is plasmid encoded, further demonstrating the importance of horizontal gene transfer in adaption to growth on aromatic compounds (Rodrı´guez-Herva et al., 2007). The diversity of toxic insults accompanying aromatic exposure and degradation has required bacteria to develop numerous detoxification mechanisms. Some strains, for example, may utilize glutathione S-transferases (GSTs) which are broadly involved in cellular detoxification against harmful xenobiotics and can protect against oxidative stress (Allocati et al., 2009). With respect to the degradation of aromatic compounds, bacterial GSTs are involved in the degradation of monocyclic aromatics such as phenol (Santos et al., 2002) and polycyclic aromatics such as anthracene (Kim et al., 1997). A GST first characterized for its involvement in bph (Bartels et al., 1999; Hofer et al., 1994), BphK, is one of the most extensively studied bacterial GSTs and may have a role in multiple pathways (Allocati et al., 2009). In S. yanoikuyae B1, for instance, the presence of a bphK gene allows the organism to grow more efficiently on m-toluate (Bae et al., 2003). Through expression of GSTs such as BphK, bacteria can achieve efficient degradation of aromatic compounds that would otherwise preclude growth due to toxicity. More detail on GSTs and their role in aromatic degradation can be found in the extensive review of Allocati et al. (2009). As with biphenyl, naphthalene has been reported to be toxic to some bacteria (Ahn et al., 1998; Garcia et al., 1998; Park and Madsen, 2004; Pumphrey and Madsen, 2007) and its metabolism can lead to the production of ROS (Kang et al., 2006). Intriguingly, the addition of antioxidants such as ascorbate was found to reduce the inhibitory effects of ROS and increase biodegradation efficiency (Kang et al., 2006). In a follow-up study, it was found that overexpression of antioxidant enzymes enhanced naphthalene biodegradation in Pseudomonas sp. strain AS1 (Kang et al., 2007). Specifically, the authors found that expression of a ferredoxin– NADPþ reductase (Fpr) or superoxide dismutase (SOD) in tandem with alkyl hydroperoxide reductase (AhpC) conferred resistance to oxidative stress and simultaneously increased growth rate. These results show that

20


overexpression of antioxidant enzymes represents an effective bacterial strategy for enhancing biodegradation and are consistent with a proteomic analysis of P. putida KT2440 which demonstrated that the production of proteins involved in oxidative stress defense such as SOD and AhpC increased following pollutant exposure (Santos et al., 2004).

C. Direct modulation of substrate range or kinetic parameters While changes in enzyme substrate range or kinetics seem trivial—especially when compared to large-scale vertical gene duplications or horizontal transfer—they can play an important role in the development of new catabolic functions. These changes are usually the result of errors in DNA replication and repair systems, which occur at a low yet detectable rate (Kunkel, 2004). The low mutation rate is not constant, however, and can increase due to the induction of error-prone polymerases under stressed conditions (Bull et al., 2001; Foster, 2007; Goodman, 2002; Nohmi, 2006; Tegova et al., 2004), the formation of sequence-dependent secondary structures (Gerischer and Ornston, 1995; Hartnett et al., 1990; Wright, 2000), or the transient formation of single-stranded DNA during transcription (Wright, 2000). Point mutations, insertions, and small deletions formed during DNA replication and repair have been observed in nearly every aromatic catabolic pathway (van der Meer, 2002). The effects of these mutations on the properties of catabolic enzymes are generally well understood and several examples are discussed below.

1. Increased efficiency or substrate range In genes encoding catabolic enzymes for aromatic degradation, relatively simple base pair mutations are capable of altering substrate specificity or kinetic parameters (van der Meer et al., 1992). These subtle changes can have profound effects on biodegradation efficiency and, in some cases, result in expansion of growth substrate range. In laboratory systems, spontaneous modification of enzyme kinetic properties or substrate specificity has led to growth on new aromatic substrates. In the TOL plasmid pWWO of P. putida mt-2, cleavage of 4-ethylcatechol, an intermediate in 4ethylbenzoate metabolism, results in suicide inhibition of the C23O XylE. When selected on 4-ethylbenzoate, however, spontaneous mutants were isolated with substitutions in XylE which rendered it resistant to inhibition by 4-ethylcatechol and in turn allowed mt-2 to grow on 4-ethylbenzoate (Ramos et al., 1987). A later study further characterized two spontaneous C23O mutants resistant to 4-ethylcatechol-mediated inactivation and developed a quantitative relationship between C23O inactivation resistance and cell growth rate (Cerdan et al., 1994). In other pathways subject to C23O inhibition such as that for bph, similar mutations have produced ring cleaving dioxygenases resistant to suicide inactivation by


21

the relevant catechols (Vaillancourt et al., 2005). This expansion of growth substrate specificity by simple base pair mutations was also observed in the TOL plasmid upper pathway. Mutants were isolated which contained substitutions in toluene oxidase that expanded the substrate range to include p-ethyltoluene (Abril et al., 1989). Additional examples of mutations that expand substrate range by increasing the rate of substrate transformation are common in the scientific literature and have been reviewed elsewhere (Khomenkov et al., 2008; van der Meer et al., 1992). In recent years, there has been considerable interest in the directed evolution of catabolic proteins for use in bioremediation or biocatalysis (Diaz, 2004; Timmis and Pieper, 1999; Urgun-Demirtas et al., 2006; Yuan et al., 2005). Much of this work has been based on information gathered from spontaneous mutants as described above. Using a variety of techniques, significant progress has been made in the generation of catalytically improved biphenyl (Furukawa et al., 2004; Kumamaru et al., 1998; Suenaga et al., 2001), naphthalene (Fortin et al., 2005; Gibson and Parales, 2000; Parales et al., 1998), and dioxin dioxygenases (Furukawa, 2000). Other catabolic proteins involved in aromatic degradation such as C23Os have also been successfully engineered with improved catalytic efficiency or broadened substrate range (Cao et al., 2009).

2. Catabolic enzyme inactivation If the activity of a particular enzyme is inhibitory to growth, studies have shown that the isolation of mutants with abolished enzyme activity is readily accomplished. This has been clearly illustrated in cases where normal enzyme function produces a toxic metabolite or results in pathway misrouting. In the metabolism of p-hydroxybenzoate by Acinetobacter calcoaceticus, for example, protocatechuate 3,4-dioxygenase (PcaGH) generates the toxic intermediate b-carboxymuconate. When an A. calcoaceticus strain was exposed to physiological conditions where expression of PcaGH inhibited growth, Gerischer and Ornston (1995) were able to isolate numerous mutants where base substitutions in pcaG or pcaH eliminated PcaGH activity. These mutants failed to cleave protocatechuate and thus did not accumulate b-carboxymuconate. In other isolates, abolition of PcaGH activity was achieved through lengthy deletions or gene interruption by IS1236, an insertion element. Specifically, the authors implicated DNA slippage structures and subsequent mispairing as the mechanism behind increased mutation rates (Gerischer and Ornston, 1995). In natural isolates, there is also evidence for the inactivation of deleterious enzyme functions. Pseudomonas sp. P51, a strain capable of degrading chlorobenzene, contains a mosaic pathway with a chlorobenzene dioxygenase (encoded by tcbA) and a dehydrogenase (encoded by tcbB) related to enzymes involved in toluene (todC1C2BAD) or benzene

22


degradation (bedC1C2BA) (Werlen et al., 1996). A large gene fragment encoding a nonfunctional meta-cleavage dioxygenase homologous to todE was also found downstream of tcbB. The authors suggested that this gene fragment, along with one similar to todF located upstream of tcbA, were evolutionary remnants of the recombination event which eventually resulted in the capture of tcbA and tcbB. In chlorobenzene degradation, the ortho-cleavage pathway produces intermediates that can support growth, while the meta-pathway is unproductive and typically induces toxicity (Werlen et al., 1996). Since the todE-like gene was deleterious to the cell, it may have been necessary for the organisms to eliminate meta-cleavage activity to prevent pathway misrouting and toxicity. In another isolate capable of chlorobenzene degradation, Burkholderia sp. strain PS12, a gene encoding a meta-cleaving dioxygenase, was similarly inactivated (Beil et al., 1999). Intriguingly, the genomic context of the chlorobenzene degrading tec genes in strain PS12 was very similar to that of Pseudomonas sp. strain P51 in that it still retained todF and todE-like remnants. In this case, however, simple point mutations inactivated the meta-cleavage enzyme rather than a deletion. Although strains PS12 and P51 used different molecular mechanisms, the general strategy of enzymatic inactivation proved equally effective. Inactivation of genes responsible for metabolite misrouting has also been observed in several other isolates, such as those involved in the degradation of nitroaromatic compounds ( Ju and Parales, 2009).

3. ‘‘Less is more’’ The efficient turnover of catechol intermediates in aromatic degradation pathways is critical due to their toxicity and unique ability to inactivate the dioxygenases responsible for their cleavage (see Section II.C). Since catechol production is required for productive aromatic metabolism in many pathways, enzyme inactivation as described above is not always possible. Recently, a novel strategy for the reduction of catechol-induced toxicity was described in P. putida F1, the model organism harboring the tod pathway. P. putida F1 is unable to grow on styrene due to the deleterious effects of 3-vinylcatechol, an intermediate in degradation. When exposed to styrene, 3-vinylcatechol inactivates the C23O TodE and subsequently accumulates, causing marked reductions in cell viability (George et al., 2011) (Fig. 1.6). When styrene-naı¨ve F1 cells were exposed to styrene as a sole carbon source, mutants were eventually isolated which did not accumulate 3-vinylcatechol and could grow on styrene (George, 2011). Characterization of one mutant, designated SF1, revealed the presence of a single base pair mutation (C479T) in the reductase component (todA) of toluene dioxygenase (TDO) (todC1C2BA), the enzyme responsible for the production of 3-vinylcatechol from styrene (George, 2011). Further analysis revealed that this simple mutation resulted in attenuated TDO activity and a decreased rate of 3-vinylcatechol production. SF1’s todA allele


A

23

Wild type R

R O

R OH Lower

Upper

COO– OH

OH B

Increased operon transcription (single promoter) R

Upper

Lower

R

R O

OH COO– OH

OH

FIGURE 1.6 Nonproductive scenarios for growth on substrates which produce C23Oinactivating catechols. (A) The rapid production of catechol by the upper pathway inactivates C23O activity (blocked curved line) and results in catechol accumulation and toxicity (skull and bones). (B) Since the tod pathway (and many other catabolic pathways) is transcribed from a single promoter, upregulating transcription may have no effect on net catechol turnover. Increased upper pathway activity produces greater amounts of catechol and amplifies C23O inactivation, negating the effect of increased C23O activity (thicker curved line).

(todAC479T) was then expressed in styrene-naı¨ve F1 and found to be sufficient to permit growth on styrene, whereas expression of a wildtype copy of todA in SF1 reduced growth on styrene and led to 3-vinylcatechol accumulation. Unlike previously discussed strains that adapted to toxic catechol intermediates by developing resistant C23Os (Section III. C.1), SF1’s C23O activity and substrate range remained unchanged. As this particular strategy relies on a decrease rather than an increase in enzyme activity, theoretically, a large number of mutations could have resulted in reduced catechol production. Indeed, characterization of four additional styrene-adapted mutants revealed the presence of reduced TDO activity in all strains despite the absence of the mutant todA allele of SF1 (George, 2011). The net effect of decreasing 3-vinylcatechol production via this decrease in TDO activity was similar to that observed when F1’s C23O todE was overexpressed (George et al., 2011): no 3-vinylcatechol accumulated and styrene supported growth. These observations are suggestive of an alternative ‘‘less is more’’ strategy and demonstrate that there multiple ways for cells to protect themselves from the accumulation of potential

24


A Selective enhancement of lower pathway

R Upper

Lower

R

R O

OH COO– OH

OH B C230 reactivation

R Upper

R O

R OH Lower

COO– OH

OH C

[2Fe–2S] Ferredoxin reactivates oxidized C230

“Less is more”

R

R Upper

R O OH Lower COO– OH

OH

FIGURE 1.7 Productive scenarios for growth on substrates which produce C23Oinactivating catechols. (A) Although some inactivation may still take place (curved line), selective enhancement of the lower pathway increases catechol turnover and prevents total C23O inactivation. This can be accomplished through changes in gene dosage. Alternatively, it can also be achieved through mutations which produce more efficient C23Os, or those which are less susceptible to inactivation. (B) Some pathways make use of [2Fe–2S] ferredoxins to reactivate C23Os affected by catechol (curved line replaced with circular arrow). This increases catechol turnover and supports growth. (C) The ‘‘less is more’’ strategy employed by SF1 is achieved through reduced upper pathway activity which decreases catechol production and subsequent C23O inactivation (thinner curved line), thereby allowing for growth.

deleterious catechols (George, 2011) (Fig. 1.7). Given the ubiquity of catechol intermediates in aromatic degradation, it is likely that this strategy for alleviating catechol toxicity may also be effective in other catabolic pathways.

IV. CONCLUSION Microorganisms possess a wide range of genetic mechanisms and strategies that allow for adaptation to environmental changes. Current knowledge summarized in this review suggests that the endogenous genetic potential of a bacterium, combined with gene transfer and mutation,


25

allows for the evolution of bacteria capable of growth on vast array aromatic substrates. As some estimates suggest that we have characterized less than 5% of microbial diversity (Curtis et al., 2002), it is likely that continued investigations will reveal as yet undiscovered aromatic degradation pathways and provide insight into additional novel mechanisms of adaptation.

REFERENCES Abril, M. A., Michan, C., Timmis, K. N., and Ramos, J. L. (1989). Regulator and enzyme specificities of the TOL plasmid-encoded upper pathway for degradation of aromatic hydrocarbons and expansion of the substrate range of the pathway. J. Bacteriol. 171, 6782–6790. Agency for Toxic Substances and Disease Registry (ATSDR) (2007). Toxicological Profile for Benzene. U.S. Department of Health and Human Services, Washington, DC. Ahn, I. S., Ghiorse, W. C., Lion, L. W., and Shuler, M. L. (1998). Growth kinetics of Pseudomonas putida G7 on naphthalene and occurrence of naphthalene toxicity during nutrient deprivation. Biotechnol. Bioeng. 59, 587–594. Allocati, N., Federici, L., Masulli, M., and Di Ilio, C. (2009). Glutathione transferases in bacteria. FEBS J. 276, 58–75. Arai, H., Akahira, S., Ohishi, T., and Kudo, T. (1999a). Adaptation of Comamonas testosteroni TA441 to utilization of phenol by spontaneous mutation of the gene for a trans-acting factor. Mol. Microbiol. 33, 1132–1140. Arai, H., Yamamoto, T., Ohishi, T., Shimizu, T., Nakata, T., and Kudo, T. (1999b). Genetic organization and characteristics of the 3-(3-hydroxyphenyl)propionic acid degradation pathway of Comamonas testosteroni TA441. Microbiology 145(Pt. 10), 2813–2820. Armengaud, J., Happe, B., and Timmis, K. N. (1998). Genetic analysis of dioxin dioxygenase of Sphingomonas sp. Strain RW1: Catabolic genes dispersed on the genome. J. Bacteriol. 180, 3954–3966. Bae, M., Sul, W. J., Koh, S. C., Lee, J. H., Zylstra, G. J., Kim, Y. M., and Kimr, E. (2003). Implication of two glutathione S-transferases in the optimal metabolism of m-toluate by Sphingomonas yanoikuyae B1. Antonie Van Leeuwenhoek 84, 25–30. Bartels, I., Knackmuss, H. J., and Reineke, W. (1984). Suicide inactivation of catechol 2,3dioxygenase from Pseudomonas putida mt-2 by 3-halocatechols. Appl. Environ. Microbiol. 47, 500–505. Bartels, F., Backhaus, S., Moore, E. R., Timmis, K. N., and Hofer, B. (1999). Occurrence and expression of glutathione-S-transferase-encoding bphK genes in Burkholderia sp. strain LB400 and other biphenyl-utilizing bacteria. Microbiology 145(Pt. 10), 2821–2834. Beil, S., Timmis, K. N., and Pieper, D. H. (1999). Genetic and biochemical analyses of the tec operon suggest a route for evolution of chlorobenzene degradation genes. J. Bacteriol. 181, 341–346. Bull, H. J., Lombardo, M. J., and Rosenberg, S. M. (2001). Stationary-phase mutation in the bacterial chromosome: Recombination protein and DNA polymerase IV dependence. Proc. Natl. Acad. Sci. USA 98, 8334–8341. Busch, A., Lacal, J., Martos, A., Ramos, J. L., and Krell, T. (2007). Bacterial sensor kinase TodS interacts with agonistic and antagonistic signals. Proc. Natl. Acad. Sci. USA 104, 13774–13779. Butler, C. S., and Mason, J. R. (1997). Structure–function analysis of the bacterial aromatic ring-hydroxylating dioxygenases. Adv. Microb. Physiol. 38, 47–84.

26


Cao, B., Nagarajan, K., and Loh, K. C. (2009). Biodegradation of aromatic compounds: Current status and opportunities for biomolecular approaches. Appl. Microbiol. Biotechnol. 85, 207–228. Cases, I., and de Lorenzo, V. (2001). The black cat/white cat principle of signal integration in bacterial promoters. EMBO J. 20, 1–11. Cerdan, P., Wasserfallen, A., Rekik, M., Timmis, K. N., and Harayama, S. (1994). Substratespecificity of catechol 2,3-dioxygenase encoded by TOL plasmid PWWO of Pseudomonas putida and its relationship to cell-growth. J. Bacteriol. 176, 6074–6081. Chavez, F. P., Gordillo, F., and Jerez, C. A. (2006). Adaptive responses and cellular behaviour of biphenyl-degrading bacteria toward polychlorinated biphenyls. Biotechnol. Adv. 24, 309–320. Choi, E. N., Cho, M. C., Kim, Y., Kim, C. K., and Lee, K. (2003). Expansion of growth substrate range in Pseudomonas putida F1 by mutations in both cymR and todS, which recruit a ringfission hydrolase CmtE and induce the tod catabolic operon, respectively. Microbiology 149, 795–805. Cirino, P. C., and Arnold, F. H. (2002). Protein engineering of oxygenases for biocatalysis. Curr. Opin. Chem. Biol. 6, 130–135. Copley, S. D. (2000). Evolution of a metabolic pathway for degradation of a toxic xenobiotic: The patchwork approach. Trends Biochem. Sci. 25, 261–265. Curtis, T. P., Sloan, W. T., and Scannell, J. W. (2002). Estimating prokaryotic diversity and its limits. Proc. Natl. Acad. Sci. USA 99, 10494–10499. Dejonghe, W., Goris, J., El Fantroussi, S., Hofte, M., De Vos, P., Verstraete, W., and Top, E. M. (2000). Effect of dissemination of 2,4-dichlorophenoxyacetic acid (2,4-D) degradation plasmids on 2,4-D degradation and on bacterial community structure in two different soil horizons. Appl. Environ. Microbiol. 66, 3297–3304. Delgado, A., Salto, R., Marques, S., and Ramos, J. L. (1995). Single amino acids changes in the signal receptor domain of XylR resulted in mutants that stimulate transcription in the absence of effectors. J. Biol. Chem. 270, 5144–5150. Dennis, J. J. (2005). The evolution of IncP catabolic plasmids. Curr. Opin. Biotechnol. 16, 291–298. Diaz, E. (2004). Bacterial degradation of aromatic pollutants: A paradigm of metabolic versatility. Int. Microbiol. 7, 173–180. Diaz, E., and Prieto, M. A. (2000). Bacterial promoters triggering biodegradation of aromatic pollutants. Curr. Opin. Biotechnol. 11, 467–475. Dominguez-Cuevas, P., Marin, P., Busby, S., Ramos, J. L., and Marques, S. (2008). Roles of effectors in XylS-dependent transcription activation: Intramolecular domain derepression and DNA binding. J. Bacteriol. 190, 3118–3128. Eaton, R. W. (1996). p-Cumate catabolic pathway in Pseudomonas putida Fl: Cloning and characterization of DNA carrying the cmt operon. J. Bacteriol. 178, 1351–1362. Eaton, R. W. (1997). p-Cymene catabolic pathway in Pseudomonas putida F1: Cloning and characterization of DNA encoding conversion of p-cymene to p-cumate. J. Bacteriol. 179, 3171–3180. Fortin, P. D., MacPherson, I., Neau, D. B., Bolin, J. T., and Eltis, L. D. (2005). Directed evolution of a ring-cleaving dioxygenase for polychlorinated biphenyl degradation. J. Biol. Chem. 280, 42307–42314. Foster, P. L. (2007). Stress-induced mutagenesis in bacteria. Crit. Rev. Biochem. Mol. Biol. 42, 373–397. Furukawa, K. (2000). Engineering dioxygenases for efficient degradation of environmental pollutants. Curr. Opin. Biotechnol. 11, 244–249. Furukawa, K., Suenaga, H., and Goto, M. (2004). Biphenyl dioxygenases: Functional versatilities and directed evolution. J. Bacteriol. 186, 5189–5196.


27

Furusawa, Y., Nagarajan, V., Tanokura, M., Masai, E., Fukuda, M., and Senda, T. (2004). Crystal structure of the terminal oxygenase component of biphenyl dioxygenase derived from Rhodococcus sp. strain RHA1. J. Mol. Biol. 342, 1041–1052. Gaillard, M., Vallaeys, T., Vorholter, F. J., Minoia, M., Werlen, C., Sentchilo, V., Puhler, A., and van der Meer, J. R. (2006). The clc element of Pseudomonas sp. Strain B13, a genomic island with various catabolic properties. J. Bacteriol. 188, 1999–2013. Garcia, E. M., Siegert, I. G., and Suarez, P. (1998). Toxicity assays and naphthalene utilization by natural bacteria selected in marine environments. Bull. Environ. Contam. Toxicol. 61, 370–377. George, K. W. (2011). Dissertation. Adaptation of Pseudomonas putida F1 to growth on styrene Cornell University. George, K. W., Kagle, J., Junker, L., Risen, A., and Hay, A. G. (2011). Growth of Pseudomonas putida F1 on styrene requires increased catechol-2,3-dioxygenase activity, not a new hydrolase. Microbiology. 157, 89–98. Gerischer, U., and Ornston, L. N. (1995). Spontaneous mutations in pcaH and -G, structural genes for protocatechuate 3,4-dioxygenase in Acinetobacter calcoaceticus. J. Bacteriol. 177, 1336–1347. Gibson, D. T., and Parales, R. E. (2000). Aromatic hydrocarbon dioxygenases in environmental biotechnology. Curr. Opin. Biotechnol. 11, 236–243. Gibson, D. T., and Subramanian, V. (1984). Microbial degradation of aromatic hydrocarbons. In ‘‘Microbial Degradation of Organic Compounds’’ (D. T. Gibson, Ed.), pp. 181–252. Marcel Dekker, Inc., New York. Goodman, M. F. (2002). Error-prone repair DNA polymerases in prokaryotes and eukaryotes. Annu. Rev. Biochem. 71, 17–50. Harayama, S., and Rekik, M. (1989). Bacterial aromatic ring-cleavage enzymes are classified into two different gene families. J. Biol. Chem. 264, 15328–15333. Harayama, S., Kok, M., and Neidle, E. L. (1992). Functional and evolutionary relationships among diverse oxygenases. Annu. Rev. Microbiol. 46, 565–601. Hartnett, C., Neidle, E. L., Ngai, K. L., and Ornston, L. N. (1990). DNA sequences of genes encoding Acinetobacter calcoaceticus protocatechuate 3,4-dioxygenase: Evidence indicating shuffling of genes and of DNA sequences within genes during their evolutionary divergence. J. Bacteriol. 172, 956–966. Harwood, C. S., and Parales, R. E. (1996). The beta-ketoadipate pathway and the biology of self-identity. Annu. Rev. Microbiol. 50, 553–590. Hazen, T. C., Dubinsky, E. A., Desantis, T. Z., Andersen, G. L., Piceno, Y. M., Singh, N., Jansson, J. K., Probst, A., Borglin, S. E., Fortney, J. L., Stringfellow, W. T., Bill, M., et al. (2010). Deep-sea oil plume enriches indigenous oil-degrading bacteria. Science 330, 204–208. Herrick, J. B., Stuart-Keil, K. G., Ghiorse, W. C., and Madsen, E. L. (1997). Natural horizontal transfer of a naphthalene dioxygenase gene between bacteria native to a coal tar-contaminated field site. Appl. Environ. Microbiol. 63, 2330–2337. Herrmann, H., Muller, C., Schmidt, I., Mahnke, J., Petruschka, L., and Hahnke, K. (1995). Localization and organization of phenol degradation genes of Pseudomonas putida strain H. Mol. Gen. Genet. 247, 240–246. Hofer, B., Backhaus, S., and Timmis, K. N. (1994). The biphenyl polychlorinated biphenyldegradation locus (Bph) of Pseudomonas sp. LB400 encodes 4 additional metabolic enzymes. Gene 144, 9–16. Horvath, R. S. (1970). Co-metabolism of methyl- and chloro-substituted catechols by an Achromobacter sp. possessing a new meta-cleaving oxygenase. Biochem. J. 119, 871–876. Hrywna, Y., Tsoi, T. V., Maltseva, O. V., Quensen, J. F., 3rd, and Tiedje, J. M. (1999). Construction and characterization of two recombinant bacteria that grow on ortho- and para-substituted chlorobiphenyls. Appl. Environ. Microbiol. 65, 2163–2169.

28


Hugo, N., Armengaud, J., Gaillard, J., Timmis, K. N., and Jouanneau, Y. (1998). A novel [2Fe2S] ferredoxin from Pseudomonas putida mt2 promotes the reductive reactivation of catechol 2,3-dioxygenase. J. Biol. Chem. 273, 9622–9629. Hugo, N., Meyer, C., Armengaud, J., Gaillard, J., Timmis, K. N., and Jouanneau, Y. (2000). Characterization of three XylT-like [2Fe-2S] ferredoxins associated with catabolism of cresols or naphthalene: Evidence for their involvement in catechol dioxygenase reactivation. J. Bacteriol. 182, 5580–5585. Hyne, N. J. (2001). Nontechnical Guide to Petroleum Geology, Exploration, Drilling, and Production. Penwell Corporation, Oklahoma. Imlay, J. A. (2003). Pathways of oxidative damage. Annu. Rev. Microbiol. 57, 395–418. Jaspers, M. C., Suske, W. A., Schmid, A., Goslings, D. A., Kohler, H. P., and van der Meer, J. R. (2000). HbpR, a new member of the XylR/DmpR subclass within the NtrC family of bacterial transcriptional activators, regulates expression of 2-hydroxybiphenyl metabolism in Pseudomonas azelaica HBP1. J. Bacteriol. 182, 405–417. Ju, K. S., and Parales, R. E. (2009). Nitroaromatic compounds, from synthesis to biodegradation. Microbiol. Mol. Biol. Rev. 74, 250–272. Juhas, M., van der Meer, J. R., Gaillard, M., Harding, R. M., Hood, D. W., and Crook, D. W. (2009). Genomic islands: Tools of bacterial horizontal gene transfer and evolution. FEMS Microbiol. Rev. 33, 376–393. Kallastu, A., Horak, R., and Kivisaar, M. (1998). Identification and characterization of IS1411, a new insertion sequence which causes transcriptional activation of the phenol degradation genes in Pseudomonas putida. J. Bacteriol. 180, 5306–5312. Kang, Y. S., Kim, Y. J., Jeon, C. O., and Park, W. (2006). Characterization of naphthalenedegrading Pseudomonas species isolated from pollutant-contaminated sites: Oxidative stress during their growth on naphthalene. J. Microbiol. Biotechnol. 16, 1819–1825. Kang, Y. S., Lee, Y., Jung, H., Jeon, C. O., Madsen, E. L., and Park, W. (2007). Overexpressing antioxidant enzymes enhances naphthalene biodegradation in Pseudomonas sp. strain As1. Microbiology 153, 3246–3254. Kasak, L., Horak, R., Nurk, A., Talvik, K., and Kivisaar, M. (1993). Regulation of the catechol 1,2-dioxygenase- and phenol monooxygenase-encoding pheBA operon in Pseudomonas putida PaW85. J. Bacteriol. 175, 8038–8042. Kasak, L., Horak, R., and Kivisaar, M. (1997). Promoter-creating mutations in Pseudomonas putida: A model system for the study of mutation in starving bacteria. Proc. Natl. Acad. Sci. USA 94, 3134–3139. Khajamohiddin, S., Repalle, E. R., Pinjari, A. B., Merrick, M., and Siddavattam, D. (2008). Biodegradation of aromatic compounds: An overview of meta-fission product hydrolases. Crit. Rev. Microbiol. 34, 13–31. Khomenkov, V. G., Shevelev, A. B., Zhukov, V. G., Zagustina, N. A., Bezborodov, A. M., and Popov, V. O. (2008). Organization of metabolic pathways and molecular-genetic mechanisms of xenobiotic degradation in microorganisms: A review. Appl. Biochem. Microbiol. 44, 117–135. Kim, E., and Zylstra, G. J. (1999). Functional analysis of genes involved in biphenyl, naphthalene, phenanthrene, and m-xylene degradation by Sphingomonas yanoikuyae B1. J. Ind. Microbiol. Biotechnol. 23, 294–302. Kim, E., Zylstra, G. J., Freeman, J. P., Heinze, T. M., Deck, J., and Cerniglia, C. E. (1997). Evidence for the role of 2-hydroxychromene-2-carboxylate isomerase in the degradation of anthracene by Sphingomonas yanoikuyae B1. FEMS Microbiol. Lett. 153, 479–484. Klecka, G. M., and Gibson, D. T. (1981). Inhibition of catechol 2,3-dioxygenase from Pseudomonas putida by 3-chlorocatechol. Appl. Environ. Microbiol. 41, 1159–1165. Koh, S., McCullar, M. V., and Focht, D. D. (1997). Biodegradation of 2,4-dichlorophenol through a distal meta-fission pathway. Appl. Environ. Microbiol. 63, 2054–2057.


29

Kohler, H. P., Kohler-Staub, D., and Focht, D. D. (1988). Degradation of 2-hydroxybiphenyl and 2,20 -dihydroxybiphenyl by Pseudomonas sp. strain HBP1. Appl. Environ. Microbiol. 54, 2683–2688. Kohler, H. P., van der Maarel, M. J., and Kohler-Staub, D. (1993). Selection of Pseudomonas sp. strain HBP1 Prp for metabolism of 2-propylphenol and elucidation of the degradative pathway. Appl. Environ. Microbiol. 59, 860–866. Kumamaru, T., Suenaga, H., Mitsuoka, M., Watanabe, T., and Furukawa, K. (1998). Enhanced degradation of polychlorinated biphenyls by directed evolution of biphenyl dioxygenase. Nat. Biotechnol. 16, 663–666. Kunkel, T. A. (2004). DNA replication fidelity. J. Biol. Chem. 279, 16895–16898. Lacal, J., Busch, A., Guazzaroni, M. E., Krell, T., and Ramos, J. L. (2006). The TodS-TodT twocomponent regulatory system recognizes a wide range of effectors and works with DNAbending proteins. Proc. Natl. Acad. Sci. USA 103, 8191–8196. Lacal, J., Guazzaroni, M. E., Busch, A., Krell, T., and Ramos, J. L. (2008). Hierarchical binding of the TodT response regulator to its multiple recognition sites at the tod pathway operon promoter. J. Mol. Biol. 376, 325–337. Laemmli, C. M., Leveau, J. H. J., Zehnder, A. J. B., and van der Meer, J. R. (2000). Characterization of a second tfd gene cluster for chlorophenol and chlorocatechol metabolism on plasmid pJP4 in Ralstonia eutropha JMP134(pJP4). J. Bacteriol. 182, 4165–4172. Lau, P. C., Wang, Y., Patel, A., Labbe, D., Bergeron, H., Brousseau, R., Konishi, Y., and Rawlings, M. (1997). A bacterial basic region leucine zipper histidine kinase regulating toluene degradation. Proc. Natl. Acad. Sci. USA 94, 1453–1458. Lee, J. Y., Roh, J. R., and Kim, H. S. (1994). Metabolic engineering of Pseudomonas putida for the simultaneous biodegradation of benzene, toluene, and p-xylene mixture. Biotechnol. Bioeng. 43, 1146–1152. Lee, J. Y., Jung, K. H., Choi, S. H., and Kim, H. S. (1995). Combination of the tod and the tol pathways in redesigning a metabolic route of Pseudomonas putida for the mineralization of a benzene, toluene, and p-xylene mixture. Appl. Environ. Microbiol. 61, 2211–2217. Lee, K., Ryu, E. K., Choi, K. S., Cho, M. C., Jeong, J. J., Choi, E. N., Lee, S. O., Yoon, D. Y., Hwang, I., and Kim, C. K. (2006). Identification and expression of the cym, cmt, and tod catabolic genes from Pseudomonas putida KL47: Expression of the regulatory todST genes as a factor for catabolic adaptation. J. Microbiol. 44, 192–199. Mahillon, J., and Chandler, M. (1998). Insertion sequences. Microbiol. Mol. Biol. Rev. 62, 725–774. Mason, J. R., and Cammack, R. (1992). The electron-transport proteins of hydroxylating bacterial dioxygenases. Annu. Rev. Microbiol. 46, 277–305. Merlin, C., Springael, D., Mergeay, M., and Toussaint, A. (1997). Organisation of the bph gene cluster of transposon Tn4371, encoding enzymes for the degradation of biphenyl and 4-chlorobiphenyl compounds. Mol. Gen. Genet. 253, 499–506. Merlin, C., Springael, D., and Toussaint, A. (1999). Tn4371: A modular structure encoding a phage-like integrase, a Pseudomonas-like catabolic pathway, and RP4/Ti-like transfer functions. Plasmid 41, 40–54. Meulien, P., Downing, R. G., and Broda, P. (1981). Excision of the 40 kb segment of the TOL plasmid from Pseudomonas putida mt-2 involves direct repeats. Mol. Gen. Genet. 184, 97–101. Milennium Assessment (2005). Ecosystems and Human Well-Being: Synthesis. Island Press, Washington, DC. Monti, M. R., Smania, A. M., Fabro, G., Alvarez, M. E., and Argarana, C. E. (2005). Engineering Pseudomonas fluorescens for biodegradation of 2,4-dinitrotoluene. Appl. Environ. Microbiol. 71, 8864–8872. Muller, T. A., Werlen, C., Spain, J., and van der Meer, J. R. (2003). Evolution of a chlorobenzene degradative pathway among bacteria in a contaminated groundwater mediated by a genomic island in Ralstonia. Environ. Microbiol. 5, 163–173.

30


Newby, D. T., and Pepper, I. L. (2002). Dispersal of plasmid pJP4 in unsaturated and saturated 2,4-dichlorophenoxyacetic acid contaminated soil. FEMS Microbiol. Ecol. 39, 157–164. Newby, D. T., Gentry, T. J., and Pepper, I. L. (2000a). Comparison of 2,4-dichlorophenoxyacetic acid degradation and plasmid transfer in soil resulting from bioaugmentation with two different pJP4 donors. Appl. Environ. Microbiol. 66, 3399–3407. Newby, D. T., Josephson, K. L., and Pepper, I. L. (2000b). Detection and characterization of plasmid pJP4 transfer to indigenous soil bacteria. Appl. Environ. Microbiol. 66, 290–296. Nishi, A., Tominaga, K., and Furukawa, K. (2000). A 90-kilobase conjugative chromosomal element coding for biphenyl and salicylate catabolism in Pseudomonas putida KF715. J. Bacteriol. 182, 1949–1955. Nohmi, T. (2006). Environmental stress and lesion-bypass DNA polymerases. Annu. Rev. Microbiol. 60, 231–253. Nurk, A., Tamm, A., Horak, R., and Kivisaar, M. (1993). In-vivo-generated fusion promoters in Pseudomonas putida. Gene 127, 23–29. Ohta, Y., Maeda, M., and Kudo, T. (2001). Pseudomonas putida CE2010 can degrade biphenyl by a mosaic pathway encoded by the tod operon and cmtE, which are identical to those of P. putida F1 except for a single base difference in the operator-promoter region of the cmt operon. Microbiology 147, 31–41. Parales, R. E., Ontl, T. A., and Gibson, D. T. (1997). Cloning and sequence analysis of a catechol 2,3-dioxygenase gene from the nitrobenzene-degrading strain Comamonas sp. JS765. J. Ind. Microbiol. Biotechnol. 19, 385–391. Parales, R. E., Emig, M. D., Lynch, N. A., and Gibson, D. T. (1998). Substrate specificities of hybrid naphthalene and 2,4-dinitrotoluene dioxygenase enzyme systems. J. Bacteriol. 180, 2337–2344. Park, W., and Madsen, E. L. (2004). Characterization in Pseudomonas putida Cg1 of nahR and its role in bacterial survival in soil. Appl. Microbiol. Biotechnol. 66, 209–216. Park, D. W., Chae, J. C., Kim, Y., Iida, T., Kudo, T., and Kim, C. K. (2002). Chloroplast-type ferredoxin involved in reactivation of catechol 2,3-dioxygenase from Pseudomonas sp. S 47. J. Biochem. Mol. Biol. 35, 432–436. Park, W., Jeon, C. O., Cadillo, H., DeRito, C., and Madsen, E. L. (2004). Survival of naphthalene-degrading Pseudomonas putida NCIB 9816-4 in naphthalene-amended soils: Toxicity of naphthalene and its metabolites. Appl. Microbiol. Biotechnol. 64, 429–435. Pavel, H., Forsman, M., and Shingler, V. (1994). An aromatic effector specificity mutant of the transcriptional regulator DmpR overcomes the growth constraints of Pseudomonas sp. strain CF600 on para-substituted methylphenols. J. Bacteriol. 176, 7550–7557. Perez-Pantoja, D., Ledger, T., Pieper, D. H., and Gonzalez, B. (2003). Efficient turnover of chlorocatechols is essential for growth of Ralstonia eutropha JMP134(pJP4) in 3-chlorobenzoic acid. J. Bacteriol. 185, 1534–1542. Peters, M., Heinaru, E., Talpsep, E., Wand, H., Stottmeister, U., Heinaru, A., and Nurk, A. (1997). Acquisition of a deliberately introduced phenol degradation operon, pheBA, by different indigenous Pseudomonas species. Appl. Environ. Microbiol. 63, 4899–4906. Pieper, D. H., and Reineke, W. (2000). Engineering bacteria for bioremediation. Curr. Opin. Biotechnol. 11, 262–270. Polissi, A., and Harayama, S. (1993). In vivo reactivation of catechol 2,3-dioxygenase mediated by a chloroplast-type ferredoxin—A bacterial strategy to expand the substrate-specificity of aromatic degradative pathways. EMBO J. 12, 3339–3347. Powlowski, J., and Shingler, V. (1994). Genetics and biochemistry of phenol degradation by Pseudomonas sp. CF600. Biodegradation 5, 219–236. Pumphrey, G. M., and Madsen, E. L. (2007). Naphthalene metabolism and growth inhibition by naphthalene in Polaromonas naphthalenivorans strain CJ2. Microbiology 153, 3730–3738.


31

Ramos, J. L., Stolz, A., Reineke, W., and Timmis, K. N. (1986). Altered effector specificities in regulators of gene expression: TOL plasmid xylS mutants and their use to engineer expansion of the range of aromatics degraded by bacteria. Proc. Natl. Acad. Sci. USA 83, 8467–8471. Ramos, J. L., Wasserfallen, A., Rose, K., and Timmis, K. N. (1987). Redesigning metabolic routes: Manipulation of TOL plasmid pathway for catabolism of alkylbenzoates. Science 235, 593–596. Ramos, J. L., Marques, S., and Timmis, K. N. (1997). Transcriptional control of the Pseudomonas TOL plasmid catabolic operons is achieved through an interplay of host factors and plasmid-encoded regulators. Annu. Rev. Microbiol. 51, 341–373. Ramos, J. L., Duque, E., Gallegos, M. T., Godoy, P., Ramos-Gonzalez, M. I., Rojas, A., Teran, W., and Segura, A. (2002). Mechanisms of solvent tolerance in gram-negative bacteria. Annu. Rev. Microbiol. 56, 743–768. Ravatn, R., Studer, S., Springael, D., Zehnder, A. J., and van der Meer, J. R. (1998). Chromosomal integration, tandem amplification, and deamplification in Pseudomonas putida F1 of a 105-kilobase genetic element containing the chlorocatechol degradative genes from Pseudomonas sp. strain B13. J. Bacteriol. 180, 4360–4369. Reineke, W. (1998). Development of hybrid strains for the mineralization of chloroaromatics by patchwork assembly. Annu. Rev. Microbiol. 52, 287–331. Rodrı´guez-Herva, J., Garcia, V., Hurtado, A., Seguar, A., and Ramos, J. L. (2007). The ttgGHI solvent efflux pump operon of Pseudomonas putida DOT-T1E is located on a large selftransmissible plasmid. Environ. Microbiol. 9, 1550–1561. Rojo, F., Pieper, D. H., Engesser, K. H., Knackmuss, H. J., and Timmis, K. N. (1987). Assemblage of ortho cleavage route for simultaneous degradation of chloro- and methylaromatics. Science 238, 1395–1398. Ronchel, M. C., Ramos-Diaz, M. A., and Ramos, J. L. (2000). Retrotransfer of DNA in the rhizosphere. Environ. Microbiol. 2, 319–323. Sala-Trepat, J. M., Murray, K., and Williams, P. A. (1972). The metabolic divergence in the meta cleavage of catechols by Pseudomonas putida NCIB 10015. Physiological significance and evolutionary implications. Eur. J. Biochem. 28, 347–356. Santos, P. M., Mignogna, G., Heipieper, H. J., and Zennaro, E. (2002). Occurrence and properties of glutathione S-transferases in phenol-degrading Pseudomonas strains. Res. Microbiol. 153, 89–98. Santos, P. M., Benndorf, D., and Sa-Correia, I. (2004). Insights into Pseudomonas putida KT2440 response to phenol-induced stress by quantitative proteomics. Proteomics 4, 2640–2652. Schweigert, N., Hunziker, R. W., Escher, B. I., and Eggen, R. I. L. (2001a). Acute toxicity of (chloro-)catechols and (chloro-)catechol-copper combinations in Escherichia coli corresponds to their membrane toxicity in vitro. Environ. Toxicol. Chem. 20, 239–247. Schweigert, N., Zehnder, A. J. B., and Eggen, R. I. L. (2001b). Chemical properties of catechols and their molecular modes of toxic action in cells, from microorganisms to mammals. Environ. Microbiol. 3, 81–91. Shingler, V. (2003). Integrated regulation in response to aromatic compounds: From signal sensing to attractive behaviour. Environ. Microbiol. 5, 1226–1241. Shingler, V., and Pavel, H. (1995). Direct regulation of the ATPase activity of the transcriptional activator DmpR by aromatic compounds. Mol. Microbiol. 17, 505–513. Siguier, P., Filee, J., and Chandler, M. (2006). Insertion sequences in prokaryotic genomes. Curr. Opin. Microbiol. 9, 526–531. Sinclair, M. I., Maxwell, P. C., Lyon, B. R., and Holloway, B. W. (1986). Chromosomal location of TOL plasmid DNA in Pseudomonas putida. J. Bacteriol. 168, 1302–1308. Springael, D., and Top, E. M. (2004). Horizontal gene transfer and microbial adaptation to xenobiotics: New types of mobile genetic elements and lessons from ecological studies. Trends Microbiol. 12, 53–58.

32


Springael, D., Ryngaert, A., Merlin, C., Toussaint, A., and Mergeay, M. (2001). Occurrence of Tn4371-related mobile elements and sequences in (chloro)biphenyl-degrading bacteria. Appl. Environ. Microbiol. 67, 42–50. Suenaga, H., Mitsuoka, M., Ura, Y., Watanabe, T., and Furukawa, K. (2001). Directed evolution of biphenyl dioxygenase: Emergence of enhanced degradation capacity for benzene, toluene, and alkylbenzenes. J. Bacteriol. 183, 5441–5444. Tegova, R., Tover, A., Tarassova, K., Tark, M., and Kivisaar, M. (2004). Involvement of errorprone DNA polymerase IV in stationary-phase mutagenesis in Pseudomonas putida. J. Bacteriol. 186, 2735–2744. Tesoro Petroleum Companies, Inc. (2003). Material safety data sheet: gasoline. http://firstfuelbank.com/msds/Tesoro.pdf. Accessed February 25, 2011. Thomas, C. M. (2000). The Horizontal Gene Pool. Harwood Academic Publishers, Amsterdam, NL. Timmis, K. N., and Pieper, D. H. (1999). Bacteria designed for bioremediation. Trends Biotechnol. 17, 201–204. Top, E. M., and Springael, D. (2003). The role of mobile genetic elements in bacterial adaptation to xenobiotic organic compounds. Curr. Opin. Biotechnol. 14, 262–269. Top, E. M., Springael, D., and Boon, N. (2002). Catabolic mobile genetic elements and their potential use in bioaugmentation of polluted soils and waters. FEMS Microbiol. Ecol. 42, 199–208. Trefault, N., De la Iglesia, R., Molina, A. M., Manzano, M., Ledger, T., Perez-Pantoja, D., Sanchez, M. A., Stuardo, M., and Gonzalez, B. (2004). Genetic organization of the catabolic plasmid pJP4 from Ralstonia eutropha JMP134 (pJP4) reveals mechanisms of adaptation to chloroaromatic pollutants and evolution of specialized chloroaromatic degradation pathways. Environ. Microbiol. 6, 655–668. Tropel, D., and van der Meer, J. R. (2004). Bacterial transcriptional regulators for degradation pathways of aromatic compounds. Microbiol. Mol. Biol. Rev. 68, 474–486. Tropel, D., Meyer, C., Armengaud, J., and Jouanneau, Y. (2002). Ferredoxin-mediated reactivation of the chlorocatechol 2,3-dioxygenase from Pseudomonas putida GJ31. Arch. Microbiol. 177, 345–351. Tsuda, M., Tan, H. M., Nishi, A., and Furukawa, K. (1999). Mobile catabolic genes in bacteria. J. Biosci. Bioeng. 87, 401–410. Urgun-Demirtas, M., Stark, B., and Pagilla, K. (2006). Use of genetically engineered microorganisms (GEMs) for the bioremediation of contaminants. Crit. Rev. Biotechnol. 26, 145–164. Vaillancourt, F. H., Labbe, G., Drouin, N. M., Fortin, P. D., and Eltis, L. D. (2002). The mechanism-based inactivation of 2,3-dihydroxybiphenyl 1,2-dioxygenase by catecholic substrates. J. Biol. Chem. 277, 2019–2027. Vaillancourt, F. H., Fortin, P. D., Labbe, G., Drouin, N. M., Karim, Z., Agar, N. Y., and Eltis, L. D. (2005). Molecular basis for the substrate selectivity of bicyclic and monocyclic extradiol dioxygenases. Biochem. Biophys. Res. Commun. 338, 215–222. Vaillancourt, F. H., Bolin, J. T., and Eltis, L. D. (2006). The ins and outs of ring-cleaving dioxygenases. Crit. Rev. Biochem. Mol. Biol. 41, 241–267. van der Meer, J. R. (2002). Evolution of metabolic pathways for degradation of environmental pollutants. In ‘‘Encyclopedia of Environmental Microbiology’’ (G. Bitton, Ed.), pp. 1194–1207. John Wiley & Sons, New York. van der Meer, J. R. (2006). Environmental pollution promotes selection of microbial degradation pathways. Front. Ecol. Environ. 4, 35–42. van der Meer, J. R., and Sentchilo, V. (2003). Genomic islands and the evolution of catabolic pathways in bacteria. Curr. Opin. Biotechnol. 14, 248–254.


33

van der Meer, J. R., Roelofsen, W., Schraa, G., and Zehnder, A. J. B. (1987). Degradation of low concentrations of dichlorobenzenes and 1,2,4-trichlorobenzene by Pseudomonas sp. strain P51 in nonsterile soil columns. FEMS Microbiol. Lett. 45, 333–341. van der Meer, J. R., Zehnder, A. J., and de Vos, W. M. (1991). Identification of a novel composite transposable element, Tn5280, carrying chlorobenzene dioxygenase genes of Pseudomonas sp. strain P51. J. Bacteriol. 173, 7077–7083. van der Meer, J. R., Devos, W. M., Harayama, S., and Zehnder, A. J. B. (1992). Molecular mechanisms of genetic adaptation to xenobiotic compounds. Microbiol. Rev. 56, 677–694. van der Meer, J. R., Werlen, C., Nishino, S. F., and Spain, J. C. (1998). Evolution of a pathway for chlorobenzene metabolism leads to natural attenuation in contaminated groundwater. Appl. Environ. Microbiol. 64, 4185–4193. van der Meer, J. R., Ravatn, R., and Sentchilo, V. (2001). The clc element of Pseudomonas sp. strain B13 and other mobile degradative elements employing phage-like integrases. Arch. Microbiol. 175, 79–85. Ward, G., Parales, R. E., and Dosoretz, C. G. (2004). Biocatalytic synthesis of polycatechols from toxic aromatic compounds. Environ. Sci. Technol. 38, 4753–4757. Werlen, C., Kohler, H. P. E., and van derMeer, J. R. (1996). The broad substrate chlorobenzene dioxygenase and cis-chlorobenzene dihydrodiol dehydrogenase of Pseudomonas sp. strain P51 are linked evolutionarily to the enzymes for benzene and toluene degradation. J. Biol. Chem. 271, 4009–4016. Worsey, M. J., and Williams, P. A. (1975). Metabolism of toluene and xylenes by Pseudomonas putida mt-2: Evidence for a new function of the TOL plasmid. J. Bacteriol. 124, 7–13. Wright, B. E. (2000). A biochemical mechanism for nonrandom mutations and evolution. J. Bacteriol. 182, 2993–3001. Yuan, L., Kurek, I., English, J., and Keenan, R. (2005). Laboratory-directed protein evolution. Microbiol. Mol. Biol. Rev. 69, 373–392. Zhou, L. M., Timmis, K. N., and Ramos, J. L. (1990). Mutations leading to constitutive expression from the TOL plasmid meta-cleavage pathway operon are located at the Cterminal end of the positive regulator protein XylS. J. Bacteriol 172, 3707– 3710. Zylstra, G. J., and Gibson, D. T. (1989). Toluene degradation by Pseudomonas putida F1. Nucleotide sequence of the todC1C2BADE genes and their expression in Escherichia coli. J. Biol. Chem. 264, 14940–14946. Zylstra, G. J., McCombie, W. R., Gibson, D. T., and Finette, B. A. (1988). Toluene degradation by Pseudomonas putida F1: Genetic organization of the tod operon. Appl. Environ. Microbiol. 54, 1498–1503.

CHAPTER

2 Recent Advances in Hantavirus Molecular Biology and Disease Islam T. M. Hussein, Abdul Haseeb, Absarul Haque, and Mohammad A. Mir1

36 38 38 39 41 45 50 53 53 54 55 56 57 58 58 59 61

Contents

I. Introduction II. Hantavirus Molecular Biology A. Hantavirus structure B. Replication cycle of hantaviruses C. Nucleocapsid protein (N) D. RNA-dependent RNA polymerase E. Glycoproteins Gn and Gc III. Hantavirus Disease A. Epidemiology B. Reservoir hosts of hantaviruses C. Diseases caused by hantaviruses in humans D. Pathogenesis E. Diagnosis F. Therapy G. Vaccines IV. Future Prospects References

Abstract

Hantaviruses are emerging zoonotic pathogens that belong to the Bunyaviridae family. They have been classified as category A pathogens by CDC (centers for disease control and prevention). Hantaviruses pose a serious threat to human health because their infection causes two highly fatal diseases, hemorrhagic fever with

Department of Microbiology, Molecular Genetics and Immunology, University of Kansas, Medical Center, Kansas City, Kansas, USA 1 Corresponding author. e-mail address: [email protected] Advances in Applied Microbiology, Volume 74 ISSN 0065-2164, DOI: 10.1016/B978-0-12-387022-3.00006-9

#


35

36

Islam T. M. Hussein et al.

renal syndrome (HFRS) and hantavirus cardiopulmonary syndrome (HCPS). These pathogens are transmitted to humans through aerosolized excreta of their infected rodent hosts. Hantaviruses have a tripartite-segmented negative-sense RNA genome. The three genomic RNA segments, S, M, and L, encode a nucleocapsid protein (N), a precursor glycoprotein that is processed into two envelope glycoproteins (Gn and Gc) and the viral RNA-dependent RNA polymerase (RdRp), respectively. N protein is the major structural component of the virus, its main function is to protect and encapsidate the three genomic RNAs forming three viral ribonucleocapsids. Recent studies have proposed that N in conjunction with RdRp plays important roles in the transcription and replication of viral genome. In addition, N preferentially facilitates the translation of viral mRNA in cells. Glycoproteins, Gn and Gc, play major roles in viral attachment and entry to the host cells, virulence, and assembly and packaging of new virions in infected cells. RdRp functions as RNA replicase and transcriptase to replicate and transcribe the viral RNA and is also thought to have endonuclease activity. Currently, no antiviral therapy or vaccine is available for the treatment of hantavirus-associated diseases. Understanding the molecular details of hantavirus life cycle will help in the identification of targets for antiviral therapeutics and in the design of potential antiviral drug for the treatment of HFRS and HCPS. Due to the alarming fatality of hantavirus diseases, development of an effective vaccine against hantaviruses is a necessity.

I. INTRODUCTION Two major outbreaks that occurred in the past led to the discovery of hantaviruses and directed the attention of researchers toward the hantavirus-associated diseases. The first outbreak occurred during the Korean War (1950–1953), with more than 3000 cases of illness reported among the United Nations troops. The disease was initially named ‘‘Korean hemorrhagic fever’’ and is now commonly referred to as hemorrhagic fever with renal syndrome (HFRS). The second outbreak, which occurred in the Four Corners region of the United States in 1993, triggered the attention of the World Health Organization toward a new highly lethal disease. The disease was initially referred to as four-corner disease and is now called hantavirus pulmonary syndrome (HPS) or hantavirus cardiopulmonary syndrome (HCPS). Hantaviruses cause serious human illness with a mortality rate that ranges from 15% (for HFRS) to 50% (for HCPS). Twenty-five years after the Korean War, the etiologic agent of this disease, the Hantaan virus (HTNV), was identified in the striped field mouse (Apodemus agrarius) (Lee et al., 1981a). Further studies revealed that the newly identified virus belonged to the Bunyaviridae family. In 1981, a new genus termed as

Hantavirus Molecular Biology and Disease

37

‘‘hantavirus’’ was introduced into the Bunyaviridae family, which included the viruses that cause HFRS (Fig. 2.1). Further investigations revealed that, unlike other members of the Bunyaviridae family, hantaviruses did not have an arthropod vector and exclusively establish a persistent infection in the population of their specific rodent hosts. The landmark discovery of HTNV encouraged more efforts to identify the etiologic agents of HFRS-related disease in Asia, Europe, and the United States. These efforts led to the discovery of Haantan-like viruses in other rodents from Far East Russia, China, and South Korea. Dobrava virus (DOBV), a distinct hantavirus, was isolated from Apodemus flavicollis, A. Agrarius, and Apodemus ponticus in Europe (Avsic-Zupanc et al., 1995, 2000; Golovljova et al., 2000; Jakab et al., 2007; Klempa et al., 2005, 2008; Klingstrom et al., 2006a). In 1980s, a rat-born Seoul virus (SEOV) was found to cause HFRS in urban areas of Asia (Chan et al., 1987; Kim et al., 1995). Around the same time period, Puumala virus (PUUV), the etiologic agent of nephropathia epidemica, a milder form of HFRS reported in 1930s in Europe, was identified in bank voles (Myodes glareolus) (Clement et al., 2003). Despite these early efforts, the lack of Paramyxoviridae Filoviridae Non segmented Rhabdoviridae Bornaviridae Bunyavirus Negative-stranded RNA viruses

Hantavirus Bunyaviridae

Nairovirus Phlebovirus

Segmented

Tospovirus Orthomyxoviridae Arenaviridae

FIGURE 2.1 Classification of negative-stranded RNA viruses: negative-stranded RNA viruses are classified into two main groups: those with nonsegmented and segmented genomic RNA. The nonsegmented group includes four families: Paramyxoviridae, Filoviridae, Rhabdoviridae, and Bornaviridae. RNA viruses whose genomes are segmented have been further classified into three families: Bunyaviridae, Orthomyxoviridae, and Arenaviridae. Hantaviruses are classified with the family Bunyaviridae, which includes five genera: Bunyavirus, Hantavirus, Nairovirus, Phlebovirus, and Tospovirus.

38


advanced technology delayed the identification of etiologic agents of HFRS. However, the etiologic agent and rodent reservoir of HCPS, the Sin Nombre virus (SNV), were discovered within few weeks after the 1993s outbreak in the Four Corners region of the United States (Hjelle et al., 1994; Nichol et al., 1993). After this outbreak, other hantavirus strains causing HCPS were discovered and more than 2000 cases of HCPS have been reported throughout the Americas (Aquino et al., 2003; Barclay and Rubinstein, 1997; Bayard et al., 2004; Bohlman et al., 2002; Chu et al., 2006; Fulhorst et al., 1997; Hjelle et al., 1994, 1996; Jiang et al., 1997; Johnson et al., 1997; Jones et al., 2008; Nichol et al., 1993; Vincent et al., 2000). Currently, the hantavirus genus includes more than 21 species and more than 30 genotypes (Avsic-Zupanc et al., 2000; Chu et al., 2006), although many more remain undiscovered due to the lack of advanced technology in other parts of the world including Africa, Middle East, and the Indian subcontinent. For example, recent discoveries of shrewborn hantavirus in several countries support the existence of other hantavirus species worldwide (Arai et al., 2007; Kang et al., 2009a). Annually, 150,000–200,000 cases of hantavirus infection are reported worldwide. Due to high emergence and significant mortality of hantavirus infections, a serious attention from research scientists and world health organizations is required to promote public awareness and accelerate efforts for the treatment of hantavirus-associated diseases. Here, we summarize the findings about the microbiology of hantaviruses with an emphasis on nucleocapsid protein, glycoproteins Gn, Gc, and RNA-dependent RNA polymerase (RdRp). A brief synopsis about different aspects of hantavirus-associated disease is also presented in this chapter.

II. HANTAVIRUS MOLECULAR BIOLOGY A. Hantavirus structure Hantavirus particles generally appear spherical or pleomorphic in electron micrographs (Fig. 2.2). They range in diameter from 80 to 120 nm (Goldsmith et al., 1995; Martin et al., 1985). Hantaviruses have tripartite single-stranded negative-sense RNA genome that encodes an RdRp (large or L segment), two glycoproteins (medium or M segment), and a nucleocapsid protein (N) (small or S segment). The negative-sense genomes serve as templates for producing positive-sense complementary RNA (cRNA) and messenger RNA (mRNA). The total size of the RNA genomes ranges from 11,845 nucleotides for HTNV to 12,317 nucleotides for SNV (Hooper and Schmaljohn, 2001; Jonsson et al., 2010; Schmaljohn et al., 1985, 1986). The viral RNA segments are coated with the nucleocapsid protein forming three helical ribonucleoprotein (RNP) complexes (Fig. 2.2). The viral RdRp is associated with these nucleocapsids (Dahlberg et al., 1977; Obijeski et al., 1976). The sequences at both the 30 and 50 termini of each RNA segment are


39

FIGURE 2.2 Hantavirus structure: (A) pictorial representation of hantavirus particle, showing three nucleocapsids enveloped in a lipid bilayer. Glycoproteins Gn and Gc buried in the lipid membrane are shown. (B) Thin-section electron micrograph of an SNV isolate from the outbreak of HCPS that occurred in the southwestern United States in 1993. Electron micrograph was obtained from the CDC website with permission. http:// www.cdc.gov/ncidod/diseases/hanta/hps/noframes/hpsem.htm.

complementary forming ‘‘panhandle’’ structures that are specifically recognized by the N protein and were shown to be important for viral transcription and replication. Viral mRNAs contain untranslated regions (UTRs) at both their 50 and 30 ends that flank the open reading frame (ORF) of the encoded viral protein. The 50 UTR of viral mRNA contains trinucleotide repeats (UAGUAGUAG), which were suggested to be involved in N-mediated translation initiation of viral mRNA. Similarly, a triplet repeat at the 30 UTR of vRNA has been proposed to play a role in the prime and realign mechanism of transcription initiation by the viral RdRp (Hewlett et al., 1977; Mir and Panganiban, 2004, 2005, 2010; Pettersson and von Bonsdorff, 1975; Raju and Kolakofsky, 1989). Hantaviruses are enveloped with an outer lipid bilayer derived from the Golgi membranes (Fig. 2.2). The envelope carries two viral-encoded glycoproteins, Gn and Gc, in the form of heterodimers which assemble into higher order oligomers and appear as projections or spikes on the outer surface of the virion (Hepojoki et al., 2010a; Huiskonen et al., 2009, 2010; Overby et al., 2008). The virion consists of > 50% protein, 20–30% lipid, 7% carbohydrate, and 2% RNA (McCaughey and Hart, 2000; Obijeski and Murphy, 1977). Hantaviruses can be easily inactivated by treatment with lipid solvents or nonionic detergents, which destroy the viral envelope (Schmaljohn and Nichol, 2007).

B. Replication cycle of hantaviruses Hantaviruses infect multiple cell lines including endothelial, epithelial, macrophages, dendritic, and lymphocytes. The replication cycle begins with the attachment of virus particles to host cell surface receptors

40


hmen

t

(Goldsmith et al., 1995; Mackow and Gavrilovskaya, 2001; Markotic et al., 2007; Raftery et al., 2002; Spiropoulou, 2001). Several studies suggest that the attachment is mediated by the interaction of viral Gn protein with integrin receptors on the surface of host cells (Fig. 2.3). The b1 and b3 integrins are considered to be the receptors for apathogenic and pathogenic hantaviruses, respectively (Gavrilovskaya et al., 1998, 1999; Larson et al., 2005). However, it has been reported that hantaviruses can infect cells lacking integrin receptors, suggesting that integrins may not be the only receptors that hantaviruses employ to attach to host cells (Mou et al., 2006). After attachment, cell entry is mediated via clatherin-coated pits, and virions are ultimately delivered to lysosomes (Fig. 2.3). Within the endolysosomal compartment, virions are uncoated and three viral capsids are released into the cytoplasm ( Jin et al., 2002). Viral RdRp initiates transcription and generates three mRNAs, one from each viral RNA segment S, M, and L. The translation of S and L segment-derived mRNAs occurs on free ribosomes, while M segment-derived mRNAs

Attac

vRNA Entry Capsids

Rep

lica

tion

Uncoating

Transcription

Exit G1/G2

cRNA

Translation G1/G2 Golgi complex

Nucleus

FIGURE 2.3 Hantavirus replication cycle: virus particle attaches to the integrin receptors on host cell surface. After entry, uncoating takes place, followed by mRNA synthesis by RdRp. The three mRNA molecules are translated by host cell translation machinery, generating viral proteins. Viral genome is synthesized by RdRp via a cRNA intermediate and is packaged into new virus particles that bud off the host cell.


41

are translated on the rough endoplasmic reticulum (ER). The glycoprotein precursor is intrinsically cleaved at a highly conserved amino acid motif, WAASA, generating two glycoproteins, Gn and Gc, respectively (Lober et al., 2001; Ruusala et al., 1992; Spiropoulou, 2001). After glycosylation in the ER, both Gn and Gc are transported to the Golgi complex, laying the foundations for final destinations where virions are maturated (Antic et al., 1992a; Plyusnin et al., 1997; Ravkov et al., 1998; Ruusala et al., 1992). After initial rounds of transcription, viral RdRp switches to the replication mode and generates three viral genomic RNAs, which are encapsidated by the viral N protein to form three nucleocapsids. Mature viral nucleocapsids are transported to specific destinations on the Golgi membrane that are studded with the Gn and Gc proteins. However, the molecular mechanisms governing the specific recognition and encapsidation of viral genome by N, transport of capsids to Golgi, and budding of nascent virions into and out of Golgi complex are still unknown. Mechanisms of incorporation of viral RdRp into virus particles are fascinating and need more attention. There are reports suggesting that the assembly and maturation of New World hantaviruses take place on the plasma membrane (Ravkov and Compans, 2001). The evidence supporting this pathway for assembly and maturation is based on observations that SNV and black Creek Canal virus (BCCV) particles were not observed in infected cells. However, localization of N protein from both old and new word hantaviruses on Golgi apparatus favors the possibility of their maturation on Golgi (Ramanathan and Jonsson, 2008; Ramanathan et al., 2007; Ravkov and Compans, 2001; Spiropoulou et al., 2003).

C. Nucleocapsid protein (N) Hantavirus nucleocapsid protein (N) is the major viral structural component; its main function is to protect and encapsidate the viral RNA forming viral RNP complex. It is encoded by the S segment vRNA and is abundantly expressed in the cytoplasm of infected cells. Therefore, it is considered as the most predominant viral antigen in the serologic response to infection. Hantaviral N protein has a molecular mass of approximately 50 kDa and contains 429–433 amino acid residues (de Carvalho Nicacio et al., 2001; Plyusnin et al., 1997; Van Epps et al., 1999). It is highly conserved across hantaviruses and contains cross-reactive epitopes that encompass the first 100 amino acids at N-terminal (Elgh et al., 1996; Gott et al., 1997; Schmaljohn et al., 1986, 1987; Yamada et al., 1995). Further, serotype-specific conformational epitopes have been detected in about half of the C-termini of N proteins by serotype-specific monoclonal antibodies (MAbs) (Ruo et al., 1991; Yoshimatsu et al., 1996).

42


1. Biological functions of N protein N protein is a multifunctional molecule that interacts with host cellular proteins and plays several important roles to facilitate hantavirus replication in infected cells.

a. Viral RNA encapsidation and assembly Several studies showed that N binds specifically to a region near the 50 terminus of vRNA, which is indicative of its function during encapsidation (Gott et al., 1993; Jonsson et al., 2001; Mir and Panganiban, 2005; Osborne and Elliott, 2000; Severson et al., 1999). It was also reported that N differentially interacts with minussense vRNA, plus-sense cRNA, and mRNA. Cell culture-based experiments have demonstrated that encapsidation requires full-length vRNA or cRNA molecules, since no mRNA molecules were detected in viral nucleocapsids ( Jin and Elliott, 1993). Moreover, SNV N protein was shown to play the role of RNA chaperone that facilitates transient dissociation of misfolded RNA structures and creates opportunities for the folding of RNA into biologically functional and thermodynamically stable higher order structures (Mir and Panganiban, 2005). A three-dimensional (3D) structural model of N protein is still lacking. However, the functionally important regions have been identified in several mutagenesis studies. The RNA-binding domain of N protein was mapped to the central conserved region that extends from amino acid 175 to 217 (Li et al., 2002). In addition, the lysine residues dispersed between positions 175 and 429 and three more residues, including E192, Y206, and S217 located in the RNA-binding domain, were also shown to be important for RNA binding (Severson et al., 2005). Hantavirus N protein undergoes protein–protein interaction and forms stable trimers both in vivo and in vitro. It has been shown that trimeric N recognizes the vRNA panhandle with specificity and high affinity (Mir et al., 2006). N-panhandle interaction has been proposed to mediate the selective encapsidation and packaging of vRNA genome during virus assembly (Alfadhli et al., 2001, 2002; Kaukinen et al., 2001, 2003, 2004; Mir and Panganiban, 2004). Using the MultiCoil prediction algorithm (Wolf et al., 1997), coiled-coils spanning amino acid residues (1–34 and 38–80) have been predicted for the N protein of several hantaviruses, including HTNV, SEOV, SNV, Prospect Hill virus (PHV), and Tula virus (TULV) (Alfadhli et al., 2001). Both N- and C-terminal regions have been implicated in mediating the homotypic interaction between N molecules, and putative coiled-coil motifs in the N-terminal region of N protein have been proposed to facilitate trimerization. A ‘‘head-to-head, tail-to-tail’’ model for trimerization was proposed, where three monomers of N protein are brought together via interaction of their N-terminal coiledcoil domains. This initial contact is followed by the interaction between


43

the C-terminal a-helices that form a shared hydrophobic space and thus consolidate the trimer formation (Alfadhli et al., 2001, 2002; Kaukinen et al., 2001, 2003, 2004).

b. Transcription and translation Apart from its role in packaging, N has been found to be involved in transcription and replication of viral genome in conjunction with viral RdRp or by interacting with template RNA during replication (Blakqori et al., 2003; Bridgen and Elliott, 1996; Ikegami et al., 2005; Kohl et al., 2004; Pinschewer et al., 2003). Viral mRNAs are translated by the host cell translation machinery. Recent studies have shown that SNV N protein binds to mRNA 50 caps and 40S ribosomal subunit with high affinity and specificity (Fig. 2.4). These interesting findings have suggested that, in the host cell cytoplasm where cellular transcripts are competing for the same translation machinery, N protein facilitates the translation of hantaviral mRNAs by

FIGURE 2.4 A model depicting the roles of N in transcription and translation initiation: (A) cellular mRNA targeted to P-bodies for decay. N moves to P-bodies and binds to mRNA caps. (B) N with a capped RNA primer loaded at the cap-binding site specifically binds the 30 terminus of vRNA and favors primer annealing. (C) Trimeric N binds the triplet repeat sequence of viral mRNA 50 UTR. N also binds 40S ribosomal subunit and loads it onto the 50 UTR.

44


preferentially loading ribosomes onto capped viral transcripts (Klempa et al., 2008). Further studies have shown that the sequence GUAGUAG, in the 50 UTR of hantavirus mRNA, is sufficient for preferential N-mediated translation initiation (Mir and Panganiban, 2010). N-mediated translation initiation mechanism is different from the complex translation initiation strategy used by eukaryotic cells. It is likely that N initiates the translation of capped mRNA without the requirement of eIF4F complex, an amalgam of three initiation factors, eIF4E, eIF4G, and eIF4A (Panganiban and Mir, 2009). In host cells, after the completion of translation, the mRNAs are deadenylated and decapped, followed by degradation by exonucleolytic digestion from both 50 and 30 ends. In the 50 –30 degradation pathway, the decapping enzyme DCP2 in coordination with several other proteins removes the 50 mRNA cap, followed by degradation by exonuclease XRN1. This degradation mechanism takes place in discrete cytoplasmic foci called processing bodies (P-bodies) (Beckham and Parker, 2008). Interestingly, a detailed analysis by confocal imaging revealed that N resides in cellular P-bodies. Further studies revealed that cellular 50 capped mRNA oligoribonucleotides are rescued by N in virus-infected cells and stored in P-bodies for the later use as primers by the viral RdRp during transcription initiation (Mir et al., 2008). Moreover, it was reported that N protein has distinct cap and RNA-binding sites that independently interact with mRNA cap and viral genomic RNA, respectively. In addition, N can simultaneously bind to both mRNA cap and vRNA. N undergoes distinct conformational changes after binding to mRNA cap, vRNA, or both. The conformationally altered N with a capped primer loaded at the cap-binding site specifically binds the conserved 30 nine nucleotides of vRNA (30 AUCAUCAUC) and assists the bound primer to anneal at the 30 terminus. The annealed primer is later elongated by the RdRp and a nascent viral mRNA is synthesized. Therefore, it was suggested that the cap-binding site of N, in conjunction with RdRp, plays key roles during the transcription and replication initiation of vRNA genome (Mir et al., 2010). Taken together, these observations imply that N has a role in generating the capped RNA primer in P-bodies and also assists the capped primer to anneal to the 30 terminus of vRNA template during transcription initiation (Fig. 2.4).

c. Interaction with cellular proteins N protein performs some ambassadorial functions by interacting with some cellular proteins to facilitate viral dissemination in infected cells. For example, interaction of N with actin filaments has been reported to mediate the transport of newly synthesized viral RNPs to the plasma membrane (Ravkov et al., 1998). It was also shown that N protein interacts with the Fas-mediated apoptosis enhancer Daxx, a death-domain adaptor protein, which transduces death

45


signals through the Jun N-terminal kinase (JNK) pathway (Yang et al., 1997). In similar studies, interaction of N protein with Daxx and small ubiquitin-like modifiers (SUMO-1) pathway components in infected cells has been reported (Alfadhli et al., 2002; Cho et al., 2002; Kaukinen et al., 2003; Lee et al., 2003).

D. RNA-dependent RNA polymerase The protein encoded by the largest ‘‘L’’ segment of hantaviruses is an RdRp. The RdRp is responsible for the replication and transcription of the viral genome. The negative-sense hantaviral RNAs (vRNAs) are copied by the L protein to produce positive-sense cRNA intermediates that serve as templates for genome replication and mRNA synthesis (Fig. 2.5).

1. Sequence homology and structural motifs Sequences of several hantavirus genomes are now available. From the available sequencing data, it is evident that the size of the L RNA of hantaviruses ( 6.5 kb) is similar to those of other members of the Bunyaviridae family, except for the Tospovirus ( 8.9 kb) and the Nairovirus ( 12 kb) genera. The reverse complement of the L segment of all the members of the Bunyaviridae family has a single large ORF with flanking 50 and 30 UTRs. The predicted size of the protein encoded by this ORF is approximately 250 kDa, which has also been confirmed experimentally by few groups (Elliott et al., 1984; Kukkonen et al., 2004). There are a total of four classes of nucleotide polymerases: DNAdependent DNA polymerase (DdDp), DNA-dependent RNA polymerase (DdRp), RNA-dependent DNA polymerase (RdDp or reverse transcriptase), and RdRp. Amino acid sequence comparisons of all the nucleotide polymerases have revealed an interesting pattern of conserved motifs UTR

ORF

cRNA (+) 5⬘

UTR 3⬘

Replication vRNA (–) 3⬘

5⬘ Transcription

mRNA (+) 5⬘

3⬘

FIGURE 2.5 Schematic representation of reactions carried out by RdRp during the hantaviral life cycle: vRNA carried by the virus in the host cell is first transcribed to produce mRNA, which is translated into viral proteins. Later in the life cycle, vRNA is amplified for packaging through a cRNA intermediate. vRNA and cRNA are exact reverse complement copies of each other. mRNA in some cases has a shorter 30 UTR (depicted by dashed line).

46


N-ter

F

A

B

C

D

E

C-ter

RdRp RdDp DdDp DdRp

FIGURE 2.6 Conserved structural motifs among four classes of nucleotide polymerases: motifs A–D are found in all the polymerases. Motif E is found in polymerases that use RNA as a template. There is an extra motif F found only in RdRps.

found in different classes of polymerases (Fig. 2.6; Hansen et al., 1997; Joyce and Steitz, 1995). Motifs A–D are shared by all polymerases. Motif E is found in those polymerases that recognize RNA as a template (reverse transcriptases and RdRPs). An extra motif F (also called as pre-motif A) is specifically found in RdRps (Bruenn, 2003; Hansen et al., 1997; Kamer and Argos, 1984; Muller et al., 1994; O’Reilly and Kao, 1998; Poch et al., 1989; Toh et al., 1985; Tordo et al., 1988). The amino acid residues from these conserved motifs play critical roles in different aspects of the nucleotidyl transferase reaction of these enzymes, which include binding of metal ions, nucleoside triphosphates (NTPs), and the RNA/DNA template. Few additional conserved regions have been found in the amino acid sequences of RdRps of segmented negative strand RNA viruses (Aquino et al., 2003; de Haan et al., 1991; Kukkonen et al., 1998; Muller et al., 1994; Nemirov et al., 2003; Stohwasser et al., 1991); however, the functional importance of these motifs has not yet been validated. Based on the organization of the conserved motifs in the amino acid sequence, the hantavirus RdRp is predicted to have a structure similar to other polymerases. The crystal structures of all polymerases show a common domain organization, which resembles a ‘‘right hand,’’ and it includes the subdomains denoted as ‘‘finger,’’ ‘‘palm,’’ and ‘‘thumb’’ (Kohlstaedt et al., 1992). Among all known RdRps, an additional N-terminal domain is found that bridges the fingers and thumb domains to make a tunnel-like structure, which is otherwise open U-shaped in most of the other polymerases (Butcher et al., 2001; Lesburg et al., 1999; Ng et al., 2008). The crystal structure of hepatitis C virus shows the palm subdomain to be constituted of b-strand of motifs A and C and a-helix of motif D. Motifs B and F, folded into an a-helix, form the fingers subdomain and the motif E, in form of an antiparallel b-sheet, is present between the palm and the thumb subdomains (Lesburg et al., 1999). Conserved Arg and Lys residues in motif F and a highly conserved Asp in motif A play critical roles in binding of the RdRp with NTPs (Arnold and Cameron, 2004; Bressanelli et al., 2002; Gohara et al., 2000, 2004; Huang et al., 1998; Joyce and Steitz, 1995).


47

Like other polymerases, the RdRps also show a dependence on binding of two divalent metal ions in the active site (Arnold et al., 1999; Doublie and Ellenberger, 1998; Rothwell and Waksman, 2005; Steitz et al., 1994). The conserved Asp residues in motifs A and C play a crucial role in binding of the two metal ions. Mutation of these Asp residues has been shown to inactivate or alter the activity of several RdRps (Arnold et al., 1999; Jablonski and Morrow, 1995; Vazquez et al., 2000). In a mutational analysis of Bunyamwera RdRp, replacement of the conserved aspartates of motifs A and C abolished the polymerase activity (Dunn et al., 1995). In a large-scale mutational study carried out on RdRp of Lassa virus (family Arenaviridae), an RdRp domain (between residues 1040 and 1540) was identified and a functional element within RdRp was found, which was important for transcription but not replication of the genome (Hass et al., 2008). RdRps of both positive-stranded RNA viruses such as poliovirus as well as negative-stranded RNA viruses including Sendai virus (Cevik et al., 2003; Smallwood et al., 2002), human parainfluenza type 3 virus (Smallwood and Moyer, 2004), measles virus (Cevik et al., 2004), lymphocytic choriomeningitis virus (LCMV) (Sanchez and de la Torre, 2005), and rift valley fever virus (RVFV) (Zamoto-Niikura et al., 2009) need oligomerization to exert their polymerization function.

a. Localization The site of RNA synthesis among the viruses of the Bunyaviridae family was believed to be the cytoplasm of the host cell. This was based on a study on La Cross virus in which the newly synthesized RNA was found to be in the cytoplasmic fraction where it was monitored by pulse-labeling followed by cell fractionation (Rossier et al., 1986). But based on later studies on localization of the two viral proteins essential for RNA synthesis, namely, L and N, it is now thought that RNA synthesis among bunyaviruses is membrane associated. Kukkonen et al. (2004) studied the localization of L-GFP fusion protein and found it to localize in the perinuclear region (Kukkonen et al., 2004). This is consistent with the fact that RNA synthesis of all positive-stranded viruses is membrane associated (Salonen et al., 2005). Further studies on other viruses of this family will be required to confirm if that is a general mechanism employed by these viruses. b. Mechanism of action Similar to all the other negative-sense RNA viruses, hantavirus genomic RNA segments contain UTRs on both the 50 and the 30 ends. The termini of the UTRs are partially complementary to each other due to which the segments fold to make a panhandle-like structure (Chizhikov et al., 1995; Kukkonen et al., 1998; Meyer and Schmaljohn, 2000; Padula et al., 2002). Several minireplicon systems in

48


which reporter genes (such as Renilla luciferase or chloramphenicol acetyltransferase) are flanked by the UTRs have been developed for members of the Bunyaviridae family (Blakqori et al., 2003; Dunn et al., 1995; Flick and Pettersson, 2001; Flick et al., 2003; Lopez et al., 1995). Studies based on these minireplicon systems have shown that 50 and 30 UTRs serve as promoters for replication of the segments and transcription of the encoded reading frames. The UTRs are encapsidated by nucleocapsid protein and associate with RdRp both in the host cells and in the virion, and only these nucleocapsids are believed to be functional templates for mRNA synthesis and RNA replication by the viral RdRp.

c. mRNA synthesis The viral RdRp is responsible for the synthesis of viral mRNA. Several unique features involved in this process are detailed here. Sequence analysis of the 50 termini of mRNAs of several Bunyaviruses has revealed the presence of short (10–18 nucleotides) nontemplated 50 end sequences (Bishop et al., 1983; Garcin et al., 1995; Simons and Pettersson, 1991). These sequences were later found to be derived from the 50 ends of host mRNAs by a mechanism similar to ‘‘cap-snatching’’ originally described for influenza virus. In this mechanism, a viral endonuclease cleaves the 50 termini of the host mRNAs and uses them as primers to initiate the transcription of viral mRNAs (Bouloy et al., 1978; Krug, 1981; Krug et al., 1979; Plotch et al., 1981). The endonuclease activity required for cap-snatching is believed to reside in the viral polymerase itself (Patterson et al., 1984). Based on the observation that all the mRNAs contain a G residue at 1 position, a prime and realign mechanism for mRNA synthesis in the Bunyaviridae family was proposed. In this mechanism, the G residue of the capped primer snatched from the host mRNA aligns with a C residue, upstream of the 30 terminus of the viral template. The primer is elongated few nucleotides by RdRp. The extended primer detaches and realigns to the 30 end of the terminus and then extends further to complete the mRNA synthesis (Garcin et al., 1995). Termination of mRNA synthesis and polyadenylation of the synthesized mRNA has been shown to vary among S, M, and L segments (Hutchinson et al., 1996). Synthesis of S segment mRNA terminates downstream of a CCACCC motif found around 200 nucleotide downstream of the 50 end of the vRNA template. The S segment mRNA is not polyadenylated (Hutchinson et al., 1996). The synthesis of the L mRNA extends to the 50 terminus of the L vRNA and is also not polyadenylated (Hutchinson et al., 1996). However, the M segment of SNV was found to be polyadenylated and a potential polyadenylation–transcription termination signal was mapped and was found to be conserved among all hantaviruses (Hutchinson et al., 1996).


49

d. RNA replication Replication of the vRNA genome is carried out by the viral RdRp through an antigenomic RNA (cRNA) intermediate. Unlike mRNA, the cRNA is exact complement of the viral genomic segments without any cap structure and host-derived sequences (Bishop et al., 1983; Eshita et al., 1985; Garcin et al., 1995; Obijeski et al., 1980; Raju and Kolakofsky, 1987). The cRNA was shown to be encapsidated by the N protein (Patterson et al., 1984). The 50 terminus of the vRNA in hantaviruses was found to contain uridine monophosohate (pU) instead of triphosphate (pppU; Chizhikov et al., 1995; Garcin et al., 1995). This observation was explained by a prime and realign mechanism for cRNA and vRNA synthesis (Garcin et al., 1995). Briefly, after elongation by few nucleotides, the capped RNA primer with a terminal ‘‘G’’ residue slides back on the 30 triplet repeat sequence of cRNA template, leaving a protruding GTP that is nonaligned with the 30 end. The protruding GTP is subsequently cleaved off by the polymerase leaving a pU residue at the 50 end (Garcin et al., 1995). e. Error, evolution, and editing The promiscuous nature of polymerization by RdRps due to lack of proof reading ability is thought to be the primary source of evolution in RNA viruses. This gives the virus an ability to replicate in different hosts and become pathogenic to humans. But the virus has to maintain a close window of variation to avoid the ‘‘error catastrophe’’ as too much of these errors may lead to generation of nonviable genome and reduction of the overall viral fitness (Crotty and Andino, 2002). Drake et al. have estimated an error rate of approximately 1 mutation/replication/genome for hantaviruses (Drake, 1999). The mean rate of evolutionary change in hantaviruses has been approximated to be within the range of 102–104 substitutions/site/year (Ramsden et al., 2008). RNA viruses employ several mechanisms to generate useful variations and to edit deleterious ones, which include reassortment and recombination (Barr and Fearns, 2010). Natural reassortment has been detected among hantaviruses (Henderson et al., 1995; Li et al., 1995; Plyusnin et al., 1997), and several experimental reassortants have been generated in the laboratory by infecting different strains of the same hantavirus (Ebihara et al., 2000; Rodriguez et al., 1998). Recently, Handke et al. were able to generate in vitro reassortants between pathogenic PUUV and nonpathogenic PHV (Handke et al., 2010). RNA recombination in viruses usually occurs by a ‘‘copy choice’’ mechanism, in which a replicating polymerase stops copying one RNA molecule and switches to another (Copper et al., 1974; Worobey and Holmes, 1999). A recombinant TULV was successfully generated through homologous recombination by infecting Vero E6 cells with one strain and by providing S cRNA of another TULV strain via plasmid-driven expression (Plyusnin et al., 2002). A recent study in Bunyamwera virus (BUNV)

50


has shown that recombination may also occur near the end of the templates to keep the ends intact (Walter and Barr, 2010).

E. Glycoproteins Gn and Gc 1. Expression, structural organization, and trafficking The envelope glycoproteins are encoded by the viral M segment RNA. They are expressed as a glycoprotein precursor (GPC) polypeptide, which is cleaved by host signal peptidase in the ER lumen to generate N-terminal (G1 or Gn) and C-terminal (G2 or Gc) fragments (Fig. 2.7; Fazakerley and Ross, 1989; Spiropoulou et al., 2003). Cleavage occurs after a highly conserved pentapeptide WAASA motif (Lober et al., 2001). The GPC has neither been detected in infected cells nor in cells transfected with a plasmid encoding M segment mRNA, but was only detectable by in vitro translation of RNA transcripts. Therefore, it was suggested that the proteolytic processing of the GPC occurs during translation (Pensiero and Hay, 1992; Schmaljohn et al., 1987; Ulmanen et al., 1981). Gn and Gc are type I transmembrane proteins modified by N-linked glycosylation, with the N-terminus exposed on the surface of the virion and the C-terminus anchored in the membrane (Shi and Elliott, 2004). They have high tendency to form disulfide bridges due to their high content of cysteine residues (4–7%) (Antic et al., 1992a). SNV GPC is 1140 amino acids in length. The Gn of SNV consists of 652 residues (MW 75 kDa), with a predicted transmembrane domain and a cytoplasmic tail (CT) domain of 142 amino acids. Gc is 488 residues in length (MW 55 kDa) and has a predicted shorter CT of eight amino acids (Spiropoulou et al., 2003). HTNV GPC has four hydrophobic domains. Domain I extends from position 1 to 17. Domains II (position 441–515) and Cleavage site (WAASA) GPC

N

C

I

ll

lV

lll

Gn (G1)

Gc (G2)

FIGURE 2.7 Schematic illustration of the hantavirus glycoprotein precursor (GPC) and its maturation: GPC is cotranslationally cleaved in the ER by host signal peptidase at a conserved WAASA motif, generating Gn (G1) and Gc (G2). The N-terminus, C-terminus, and the four hydrophobic regions (I–IV) are indicated.


51

IV (position 1097–1127) are the transmembrane domains of Gn and Gc, respectively (Spiropoulou et al., 1994). Domain II is very long and the actual membrane spanning portion is not well defined. Domain III extends from position 627 to 648 and ends with the highly conserved pentapeptide motif WAASA before the N-terminus of Gc (Spiropoulou, 2001). Most enveloped viruses bud from the plasma membrane into the extracellular space. However, in the case of family Bunyaviridae, many members of this family mature by budding into the lumen of the Golgi complex. Progeny virions are then released by fusion of secretory vesicles with the plasma membrane (Kuismanen et al., 1982; Schmaljohn and Nichol, 2007). Additional evidence suggests that the assembly of New World hantaviruses (e.g., SNV and BCCV) may occur at the plasma membrane (Ravkov et al., 1997). The reason why some viruses have evolved to bud from the Golgi membranes is poorly understood, but it might be due to the presence of Golgi retention signals in viral glycoproteins, which allow them to accumulate in the Golgi complex after their maturation in the ER (Antic et al., 1992b; Shi and Elliott, 2002; Spiropoulou et al., 2003). There is no clear consensus on the exact location of these Golgi targeting signals in different members of family Bunyaviridae, and it appears that different viruses, even within a single genus, have their own strategies for Golgi targeting (Shi et al., 2004). For members of the Bunyavirus genus, this signal was mapped to the N-terminus of Gc (Lappin et al., 1994). The same signal was reported to be located in the transmembrane domain and CT of Gn for members of genus Phlebovirus (Andersson et al., 1997; Gerrard and Nichol, 2002; Matsuoka et al., 1994). It was also suggested that translocation of Gn and Gc from the ER to the Golgi complex depends on their interaction and heterodimerization because neither of them was able to leave the ER when expressed individually (Deyde et al., 2005; Ruusala et al., 1992; Shi and Elliott, 2002; Spiropoulou et al., 2003).

2. Roles in virus biology Glycoproteins, Gn and Gc, play a significant role in the biology of hantaviruses, including virus entry to host cells, virulence, and assembly and packaging of new virions in infected cells.

a. Attachment and entry Hantaviruses enter the cells via receptormediated endocytosis. The acidic environment of the endosome facilitates fusion between the viral and cellular membranes, with a consequent release of the viral nucleocapsids into the cytoplasm ( Jin et al., 2002). Viral glycoproteins mediate attachment of virus to endothelial cells via interaction with surface b3 integrin receptors (Arikawa et al., 1989; Gavrilovskaya et al., 1998, 1999).

52


b. Assembly and packaging Gn has a 142-residue C-terminal CT, which plays important roles in viral assembly and host–pathogen interactions. The lack of a matrix protein in hantaviruses, which plays an important role in the assembly and budding of many enveloped viruses, is thought to be substituted by a direct interaction between the nucleocapsid and envelope glycoproteins. Several lines of evidence seem to substantiate this model. The CT of Gn of Uukuniemi virus was shown by alanine scanning mutagenesis to mediate the packaging of a minigenome encoding a reporter gene into virus-like particles (VLPs; Overby et al., 2007). Also, the CT domain of both Gn and Gc was shown to be indispensible for the replication of BUNV (Shi et al., 2007). A highly conserved cysteine/ histidine-rich region of the Gn tail was recently shown by NMR spectroscopy to form two classical zinc fingers (Estrada et al., 2009). This suggested an involvement of this domain in nucleic acid binding or protein–protein interactions. However, it is not yet clear whether the CT interacts with the RNA genome or the protein component of the RNP during assembly. More recently, the N protein of PUUV was shown to coimmunoprecipitate with the glycoprotein complex. Mapping of the interaction sites revealed that the N protein has multiple binding sites in the CT of Gn and was also able to bind to the predicted CT of Gc (Hepojoki et al., 2010b). c. Virulence A clear association between viral glycoproteins and virulence was established by comparing the pathogenicity of virulent and attenuated or genetically reassorted strains of HTNV in an experimental newborn mouse model. Sequence comparison identified single amino acid changes in Gn and Gc as the genetic determinants responsible for the observed difference in viral virulence (Ebihara et al., 2000; Isegawa et al., 1994). Gn consists of an external domain, a transmembrane domain and a C-termianl CT (Spiropoulou, 2001). The CT of Gn appears to be a multifunctional virulence determinant that helps the virus to evade host innate responses and ensures productive infection in human endothelial cells. It contains immunoreceptor tyrosine-based activation motifs (ITAMs). These are cell-signaling elements involved in regulating the functions of immune and endothelial cells and their presence in Gn CT suggests a direct role in hantavirus pathogenesis (Geimonen et al., 2003b). RIG-I (retinoic acid-inducible gene I) is an RNA helicase that triggers the cellular innate interferon (IFN) immune response upon detection of viral double-stranded RNA (Yoneyama et al., 2004). The Gn CT of the pathogenic New York-1 virus (NY-1 V) was shown to inhibit the RIG-I-directed IFN responses (Alff et al., 2006, 2008). To counteract this effect, it seems that cells have evolved a mechanism for downregulating Gn expression by targeting this protein for proteosomal degradation via polyubiquitination of the CT (Geimonen et al., 2003a). These findings have created some


53

controversy in the literature. It was initially reported that the Gn CTs of pathogenic (NY-1 V, HTNV, and Andes virus ANDV), but not the nonpathogenic hantavirus PHV, are exclusively degraded via the ubiquitin– proteasome pathway in COS7 cells (Sen et al., 2007). However, this view was challenged by Wang et al., who reported that the Gn CTs of the nonpathogenic TULV and PHV were proteosomally degraded in HEK293 and Vero E6 cells. Therefore, it was concluded that this degradation is not related to viral pathogenesis (Wang et al., 2009).

III. HANTAVIRUS DISEASE A. Epidemiology Unlike other members of the Bunyaviridae family, hantaviruses are not transmitted by biting insects. Human infection occurs accidentally via inhalation of contaminated rodent excreta (Hart and Bennett, 1999). Rodents are the main natural reservoir, where the virus can establish asymptomatic persistent infection. Within rodents, the disease is transmitted horizontally, mainly through aggressive behavior such as biting and scratching (Lee et al., 1981a). It is mainly a rural disease associated with the risk factors of farming, hunting, and camping because those activities bring humans into close contact with the rodent reservoirs (Simpson et al., 2010). The only hantavirus that causes disease in urban areas is SEOV because its host is the domestic rat (Mir, 2010). Human-tohuman transmission is very rare and was reported only for ANDV in Argentina (Padula et al., 1998). Approximately 150,000–200,000 cases of hantavirus infection are reported annually worldwide (Hart and Bennett, 1999). The geographic distribution of the disease reflects the distribution of the rodent host, hence the classification of hantaviruses into two main groups: Old World and New World ( Jonsson et al., 2010). Occurrence of HFRS and HCPS depends on the type of virus causing the infection (Borges et al., 2006). HFRS is primarily considered a Eurasian disease, which was first reported in Korea in 1951 and is caused by Old World hantaviruses, for example, HTNV, SEOV, DOBV, and PUUV. More than 150,000 cases of HFRS are reported annually, half of which occur in China only (Peters et al., 1999). HCPS appears to be confined to the Americas and is associated with New World hantaviruses, for example, SNV, NY-1 V, and ANDV. It was first discovered in 1993 in the Four Corners region of the southwestern United States (Nichol et al., 1993). Approximately 300 cases of HCPS are reported each year in North and South America (Muranyi et al., 2005). SNV, which infects deer mouse, is the major cause of HCPS in North America (Fabbri and Maslow, 2001).

54


B. Reservoir hosts of hantaviruses Rodents of the subfamily Murinae within family Muridae and the subfamilies Arvicolinae, Neotominae, and Sigmodontinae within family Cricetidae are considered to be the main natural reservoir hosts of hantaviruses. Old World hantaviruses, which cause HFRS, are carried by rodents of Myodes, Rattus, and Apodemus genera and New World hantaviruses, which cause HCPS are carried by the rodents of the subfamily Sigmodontinae. Recently, numerous novel hantaviruses from various sorcid and talpid insectivores have been discovered (Arai et al., 2007; Kang et al., 2009a,b; Song et al., 2007a,b,c), but none of these viruses have been associated with any human disease. An apparent coevolution of hantaviruses with their reservoir hosts is evident by the commonly observed close association of each hantavirus species with a certain rodent species. Moreover, phylogenetic analyses of hantavirus sequences and mitochondrial sequences from their rodent hosts suggest a long-standing coevolutionary history of hantaviruses with their rodent carriers ( Jackson and Charleston, 2004; Plyusnin, 2002; Plyusnin and Morzunov, 2001). Depending on the population density, up to 50% of any given population of rodents are seropostive and considered as silent carriers for hantaviruses. Through experimental infections and field surveys, it has been shown that the transmission among rodents is exclusively horizontal and it occurs via inhalation of infected aerosols, through saliva or excreta, biting, and other aggressive behavioral interactions (Botten et al., 2002; Glass et al., 1988; Hinson et al., 2004; Hutchinson et al., 2000; Kariwa et al., 1998; Lee et al., 1981a,b; Root et al., 2004). Unlike other rodent-borne viruses such as arenaviruses, there is no vertical transmission from the dam to its offspring, and maternal antibodies can protect offspring from infection for several months (Kallio et al., 2006; Taruishi et al., 2008). This view has been recently disputed by Calisher et al. (2009), and several scenarios have been presented for the transmission of hantaviruses from infected to uninfected hosts (Calisher et al., 2009). In general, infection of the rodent hosts by their respective hantaviruses is thought to be asymptomatic and no overt disease is produced. However, several recent studies have reported development of some disease symptoms in the reservoir hosts such as pulmonary edema and periportal hepatitis in white-footed mouse Peromyscus leucopus infected with NY-1 V (Lyubsky et al., 1996) and SNV-infected deer mouse Peromyscus maniculatus (Netski et al., 1999). Infection of hantavirus in the rodent host may also cause growth retardation as observed in Rattus norvegicus infected with SEOV (Childs et al., 1989) and P. maniculatus infected with SNV (Kanerva et al., 1998). Nevertheless, the general absence of an overt illness in rodent hosts despite a persistent and lifelong


55

infection is believed to be a smart way of survival for hantaviruses, by which they avoid killing their own hosts. It also highlights an amicable relationship between the virus and its host that was developed over hundreds of thousands of years of mutual interaction. The survival of rodent hosts in spite of the chronic viral infection is mediated by a combination of host’s unique immunity against the virus and the viral genetic variability, which helps it evade the host’s defense mechanisms (Schonrich et al., 2008). Persistently infected rodents shed the virus in urine, feces, and saliva. Inhalation of virus-contaminated aerosols is the major route of transmission to humans. However, the general view that rodents are the only infection source for humans has been disputed by Zeier et al. (2005). It has been suggested that the close proximity of domestic animals such as cats, dogs, pigs, and cattle with rodents may cause some transmission events of hantaviruses to these animals. This species jump may cause the virus to evolve differently and result in more dangerous forms of the virus, which could be more pathogenic to humans (Zeier et al., 2005).

C. Diseases caused by hantaviruses in humans In contrast to the asymptomatic infection of the rodent reservoir, hantavirus infection to humans results in two disease forms: HFRS and HCPS. In both syndromes, vascular endothelial cells show increased permeability and both are believed to result from host immune responses to infection, rather than damage caused by the viruses themselves (Khaiboullina and St. Jeor, 2002).

1. Hemorrhagic fever with renal syndrome The incubation period of HFRS is about 3 weeks but can range from 10 days to 6 weeks ( Jonsson et al., 2010; Kramski et al., 2009). The clinical course is classically subdivided into five overlapping phases: febrile, hypotensive, oliguric, diuretic, and convalescent (Schmaljohn and Hjelle, 1997). The febrile phase lasts for 3–5 days and is characterized by flu-like symptoms, thirst, nausea, and vomiting. The febrile phase is followed by a hypotensive phase, which lasts for few hours to 2 days, and is characterized by thrombocytopenia and petechial hemorrhage, retroperitoneal edema, and abdominal pain. The subsequent oliguric phase lasts for few days to two weeks. The mortality rate is 15%, with the majority of deaths occurring during the hypotensive and oliguric phases due to the complications of vascular leakage, renal failure, and acute shock. Patients who survive these complications usually progress into the diuretic phase and show improved renal function, with subsequent convalescence and recovery (Peters et al., 1999).

56


2. Hantavirus cardiopulmonary syndrome The incubation period of HCPS ranges from 9 to 33 days (Young et al., 2000). The clinical disease course of HCPS is usually divided into prodromal, cardiopulmonary, and diuretic or convalescent phases. The prodromal stage, which usually lasts 3–6 days, is characterized by flu-like symptoms such as fever, headache, chills, and muscle pain. Abdominal pain, nausea, vomiting, and dizziness may occur. This phase usually progresses rapidly to a severe respiratory disease characterized by nonproductive cough and dyspnea. In contrast to HFRS, the increased permeability and fluid leakage occur exclusively in the lungs instead of the kidneys. This bilateral pulmonary edema is usually visible in chest X-rays (Boroja et al., 2002). Thrombocytopenia, hemoconcentration, and leukocytosis are the most prominent hematologic findings (Simpson et al., 2010). Rapid deterioration, cardiac insufficiency, and respiratory failure caused by edema and hypotention, shock, and death may occur within 2–10 days after the onset of illness in almost 50% of cases. Patients who survive the acute phase recover within 5–7 days and enter the diuretic phase (Nolte et al., 1995; Zaki et al., 1995).

D. Pathogenesis Hantavirus infection occurs via inhalation of infectious virus particles into the lungs. The inhaled viruses bind to b3 integrin receptors on the surface of pulmonary endothelial cells, macrophages, and dendritic cells (DCs), where virus replication mainly occurs (Gavrilovskaya et al., 1998). Capillary leakage is the hallmark of hantavirus infection; however, the sequence of events from inhalation of infectious virus particles until pulmonary capillary leakage remains poorly understood. It was proposed that DCs play the role of a Trojan horse by helping the dissemination of virus throughout the body. Immature DCs located near the respiratory epithelial cells engulf inhaled virus particles, which can replicate in those cells without causing cell death. During their maturation, DCs migrate from the lungs to the lymph nodes where they present the engulfed antigen to immune cells (Raftery et al., 2002). The mechanisms by which pathogenic hantaviruses cause capillary leakage is an area of active research. Accumulating pieces of experimental evidences suggest that multiple immunopathologic mechanisms rather than direct viral-induced cytopathic effects are responsible for the disruption of vascular endothelium and the subsequent capillary leakage associated with hantavirus diseases (Borges et al., 2006; Sundstrom et al., 2001). In PUUV-infected patients, elevated levels of serum lactate dehydrogenase (LDH), indicative of cellular damage, correlated with high levels of serum perforin, granzyme B, and epithelial cell apoptosis markers. These


57

findings suggested that tissue damage is due to an immune reaction and that epithelial apoptosis contributed significantly to the damage (Klingstrom et al., 2006b). Some changes in the endothelial cell barrier functions might be related to the use of b3 integrins as receptors. It was shown that the initial interaction of the infecting virus with b3 integrin receptors disturbs the integrin-directed migration of endothelial cells, which is essential for maintaining vascular integrity (Gavrilovskaya et al., 2002). The cellular immune response to infection involves strong stimulation of hantavirusspecific cytotoxic CD8 T lymphocytes (CTLs), which are released in large numbers into the blood. The severity of the disease was shown to correlate with the number of CTLs in the blood (Kilpatrick et al., 2004). This vast excess of activated CTLs is partly due to the lack of downregulation of T-cell function as evidenced by low serum levels of cytokines released by regulatory T cells, such as TGF-b, in HCPS patients (Chen and Yang, 1990; Mills, 2004). The intense antiviral immune response can contribute to the increased permeability of endothelial cells in two ways. First, the elevated levels of inflammatory mediators and cytokines, such as TNF-a, IL-6, and IL-10, released during the acute phase by activated T cells and infected endothelial cells increase the vascular permeability resulting in pulmonary edema (Linderholm et al., 1996). Second, it was shown in transwell permeability assays that hantavirus-specific cytotoxic T lymphocytes could directly lyse human endothelial cells infected with SNV (Hayasaka et al., 2007). The exact mechanism of kidney failure in HFRS is unclear. Renal disease has been attributed to deposition of immune complexes and the presence of inflammatory cell infiltrations with subsequent tubular damage (McCaughey and Hart, 2000; Sironen et al., 2008).

E. Diagnosis Diagnosis of hantavirus infection in the clinic relies on the establishment of a history of rodent exposure, symptoms suggestive of respiratory or renal involvement, blood exams showing severe thrombocytopenia, and positive serological tests. Virus isolation from clinical samples is difficult due to the presence of high levels of neutralizing antibodies (McCaughey and Hart, 2000). High titers of IgM and IgG antibodies against hantavirus N and Gn proteins are detectable in the sera of patients during the acute phase, enabling reliable serological confirmation of infection (Bostik et al., 2000; Elgh et al., 1997; Groen et al., 1994). SNV Gn antibodies are highly specific and do not cross-react with Gn antigens of other hantaviruses (Hjelle et al., 1994; Jenison et al., 1994). The most widely used serological tests for diagnosis of HCPS are IgM capture and IgG indirect ELISAs (Li et al., 2002). A rapid test for SNV and SEOV in the form of a strip

58


immunoblot assay has been developed, which relies on antigens derived from recombinant SNV N and Gn proteins and SEOV N protein. The assay was demonstrated to be highly sensitive and specific because it could identify all patients with acute SNV infection during the early course of the disease with no false-positive results (Hjelle et al., 1997). Immunohistochemical detection of hantavirus antigens has been particularly useful in establishing retrospective diagnosis of HCPS from fixed tissue samples (Peters et al., 1999). RT-PCR assays have also been used, but results should be always interpreted with caution due to the potential of cross-contamination and have to be always substantiated by positive results from immunodiagnostic assays (Moreli et al., 2004).

F. Therapy No specific antiviral therapy is currently available for treating hantavirus infections. Management of infected patients relies solely on supportive therapy in an intensive care unit until the virus is cleared by the immune system (Muranyi et al., 2005). Ribavirin, a broad-spectrum nucleoside analogue antiviral drug, has been shown to possess, both in vitro and in vivo, inhibitory activity against hantaviruses (Huggins et al., 1986; Medina et al., 2007; Severson et al., 2003). A clinical trial showed decreased virus titers, morbidity, and mortality rates in HFRS Chinese patients (Huggins et al., 1991). Another trial on HFRS patients in Korea showed significant reduction in fatality and improved prognosis when given early in the course of the disease, as well as a reduction in the risk of renal insufficiency (Rusnak et al., 2009). Intravenous injection of ribavirin was generally well tolerated by HCPS patients (Chapman et al., 1999). However, a Placebo-controlled, double-blind trial of intravenous ribavirin in HCPS patients was ended prematurely due to low rate of enrollment (Mertz et al., 2004). Therefore, ribavirin does not seem to have any clinical application in HCPS patients due to the lack of conclusive clinical data. Passive administration of neutralizing antibodies for PUUV has been shown to protect macaques against virus challenge; however, no human clinical trials were conducted (Klingstrom et al., 2005).

G. Vaccines Despite many efforts that have been invested in developing a safe effective vaccine to protect against hantavirus infection, currently, no FDAapproved vaccine is available in the United States (Ulrich et al., 2002). A formalin-inactivated HTNV vaccine (Hantavax) produced from mouse brain-derived virus is licensed for use in Korea since the 1990s. Efficacy studies showed that 97% of human volunteers receiving this vaccine developed high titers of specific neutralizing antibodies after a booster


59

dose; however, the induced humoral immune response was short lived (Cho and Howard, 1999). Another formaline-inactivated vaccine produced from suspensions of Vero cells elicited a neutralizing antibody response in mice, which was fivefold higher than Hantavax (Choi et al., 2003). However, it is unlikely that these vaccines can provide protection against all hantaviruses. The N, Gn, and Gc proteins of hantaviruses are highly immunogenic (Bharadwaj et al., 2002). Therefore, different approaches were taken to develop recombinant vaccines using proteins expressed in baculovirus, vaccinia virus, and the yeast Saccharomyces cerevisiae (Chu et al., 1995; Schmaljohn et al., 1990; Yoshimatsu et al., 1993). Recombinant N proteins of PUUV and DOBV expressed in yeast were shown to induce a protective immune response in rodent models (Dargeviciute et al., 2002; Geldmacher et al., 2004). Moreover, deer mice immunized with recombinant deer mouse cytomegalovirus (PCMV) expressing SNV Gn developed an antibody response to SNV (Rizvanov et al., 2003). Although highly protective in mouse models, a recombinant vaccinia virus containing the S and M segments of HTNV proved to be inefficient in a phase II clinical trial. It elicited neutralizing antibodies in only 72% of the 142 participating volunteers (McClain et al., 2000). Several other studies using naked DNA vaccines containing the M or S genome segments of SNV and SEOV have shown similar success in rodent models, but no clinical administration has been reported (Bharadwaj et al., 1999; Hooper et al., 1999).

IV. FUTURE PROSPECTS Frequent emergence of zoonotic viruses is a serious concern to human health. For example, the recent emergence of H1N1 (swine flu) has created havoc in human lives, especially in pregnant women and young children who were predicted to be most susceptible for this new virus species. History witnesses the emergence and reemergence of numerous pathogens in the past century, with an estimated frequency of one new pathogen every 18 months. Many of these new pathogens were zoonotic RNA viruses, such as hantavirus, H1NI swine flu, Nipah virus, Hendra virus, Ebola virus, West Nile virus, and SARS (severe acute respiratory syndrome cornovirus; Jones et al., 2008; Morens et al., 2004). Despite their significant devastation to human lives, the ecology and natural history of these zoonotic viruses are still a mystery. The principles that govern their stable maintenance in their natural reservoirs, compulsions for switching their hosts, mechanisms of recombination and reassortment, molecular mechanisms of their transfer, survival and adaptation in the new host, are the interesting areas that remain poorly understood. The natural hosts of these zoonotic viruses continue to be the reservoirs for the generation of

60


new pathogenic strains and probable causes of future pandemics that pose serious threat to human lives. Hantaviruses cause HFRS and HCPS when transmitted to humans but maintain a persistent infection in their rodent hosts. Coevolutionary existence of hantaviruses and their rodent hosts provides insights into mechanisms by which these zonotic viruses manage to exist in the environment for millions of years with a potential to infect humans upon transmission. However, understanding the mechanisms by which hantaviruses maintain a persistent infection in their rodent hosts without a disease may provide insights into the possible approaches that could be used for the treatment of hantavirus-associated disease in humans. Excessive proinflammatory cytokine (TNF-a) and CD8þ responses in humans are hypothesized to mediate the pathogenesis of human HFRS and HCPS, suggesting that anti-TNFa therapy might be helpful during the treatment. Since HCPS is comparatively a rare disease, funding for the development of an HCPS vaccine is not a high priority for many countries. However, with the increasing number of HCPS cases, especially in South America, vaccine efforts for HCPS may receive better appreciation. Due to the requirements of high-containment laboratories for HCPScausing viruses, plasmid DNA gene gun-based approaches will be preferred for the development of an HCPS vaccine. Dissecting the molecular mechanisms of hantavirus replication in host cells will provide insights about the new potential targets for the design of antiviral therapeutic agents for the treatment of HFRS and HCPS. For example, the newly discovered translation initiation mechanism, operated by hantavirus nucleocapsid protein, demonstrates how a single viral protein lures the host cell translation machinery for the preferential translation of viral mRNAs in virus-infected cells where cellular transcripts are competing for the same translation apparatus (Mir and Panganiban, 2008). Shutting down this viral translation initiation strategy might inhibit virus replication in infected cells. In addition, viral RdRp uses capped RNA primers to initiate the transcription/replication of viral RNA genome. Capped RNA primers are generated from host cell mRNAs by a unique ‘‘cap-snatching mechanism.’’ This mechanism is well understood in influenza virus. However, recent studies suggest that hantaviral cap-snatching mechanism might be different from influenza virus (Mir et al., 2008). In-depth understanding of cap-snatching mechanism will reveal novel targets that could be used for the design of antiviral agents for the treatment of diseases caused by a broad spectrum of viruses that use cap-snatching mechanism to initiate transcription/replication of viral genome. Currently, the studies of hantavirus biology and pathogenesis are limited due to the lack of an animal model and a complete reverse genetic system for these viruses, although a minigenome system for HTNV has


61

been reported (Flick et al., 2003). Reverse genetic system will generate hopes for screening chemical libraries to identify molecules with antiviral therapeutic potential, for the development of attenuated vaccine candidates, and for the investigation of molecular mechanisms involved in hantavirus replication, gene expression, as well as virus assembly and budding.

REFERENCES Alfadhli, A., Love, Z., Arvidson, B., Seeds, J., Willey, J., and Barklis, E. (2001). Hantavirus nucleocapsid protein oligomerization. J. Virol. 75, 2019–2023. Alfadhli, A., Steel, E., Finlay, L., Bachinger, H. P., and Barklis, E. (2002). Hantavirus nucleocapsid protein coiled-coil domains. J. Biol. Chem. 277, 27103–27108. Alff, P. J., Gavrilovskaya, I. N., Gorbunova, E., Endriss, K., Chong, Y., Geimonen, E., Sen, N., Reich, N. C., and Mackow, E. R. (2006). The pathogenic NY-1 hantavirus G1 cytoplasmic tail inhibits RIG-I- and TBK-1-directed interferon responses. J. Virol. 80, 9676–9686. Alff, P. J., Sen, N., Gorbunova, E., Gavrilovskaya, I. N., and Mackow, E. R. (2008). The NY-1 hantavirus Gn cytoplasmic tail coprecipitates TRAF3 and inhibits cellular interferon responses by disrupting TBK1–TRAF3 complex formation. J. Virol. 82, 9115–9122. Andersson, A. M., Melin, L., Bean, A., and Pettersson, R. F. (1997). A retention signal necessary and sufficient for Golgi localization maps to the cytoplasmic tail of a Bunyaviridae (Uukuniemi virus) membrane glycoprotein. J. Virol. 71, 4717–4727. Antic, D., Kang, C. Y., Spik, K., Schmaljohn, C., Vapalahti, O., and Vaheri, A. (1992a). Comparison of the deduced gene products of the L, M and S genome segments of hantaviruses. Virus Res. 24, 35–46. Antic, D., Wright, K. E., and Kang, C. Y. (1992b). Maturation of Hantaan virus glycoproteins G1 and G2. Virology 189, 324–328. Aquino, V. H., Moreli, M. L., and Moraes Figueiredo, L. T. (2003). Analysis of oropouche virus L protein amino acid sequence showed the presence of an additional conserved region that could harbour an important role for the polymerase activity. Arch. Virol. 148, 19–28. Arai, S., Song, J. W., Sumibcay, L., Bennett, S. N., Nerurkar, V. R., Parmenter, C., Cook, J. A., Yates, T. L., and Yanagihara, R. (2007). Hantavirus in northern short-tailed shrew, United States. Emerg. Infect. Dis. 13, 1420–1423. Arikawa, J., Schmaljohn, A. L., Dalrymple, J. M., and Schmaljohn, C. S. (1989). Characterization of Hantaan virus envelope glycoprotein antigenic determinants defined by monoclonal antibodies. J. Gen. Virol. 70(Pt. 3), 615–624. Arnold, J. J., and Cameron, C. E. (2004). Poliovirus RNA-dependent RNA polymerase (3Dpol): Pre-steady-state kinetic analysis of ribonucleotide incorporation in the presence of Mg2þ. Biochemistry 43, 5126–5137. Arnold, J. J., Ghosh, S. K., and Cameron, C. E. (1999). Poliovirus RNA-dependent RNA polymerase (3D(pol)). Divalent cation modulation of primer, template, and nucleotide selection. J. Biol. Chem. 274, 37060–37069. Avsic-Zupanc, T., Toney, A., Anderson, K., Chu, Y. K., and Schmaljohn, C. (1995). Genetic and antigenic properties of Dobrava virus: A unique member of the Hantavirus genus, family Bunyaviridae. J. Gen. Virol. 76(Pt. 11), 2801–2808. Avsic-Zupanc, T., Nemirov, K., Petrovec, M., Trilar, T., Poljak, M., Vaheri, A., and Plyusnin, A. (2000). Genetic analysis of wild-type Dobrava hantavirus in Slovenia: Coexistence of two distinct genetic lineages within the same natural focus. J. Gen. Virol. 81, 1747–1755.

62


Barclay, C. M., and Rubinstein, G. (1997). Hantavirus in Argentina. Nature 386, 320. Barr, J. N., and Fearns, R. (2010). How RNA viruses maintain their genome integrity. J. Gen. Virol. 91, 1373–1387. Bayard, V., Kitsutani, P. T., Barria, E. O., Ruedas, L. A., Tinnin, D. S., Munoz, C., de Mosca, I. B., Guerrero, G., Kant, R., Garcia, A., Caceres, L., Gracio, F. G., et al. (2004). Outbreak of hantavirus pulmonary syndrome, Los Santos, Panama, 1999–2000. Emerg. Infect. Dis. 10, 1635–1642. Beckham, C. J., and Parker, R. (2008). P bodies, stress granules, and viral life cycles. Cell Host Microbe 3, 206–212. Bharadwaj, M., Lyons, C. R., Wortman, I. A., and Hjelle, B. (1999). Intramuscular inoculation of Sin Nombre hantavirus cDNAs induces cellular and humoral immune responses in BALB/c mice. Vaccine 17, 2836–2843. Bharadwaj, M., Mirowsky, K., Ye, C., Botten, J., Masten, B., Yee, J., Lyons, C. R., and Hjelle, B. (2002). Genetic vaccines protect against Sin Nombre hantavirus challenge in the deer mouse (Peromyscus maniculatus). J. Gen. Virol. 83, 1745–1751. Bishop, D. H., Gay, M. E., and Matsuoko, Y. (1983). Nonviral heterogeneous sequences are present at the 5’ ends of one species of snowshoe hare bunyavirus S complementary RNA. Nucleic Acids Res. 11, 6409–6418. Blakqori, G., Kochs, G., Haller, O., and Weber, F. (2003). Functional L polymerase of La Crosse virus allows in vivo reconstitution of recombinant nucleocapsids. J. Gen. Virol. 84, 1207–1214. Bohlman, M. C., Morzunov, S. P., Meissner, J., Taylor, M. B., Ishibashi, K., Rowe, J., Levis, S., Enria, D., and St. Jeor, S. C. (2002). Analysis of hantavirus genetic diversity in Argentina: S segment-derived phylogeny. J. Virol. 76, 3765–3773. Borges, A. A., Campos, G. M., Moreli, M. L., Souza, R. L., Aquino, V. H., Saggioro, F. P., and Figueiredo, L. T. (2006). Hantavirus cardiopulmonary syndrome: Immune response and pathogenesis. Microbes Infect. 8, 2324–2330. Boroja, M., Barrie, J. R., and Raymond, G. S. (2002). Radiographic findings in 20 patients with Hantavirus pulmonary syndrome correlated with clinical outcome. AJR Am. J. Roentgenol. 178, 159–163. Bostik, P., Winter, J., Ksiazek, T. G., Rollin, P. E., Villinger, F., Zaki, S. R., Peters, C. J., and Ansari, A. A. (2000). Sin nombre virus (SNV) Ig isotype antibody response during acute and convalescent phases of hantavirus pulmonary syndrome. Emerg. Infect. Dis. 6, 184–187. Botten, J., Mirowsky, K., Ye, C., Gottlieb, K., Saavedra, M., Ponce, L., and Hjelle, B. (2002). Shedding and intracage transmission of Sin Nombre hantavirus in the deer mouse (Peromyscus maniculatus) model. J. Virol. 76, 7587–7594. Bouloy, M., Plotch, S. J., and Krug, R. M. (1978). Globin mRNAs are primers for the transcription of influenza viral RNA in vitro. Proc. Natl. Acad. Sci. USA 75, 4886–4890. Bressanelli, S., Tomei, L., Rey, F. A., and De Francesco, R. (2002). Structural analysis of the hepatitis C virus RNA polymerase in complex with ribonucleotides. J. Virol. 76, 3482–3492. Bridgen, A., and Elliott, R. M. (1996). Rescue of a segmented negative-strand RNA virus entirely from cloned complementary DNAs. Proc. Natl. Acad. Sci. USA 93, 15400–15404. Bruenn, J. A. (2003). A structural and primary sequence comparison of the viral RNAdependent RNA polymerases. Nucleic Acids Res. 31, 1821–1829. Butcher, S. J., Grimes, J. M., Makeyev, E. V., Bamford, D. H., and Stuart, D. I. (2001). A mechanism for initiating RNA-dependent RNA polymerization. Nature 410, 235–240. Calisher, C. H., Peters, C. J., Douglass, R. J., and Kuenzi, A. J. (2009). Hantaviral infections of rodents: Possible scenarios. Arch. Virol. 154, 1195–1197. Cevik, B., Smallwood, S., and Moyer, S. A. (2003). The L-L oligomerization domain resides at the very N-terminus of the sendai virus L RNA polymerase protein. Virology 313, 525–536.


63

Cevik, B., Holmes, D. E., Vrotsos, E., Feller, J. A., Smallwood, S., and Moyer, S. A. (2004). The phosphoprotein (P) and L binding sites reside in the N-terminus of the L subunit of the measles virus RNA polymerase. Virology 327, 297–306. Chan, Y. C., Wong, T. W., and Yap, E. H. (1987). Haemorrhagic fever with renal syndrome: Clinical, virological and epidemiological perspectives. Ann. Acad. Med. Singapore 16, 696–701. Chapman, L. E., Mertz, G. J., Peters, C. J., Jolson, H. M., Khan, A. S., Ksiazek, T. G., Koster, F. T., Baum, K. F., Rollin, P. E., Pavia, A. T., Holman, R. C., Christenson, J. C., et al. (1999). Intravenous ribavirin for hantavirus pulmonary syndrome: Safety and tolerance during 1 year of open-label experience. Ribavirin Study Group. Antivir. Ther. 4, 211–219. Chen, L. B., and Yang, W. S. (1990). Abnormalities of T cell immunoregulation in hemorrhagic fever with renal syndrome. J. Infect. Dis. 161, 1016–1019. Childs, J. E., Glass, G. E., Korch, G. W., and LeDuc, J. W. (1989). Effects of hantaviral infection on survival, growth and fertility in wild rat (Rattus norvegicus) populations of Baltimore, Maryland. J. Wildl. Dis. 25, 469–476. Chizhikov, V. E., Spiropoulou, C. F., Morzunov, S. P., Monroe, M. C., Peters, C. J., and Nichol, S. T. (1995). Complete genetic characterization and analysis of isolation of Sin Nombre virus. J. Virol. 69, 8132–8136. Cho, H. W., and Howard, C. R. (1999). Antibody responses in humans to an inactivated hantavirus vaccine (Hantavax). Vaccine 17, 2569–2575. Cho, H. W., Howard, C. R., and Lee, H. W. (2002). Review of an inactivated vaccine against hantaviruses. Intervirology 45, 328–333. Choi, Y., Ahn, C. J., Seong, K. M., Jung, M. Y., and Ahn, B. Y. (2003). Inactivated Hantaan virus vaccine derived from suspension culture of Vero cells. Vaccine 21, 1867–1873. Chu, Y. K., Jennings, G. B., and Schmaljohn, C. S. (1995). A vaccinia virus-vectored Hantaan virus vaccine protects hamsters from challenge with Hantaan and Seoul viruses but not Puumala virus. J. Virol. 69, 6417–6423. Chu, Y. K., Milligan, B., Owen, R. D., Goodin, D. G., and Jonsson, C. B. (2006). Phylogenetic and geographical relationships of hantavirus strains in eastern and western Paraguay. Am. J. Trop. Med. Hyg. 75, 1127–1134. Clement, J., Lameire, N., Keyaerts, E., Maes, P., and Van Ranst, M. (2003). Hantavirus infections in Europe. Lancet Infect. Dis. 3, 752–753 discussion 753–754. Copper, P. D., Steiner-Pryor, A., Scotti, P. D., and Delong, D. (1974). On the nature of poliovirus genetic recombinants. J. Gen. Virol. 23, 41–49. Crotty, S., and Andino, R. (2002). Implications of high RNA virus mutation rates: Lethal mutagenesis and the antiviral drug ribavirin. Microbes Infect. 4, 1301–1307. Dahlberg, J. E., Obijeski, J. F., and Korb, J. (1977). Electron microscopy of the segmented RNA genome of La Crosse virus: Absence of circular molecules. J. Virol. 22, 203–209. Dargeviciute, A., Brus Sjolander, K., Sasnauskas, K., Kruger, D. H., Meisel, H., Ulrich, R., and Lundkvist, A. (2002). Yeast-expressed Puumala hantavirus nucleocapsid protein induces protection in a bank vole model. Vaccine 20, 3523–3531. de Carvalho Nicacio, C., Sallberg, M., Hultgren, C., and Lundkvist, A. (2001). T-helper and humoral responses to Puumala hantavirus nucleocapsid protein: Identification of Thelper epitopes in a mouse model. J. Gen. Virol. 82, 129–138. de Haan, P., Kormelink, R., de Oliveira Resende, R., van Poelwijk, F., Peters, D., and Goldbach, R. (1991). Tomato spotted wilt virus L RNA encodes a putative RNA polymerase. J. Gen. Virol. 72(Pt. 9), 2207–2216. Deyde, V. M., Rizvanov, A. A., Chase, J., Otteson, E. W., and St. Jeor, S. C. (2005). Interactions and trafficking of Andes and Sin Nombre Hantavirus glycoproteins G1 and G2. Virology 331, 307–315.

64


Doublie, S., and Ellenberger, T. (1998). The mechanism of action of T7 DNA polymerase. Curr. Opin. Struct. Biol. 8, 704–712. Drake, J. W. (1999). The distribution of rates of spontaneous mutation over viruses, prokaryotes, and eukaryotes. Ann. NY Acad. Sci. 870, 100–107. Dunn, E. F., Pritlove, D. C., Jin, H., and Elliott, R. M. (1995). Transcription of a recombinant bunyavirus RNA template by transiently expressed bunyavirus proteins. Virology 211, 133–143. Ebihara, H., Yoshimatsu, K., Ogino, M., Araki, K., Ami, Y., Kariwa, H., Takashima, I., Li, D., and Arikawa, J. (2000). Pathogenicity of Hantaan virus in newborn mice: Genetic reassortant study demonstrating that a single amino acid change in glycoprotein G1 is related to virulence. J. Virol. 74, 9245–9255. Elgh, F., Lundkvist, A., Alexeyev, O. A., Wadell, G., and Juto, P. (1996). A major antigenic domain for the human humoral response to Puumala virus nucleocapsid protein is located at the amino-terminus. J. Virol. Methods 59, 161–172. Elgh, F., Lundkvist, A., Alexeyev, O. A., Stenlund, H., Avsic-Zupanc, T., Hjelle, B., Lee, H. W., Smith, K. J., Vainionpaa, R., Wiger, D., Wadell, G., and Juto, P. (1997). Serological diagnosis of hantavirus infections by an enzyme-linked immunosorbent assay based on detection of immunoglobulin G and M responses to recombinant nucleocapsid proteins of five viral serotypes. J. Clin. Microbiol. 35, 1122–1130. Elliott, L. H., Kiley, M. P., and McCormick, J. B. (1984). Hantaan virus: Identification of virion proteins. J. Gen. Virol. 65(Pt. 8), 1285–1293. Eshita, Y., Ericson, B., Romanowski, V., and Bishop, D. H. (1985). Analyses of the mRNA transcription processes of snowshoe hare bunyavirus S and M RNA species. J. Virol. 55, 681–689. Estrada, D. F., Boudreaux, D. M., Zhong, D., St. Jeor, S. C., and De Guzman, R. N. (2009). The hantavirus glycoprotein G1 tail contains dual CCHC-type classical zinc fingers. J. Biol. Chem. 284, 8654–8660. Fabbri, M., and Maslow, M. J. (2001). Hantavirus pulmonary syndrome in the United States. Curr. Infect. Dis. Rep. 3, 258–265. Fazakerley, J. K., and Ross, A. M. (1989). Computer analysis suggests a role for signal sequences in processing polyproteins of enveloped RNA viruses and as a mechanism of viral fusion. Virus Genes 2, 223–239. Flick, R., and Pettersson, R. F. (2001). Reverse genetics system for Uukuniemi virus (Bunyaviridae): RNA polymerase I-catalyzed expression of chimeric viral RNAs. J. Virol. 75, 1643–1655. Flick, K., Hooper, J. W., Schmaljohn, C. S., Pettersson, R. F., Feldmann, H., and Flick, R. (2003). Rescue of Hantaan virus minigenomes. Virology 306, 219–224. Fulhorst, C. F., Monroe, M. C., Salas, R. A., Duno, G., Utrera, A., Ksiazek, T. G., Nichol, S. T., de Manzione, N. M., Tovar, D., and Tesh, R. B. (1997). Isolation, characterization and geographic distribution of Cano Delgadito virus, a newly discovered South American hantavirus (family Bunyaviridae). Virus Res. 51, 159–171. Garcin, D., Lezzi, M., Dobbs, M., Elliott, R. M., Schmaljohn, C., Kang, C. Y., and Kolakofsky, D. (1995). The 5’ ends of Hantaan virus (Bunyaviridae) RNAs suggest a prime-and-realign mechanism for the initiation of RNA synthesis. J. Virol. 69, 5754–5762. Gavrilovskaya, I. N., Shepley, M., Shaw, R., Ginsberg, M. H., and Mackow, E. R. (1998). beta3 Integrins mediate the cellular entry of hantaviruses that cause respiratory failure. Proc. Natl. Acad. Sci. USA 95, 7074–7079. Gavrilovskaya, I. N., Brown, E. J., Ginsberg, M. H., and Mackow, E. R. (1999). Cellular entry of hantaviruses which cause hemorrhagic fever with renal syndrome is mediated by beta3 integrins. J. Virol. 73, 3951–3959.


65

Gavrilovskaya, I. N., Peresleni, T., Geimonen, E., and Mackow, E. R. (2002). Pathogenic hantaviruses selectively inhibit beta3 integrin directed endothelial cell migration. Arch. Virol. 147, 1913–1931. Geimonen, E., Fernandez, I., Gavrilovskaya, I. N., and Mackow, E. R. (2003a). Tyrosine residues direct the ubiquitination and degradation of the NY-1 hantavirus G1 cytoplasmic tail. J. Virol. 77, 10760–10868. Geimonen, E., LaMonica, R., Springer, K., Farooqui, Y., Gavrilovskaya, I. N., and Mackow, E. R. (2003b). Hantavirus pulmonary syndrome-associated hantaviruses contain conserved and functional ITAM signaling elements. J. Virol. 77, 1638–1643. Geldmacher, A., Schmaler, M., Kruger, D. H., and Ulrich, R. (2004). Yeast-expressed hantavirus Dobrava nucleocapsid protein induces a strong, long-lasting, and highly crossreactive immune response in mice. Viral Immunol. 17, 115–122. Gerrard, S. R., and Nichol, S. T. (2002). Characterization of the Golgi retention motif of Rift Valley fever virus G(N) glycoprotein. J. Virol. 76, 12200–12210. Glass, G. E., Childs, J. E., Korch, G. W., and LeDuc, J. W. (1988). Association of intraspecific wounding with hantaviral infection in wild rats (Rattus norvegicus). Epidemiol. Infect. 101, 459–472. Gohara, D. W., Crotty, S., Arnold, J. J., Yoder, J. D., Andino, R., and Cameron, C. E. (2000). Poliovirus RNA-dependent RNA polymerase (3Dpol): Structural, biochemical, and biological analysis of conserved structural motifs A and B. J. Biol. Chem. 275, 25523–25532. Gohara, D. W., Arnold, J. J., and Cameron, C. E. (2004). Poliovirus RNA-dependent RNA polymerase (3Dpol): Kinetic, thermodynamic, and structural analysis of ribonucleotide selection. Biochemistry 43, 5149–5158. Goldsmith, C. S., Elliott, L. H., Peters, C. J., and Zaki, S. R. (1995). Ultrastructural characteristics of Sin Nombre virus, causative agent of hantavirus pulmonary syndrome. Arch. Virol. 140, 2107–2122. Golovljova, I., Vasilenko, V., Prukk, T., Brus Sjolander, K., Plyusnin, A., and Lundkvist, A. (2000). Puumala and Dobrava hantaviruses causing hemorrhagic fever with renal syndrome in Estonia. Eur. J. Clin. Microbiol. Infect. Dis. 19, 968–969. Gott, P., Stohwasser, R., Schnitzler, P., Darai, G., and Bautz, E. K. (1993). RNA binding of recombinant nucleocapsid proteins of hantaviruses. Virology 194, 332–337. Gott, P., Zoller, L., Darai, G., and Bautz, E. K. (1997). A major antigenic domain of hantaviruses is located on the aminoproximal site of the viral nucleocapsid protein. Virus Genes 14, 31–40. Groen, J., Gerding, M., Jordans, J. G., Clement, J. P., and Osterhaus, A. D. (1994). Class and subclass distribution of Hantavirus-specific serum antibodies at different times after the onset of nephropathia epidemica. J. Med. Virol. 43, 39–43. Handke, W., Oelschlegel, R., Franke, R., Wiedemann, L., Kruger, D. H., and Rang, A. (2010). Generation and characterisation of genetic reassortants between Puumala and Prospect Hill hantavirus in vitro. J. Gen. Virol. 91, 2351–2359. Hansen, J. L., Long, A. M., and Schultz, S. C. (1997). Structure of the RNA-dependent RNA polymerase of poliovirus. Structure 5, 1109–1122. Hart, C. A., and Bennett, M. (1999). Hantavirus infections: Epidemiology and pathogenesis. Microbes Infect. 1, 1229–1237. Hass, M., Lelke, M., Busch, C., Becker-Ziaja, B., and Gunther, S. (2008). Mutational evidence for a structural model of the Lassa virus RNA polymerase domain and identification of two residues, Gly1394 and Asp1395, that are critical for transcription but not replication of the genome. J. Virol. 82, 10207–10217. Hayasaka, D., Maeda, K., Ennis, F. A., and Terajima, M. (2007). Increased permeability of human endothelial cell line EA.hy926 induced by hantavirus-specific cytotoxic T lymphocytes. Virus Res. 123, 120–127.

66


Henderson, W. W., Monroe, M. C., St. Jeor, S. C., Thayer, W. P., Rowe, J. E., Peters, C. J., and Nichol, S. T. (1995). Naturally occurring Sin Nombre virus genetic reassortants. Virology 214, 602–610. Hepojoki, J., Strandin, T., Vaheri, A., and Lankinen, H. (2010a). Interactions and oligomerization of hantavirus glycoproteins. J. Virol. 84, 227–242. Hepojoki, J. M., Strandin, T., Wang, H., Vapalahti, O., Vaheri, A., and Lankinen, H. (2010b). The cytoplasmic tails of hantavirus glycoproteins interact with the nucleocapsid protein. J. Gen. Virol. 91, 2341–2350. Hewlett, M. J., Pettersson, R. F., and Baltimore, D. (1977). Circular forms of Uukuniemi virion RNA: An electron microscopic study. J. Virol. 21, 1085–1093. Hinson, E. R., Shone, S. M., Zink, M. C., Glass, G. E., and Klein, S. L. (2004). Wounding: The primary mode of Seoul virus transmission among male Norway rats. Am. J. Trop. Med. Hyg. 70, 310–317. Hjelle, B., Chavez-Giles, F., Torrez-Martinez, N., Yamada, T., Sarisky, J., Ascher, M., and Jenison, S. (1994). Dominant glycoprotein epitope of four corners hantavirus is conserved across a wide geographical area. J. Gen. Virol. 75(Pt. 11), 2881–2888. Hjelle, B., Torrez-Martinez, N., and Koster, F. T. (1996). Hantavirus pulmonary syndromerelated virus from Bolivia. Lancet 347, 57. Hjelle, B., Jenison, S., Torrez-Martinez, N., Herring, B., Quan, S., Polito, A., Pichuantes, S., Yamada, T., Morris, C., Elgh, F., Lee, H. W., Artsob, H., et al. (1997). Rapid and specific detection of Sin Nombre virus antibodies in patients with hantavirus pulmonary syndrome by a strip immunoblot assay suitable for field diagnosis. J. Clin. Microbiol. 35, 600–608. Hooper, J., and Schmaljohn, C. S. (2001). Hantavirus. In ‘‘The Springer Index of Viruses’’ (C. A. Tidona and G. Darai, Eds.), pp. 150–156. Springer-Verlag, Berlin Heidelberg New York. Hooper, J. W., Kamrud, K. I., Elgh, F., Custer, D., and Schmaljohn, C. S. (1999). DNA vaccination with hantavirus M segment elicits neutralizing antibodies and protects against seoul virus infection. Virology 255, 269–278. Huang, H., Chopra, R., Verdine, G. L., and Harrison, S. C. (1998). Structure of a covalently trapped catalytic complex of HIV-1 reverse transcriptase: Implications for drug resistance. Science 282, 1669–1675. Huggins, J. W., Kim, G. R., Brand, O. M., and McKee, K. T., Jr. (1986). Ribavirin therapy for Hantaan virus infection in suckling mice. J. Infect. Dis. 153, 489–497. Huggins, J. W., Hsiang, C. M., Cosgriff, T. M., Guang, M. Y., Smith, J. I., Wu, Z. O., LeDuc, J. W., Zheng, Z. M., Meegan, J. M., Wang, Q. N., et al. (1991). Prospective, double-blind, concurrent, placebo-controlled clinical trial of intravenous ribavirin therapy of hemorrhagic fever with renal syndrome. J. Infect. Dis. 164, 1119–1127. Huiskonen, J. T., Overby, A. K., Weber, F., and Grunewald, K. (2009). Electron cryo-microscopy and single-particle averaging of Rift Valley fever virus: Evidence for GN-GC glycoprotein heterodimers. J. Virol. 83, 3762–3769. Huiskonen, J. T., Hepojoki, J., Laurinmaki, P., Vaheri, A., Lankinen, H., Butcher, S. J., and Grunewald, K. (2010). Electron cryotomography of Tula hantavirus suggests a unique assembly paradigm for enveloped viruses. J. Virol. 84, 4889–4897. Hutchinson, K. L., Peters, C. J., and Nichol, S. T. (1996). Sin Nombre virus mRNA synthesis. Virology 224, 139–149. Hutchinson, K. L., Rollin, P. E., Shieh, W. J., Zaki, S., Greer, P. W., and Peters, C. J. (2000). Transmission of Black Creek Canal virus between cotton rats. J. Med. Virol. 60, 70–76. Ikegami, T., Peters, C. J., and Makino, S. (2005). Rift valley fever virus nonstructural protein NSs promotes viral RNA replication and transcription in a minigenome system. J. Virol. 79, 5606–5615.


67

Isegawa, Y., Tanishita, O., Ueda, S., and Yamanishi, K. (1994). Association of serine in position 1124 of Hantaan virus glycoprotein with virulence in mice. J. Gen. Virol. 75 (Pt. 11), 3273–3278. Jablonski, S. A., and Morrow, C. D. (1995). Mutation of the aspartic acid residues of the GDD sequence motif of poliovirus RNA-dependent RNA polymerase results in enzymes with altered metal ion requirements for activity. J. Virol. 69, 1532–1539. Jackson, A. P., and Charleston, M. A. (2004). A cophylogenetic perspective of RNA-virus evolution. Mol. Biol. Evol. 21, 45–57. Jakab, F., Horvath, G., Ferenczi, E., Sebok, J., Varecza, Z., and Szucs, G. (2007). Detection of Dobrava hantaviruses in Apodemus agrarius mice in the Transdanubian region of Hungary. Virus Res. 128, 149–152. Jenison, S., Yamada, T., Morris, C., Anderson, B., Torrez-Martinez, N., Keller, N., and Hjelle, B. (1994). Characterization of human antibody responses to four corners hantavirus infections among patients with hantavirus pulmonary syndrome. J. Virol. 68, 3000–3006. Jiang, W., Hou, Y., and Inouye, M. (1997). CspA, the major cold-shock protein of Escherichia coli, is an RNA chaperone. J. Biol. Chem. 272, 196–202. Jin, H., and Elliott, R. M. (1993). Characterization of Bunyamwera virus S RNA that is transcribed and replicated by the L protein expressed from recombinant vaccinia virus. J. Virol. 67, 1396–1404. Jin, M., Park, J., Lee, S., Park, B., Shin, J., Song, K. J., Ahn, T. I., Hwang, S. Y., Ahn, B. Y., and Ahn, K. (2002). Hantaan virus enters cells by clathrin-dependent receptor-mediated endocytosis. Virology 294, 60–69. Johnson, A. M., Bowen, M. D., Ksiazek, T. G., Williams, R. J., Bryan, R. T., Mills, J. N., Peters, C. J., and Nichol, S. T. (1997). Laguna Negra virus associated with HPS in western Paraguay and Bolivia. Virology 238, 115–127. Jones, K. E., Patel, N. G., Levy, M. A., Storeygard, A., Balk, D., Gittleman, J. L., and Daszak, P. (2008). Global trends in emerging infectious diseases. Nature 451, 990–993. Jonsson, C. B., Gallegos, J., Ferro, P., Severson, W., Xu, X., and Schmaljohn, C. S. (2001). Purification and characterization of the Sin Nombre virus nucleocapsid protein expressed in Escherichia coli. Protein Expr. Purif. 23, 134–141. Jonsson, C. B., Figueiredo, L. T., and Vapalahti, O. (2010). A global perspective on hantavirus ecology, epidemiology, and disease. Clin. Microbiol. Rev. 23, 412–441. Joyce, C. M., and Steitz, T. A. (1995). Polymerase structures and function: Variations on a theme? J. Bacteriol. 177, 6321–6329. Kallio, E. R., Poikonen, A., Vaheri, A., Vapalahti, O., Henttonen, H., Koskela, E., and Mappes, T. (2006). Maternal antibodies postpone hantavirus infection and enhance individual breeding success. Proc. Biol. Sci. 273, 2771–2776. Kamer, G., and Argos, P. (1984). Primary structural comparison of RNA-dependent polymerases from plant, animal and bacterial viruses. Nucleic Acids Res. 12, 7269–7282. Kanerva, M., Mustonen, J., and Vaheri, A. (1998). Pathogenesis of puumala and other hantavirus infections. Rev. Med. Virol. 8, 67–86. Kang, H. J., Bennett, S. N., Dizney, L., Sumibcay, L., Arai, S., Ruedas, L. A., Song, J. W., and Yanagihara, R. (2009a). Host switch during evolution of a genetically distinct hantavirus in the American shrew mole (Neurotrichus gibbsii). Virology 388, 8–14. Kang, H. J., Bennett, S. N., Sumibcay, L., Arai, S., Hope, A. G., Mocz, G., Song, J. W., Cook, J. A., and Yanagihara, R. (2009b). Evolutionary insights from a genetically divergent hantavirus harbored by the European common mole (Talpa europaea). PLoS ONE 4, e6149. Kariwa, H., Fujiki, M., Yoshimatsu, K., Arikawa, J., Takashima, I., and Hashimoto, N. (1998). Urine-associated horizontal transmission of Seoul virus among rats. Arch. Virol. 143, 365–374.

68


Kaukinen, P., Koistinen, V., Vapalahti, O., Vaheri, A., and Plyusnin, A. (2001). Interaction between molecules of hantavirus nucleocapsid protein. J. Gen. Virol. 82, 1845–1853. Kaukinen, P., Vaheri, A., and Plyusnin, A. (2003). Mapping of the regions involved in homotypic interactions of Tula hantavirus N protein. J. Virol. 77, 10910–10916. Kaukinen, P., Kumar, V., Tulimaki, K., Engelhardt, P., Vaheri, A., and Plyusnin, A. (2004). Oligomerization of Hantavirus N protein: C-terminal alpha-helices interact to form a shared hydrophobic space. J. Virol. 78, 13669–13677. Khaiboullina, S. F., and St Jeor, S. C. (2002). Hantavirus immunology. Viral Immunol. 15, 609–625. Kilpatrick, E. D., Terajima, M., Koster, F. T., Catalina, M. D., Cruz, J., and Ennis, F. A. (2004). Role of specific CD8þ T cells in the severity of a fulminant zoonotic viral hemorrhagic fever, hantavirus pulmonary syndrome. J. Immunol. 172, 3297–3304. Kim, Y. S., Ahn, C., Han, J. S., Kim, S., Lee, J. S., and Lee, P. W. (1995). Hemorrhagic fever with renal syndrome caused by the Seoul virus. Nephron 71, 419–427. Klempa, B., Stanko, M., Labuda, M., Ulrich, R., Meisel, H., and Kruger, D. H. (2005). Central European Dobrava Hantavirus isolate from a striped field mouse (Apodemus agrarius). J. Clin. Microbiol. 43, 2756–2763. Klempa, B., Tkachenko, E. A., Dzagurova, T. K., Yunicheva, Y. V., Morozov, V. G., Okulova, N. M., Slyusareva, G. P., Smirnov, A., and Kruger, D. H. (2008). Hemorrhagic fever with renal syndrome caused by 2 lineages of Dobrava hantavirus, Russia. Emerg. Infect. Dis. 14, 617–625. Klingstrom, J., Falk, K. I., and Lundkvist, A. (2005). Delayed viremia and antibody responses in Puumala hantavirus challenged passively immunized cynomolgus macaques. Arch. Virol. 150, 79–92. Klingstrom, J., Hardestam, J., and Lundkvist, A. (2006a). Dobrava, but not Saaremaa, hantavirus is lethal and induces nitric oxide production in suckling mice. Microbes Infect. 8, 728–737. Klingstrom, J., Hardestam, J., Stoltz, M., Zuber, B., Lundkvist, A., Linder, S., and Ahlm, C. (2006b). Loss of cell membrane integrity in puumala hantavirus-infected patients correlates with levels of epithelial cell apoptosis and perforin. J. Virol. 80, 8279–8282. Kohl, A., Hart, T. J., Noonan, C., Royall, E., Roberts, L. O., and Elliott, R. M. (2004). A bunyamwera virus minireplicon system in mosquito cells. J. Virol. 78, 5679–5685. Kohlstaedt, L. A., Wang, J., Friedman, J. M., Rice, P. A., and Steitz, T. A. (1992). Crystal structure at 3.5 A resolution of HIV-1 reverse transcriptase complexed with an inhibitor. Science 256, 1783–1790. Kramski, M., Achazi, K., Klempa, B., and Kruger, D. H. (2009). Nephropathia epidemica with a 6-week incubation period after occupational exposure to Puumala hantavirus. J. Clin. Virol. 44, 99–101. Krug, R. M. (1981). Priming of influenza viral RNA transcription by capped heterologous RNAs. Curr. Top. Microbiol. Immunol. 93, 125–149. Krug, R. M., Broni, B. A., and Bouloy, M. (1979). Are the 5’ ends of influenza viral mRNAs synthesized in vivo donated by host mRNAs? Cell 18, 329–334. Kuismanen, E., Hedman, K., Saraste, J., and Pettersson, R. F. (1982). Uukuniemi virus maturation: Accumulation of virus particles and viral antigens in the Golgi complex. Mol. Cell. Biol. 2, 1444–1458. Kukkonen, S. K., Vaheri, A., and Plyusnin, A. (1998). Completion of the Tula hantavirus genome sequence: Properties of the L segment and heterogeneity found in the 3’ termini of S and L genome RNAs. J. Gen. Virol. 79(Pt. 11), 2615–2622. Kukkonen, S. K., Vaheri, A., and Plyusnin, A. (2004). Tula hantavirus L protein is a 250 kDa perinuclear membrane-associated protein. J. Gen. Virol. 85, 1181–1189.


69

Lappin, D. F., Nakitare, G. W., Palfreyman, J. W., and Elliott, R. M. (1994). Localization of Bunyamwera bunyavirus G1 glycoprotein to the Golgi requires association with G2 but not with NSm. J. Gen. Virol. 75(Pt. 12), 3441–3451. Larson, R. S., Brown, D. C., Ye, C., and Hjelle, B. (2005). Peptide antagonists that inhibit Sin Nombre virus and hantaan virus entry through the beta3-integrin receptor. J. Virol. 79, 7319–7326. Lee, H. W., French, G. R., Lee, P. W., Baek, L. J., Tsuchiya, K., and Foulke, R. S. (1981a). Observations on natural and laboratory infection of rodents with the etiologic agent of Korean hemorrhagic fever. Am. J. Trop. Med. Hyg. 30, 477–482. Lee, H. W., Lee, P. W., Baek, L. J., Song, C. K., and Seong, I. W. (1981b). Intraspecific transmission of Hantaan virus, etiologic agent of Korean hemorrhagic fever, in the rodent Apodemus agrarius. Am. J. Trop. Med. Hyg. 30, 1106–1112. Lee, B. H., Yoshimatsu, K., Maeda, A., Ochiai, K., Morimatsu, M., Araki, K., Ogino, M., Morikawa, S., and Arikawa, J. (2003). Association of the nucleocapsid protein of the Seoul and Hantaan hantaviruses with small ubiquitin-like modifier-1-related molecules. Virus Res. 98, 83–91. Lesburg, C. A., Cable, M. B., Ferrari, E., Hong, Z., Mannarino, A. F., and Weber, P. C. (1999). Crystal structure of the RNA-dependent RNA polymerase from hepatitis C virus reveals a fully encircled active site. Nat. Struct. Biol. 6, 937–943. Li, D., Schmaljohn, A. L., Anderson, K., and Schmaljohn, C. S. (1995). Complete nucleotide sequences of the M and S segments of two hantavirus isolates from California: Evidence for reassortment in nature among viruses related to hantavirus pulmonary syndrome. Virology 206, 973–983. Li, Z., Bai, X., and Bian, H. (2002). Serologic diagnosis of Hantaan virus infection based on a peptide antigen. Clin. Chem. 48, 645–647. Linderholm, M., Ahlm, C., Settergren, B., Waage, A., and Tarnvik, A. (1996). Elevated plasma levels of tumor necrosis factor (TNF)-alpha, soluble TNF receptors, interleukin (IL)-6, and IL-10 in patients with hemorrhagic fever with renal syndrome. J. Infect. Dis. 173, 38–43. Lober, C., Anheier, B., Lindow, S., Klenk, H. D., and Feldmann, H. (2001). The Hantaan virus glycoprotein precursor is cleaved at the conserved pentapeptide WAASA. Virology 289, 224–229. Lopez, N., Muller, R., Prehaud, C., and Bouloy, M. (1995). The L protein of Rift Valley fever virus can rescue viral ribonucleoproteins and transcribe synthetic genome-like RNA molecules. J. Virol. 69, 3972–3979. Lyubsky, S., Gavrilovskaya, I., Luft, B., and Mackow, E. (1996). Histopathology of Peromyscus leucopus naturally infected with pathogenic NY-1 hantaviruses: Pathologic markers of HPS viral infection in mice. Lab. Invest. 74, 627–633. Mackow, E. R., and Gavrilovskaya, I. N. (2001). Cellular receptors and hantavirus pathogenesis. Curr. Top. Microbiol. Immunol. 256, 91–115. Markotic, A., Hensley, L., Daddario, K., Spik, K., Anderson, K., and Schmaljohn, C. (2007). Pathogenic hantaviruses elicit different immunoreactions in THP-1 cells and primary monocytes and induce differentiation of human monocytes to dendritic-like cells. Coll. Antropol. 31, 1159–1167. Martin, M. L., Lindsey-Regnery, H., Sasso, D. R., McCormick, J. B., and Palmer, E. (1985). Distinction between Bunyaviridae genera by surface structure and comparison with Hantaan virus using negative stain electron microscopy. Arch. Virol. 86, 17–28. Matsuoka, Y., Chen, S. Y., and Compans, R. W. (1994). A signal for Golgi retention in the bunyavirus G1 glycoprotein. J. Biol. Chem. 269, 22565–22573. McCaughey, C., and Hart, C. A. (2000). Hantaviruses. J. Med. Microbiol. 49, 587–599. McClain, D. J., Summers, P. L., Harrison, S. A., Schmaljohn, A. L., and Schmaljohn, C. S. (2000). Clinical evaluation of a vaccinia-vectored Hantaan virus vaccine. J. Med. Virol. 60, 77–85.

70


Medina, R. A., Mirowsky-Garcia, K., Hutt, J., and Hjelle, B. (2007). Ribavirin, human convalescent plasma and anti-beta3 integrin antibody inhibit infection by Sin Nombre virus in the deer mouse model. J. Gen. Virol. 88, 493–505. Mertz, G. J., Miedzinski, L., Goade, D., Pavia, A. T., Hjelle, B., Hansbarger, C. O., Levy, H., Koster, F. T., Baum, K., Lindemulder, A., Wang, W., Riser, L., et al. (2004). Placebocontrolled, double-blind trial of intravenous ribavirin for the treatment of hantavirus cardiopulmonary syndrome in North America. Clin. Infect. Dis. 39, 1307–1313. Meyer, B. J., and Schmaljohn, C. (2000). Accumulation of terminally deleted RNAs may play a role in Seoul virus persistence. J. Virol. 74, 1321–1331. Mills, K. H. (2004). Regulatory T cells: Friend or foe in immunity to infection? Nat. Rev. Immunol. 4, 841–855. Mir, M. A. (2010). Hantaviruses. Clin. Lab. Med. 30, 67–91. Mir, M. A., and Panganiban, A. T. (2004). Trimeric hantavirus nucleocapsid protein binds specifically to the viral RNA panhandle. J. Virol. 78, 8281–8288. Mir, M. A., and Panganiban, A. T. (2005). The hantavirus nucleocapsid protein recognizes specific features of the viral RNA panhandle and is altered in conformation upon RNA binding. J. Virol. 79, 1824–1835. Mir, M. A., and Panganiban, A. T. (2010). The triplet repeats of the Sin Nombre hantavirus 5’ UTR are sufficient in cis for nucleocapsid-mediated translation initiation. J. Virol. 84, 8937–8944. Mir, M. A., Brown, B., Hjelle, B., Duran, W. A., and Panganiban, A. T. (2006). Hantavirus N protein exhibits genus-specific recognition of the viral RNA panhandle. J. Virol. 80, 11283–11292. Mir, M. A., and Panganiban, A. T. (2008). A protein that replaces the entire cellular eIF4F complex. EMBO J. 27, 3129–3139. Mir, M. A., Duran, W. A., Hjelle, B. L., Ye, C., and Panganiban, A. T. (2008). Storage of cellular 5’ mRNA caps in P bodies for viral cap-snatching. Proc. Natl. Acad. Sci. USA 105, 19294–19299. Mir, M. A., Sheema, S., Haseeb, A., and Haque, A. (2010). Hantavirus nucleocapsid protein has distinct m7G cap- and RNA-binding sites. J. Biol. Chem. 285, 11357–11368. Moreli, M. L., Sousa, R. L., and Figueiredo, L. T. (2004). Detection of Brazilian hantavirus by reverse transcription polymerase chain reaction amplification of N gene in patients with hantavirus cardiopulmonary syndrome. Mem. Inst. Oswaldo Cruz 99, 633–638. Morens, D. M., Folkers, G. K., and Fauci, A. S. (2004). The challenge of emerging and re-emerging infectious diseases. Nature 430, 242–249. Mou, D. L., Wang, Y. P., Huang, C. X., Li, G. Y., Pan, L., Yang, W. S., and Bai, X. F. (2006). Cellular entry of Hantaan virus A9 strain: Specific interactions with beta3 integrins and a novel 70 kDa protein. Biochem. Biophys. Res. Commun. 339, 611–617. Muller, R., Poch, O., Delarue, M., Bishop, D. H., and Bouloy, M. (1994). Rift Valley fever virus L segment: Correction of the sequence and possible functional role of newly identified regions conserved in RNA-dependent polymerases. J. Gen. Virol. 75(Pt. 6), 1345–1352. Muranyi, W., Bahr, U., Zeier, M., and van der Woude, F. J. (2005). Hantavirus infection. J. Am. Soc. Nephrol. 16, 3669–3679. Nemirov, K., Vapalahti, O., Papa, A., Plyusnina, A., Lundkvist, A., Antoniadis, A., Vaheri, A., and Plyusnin, A. (2003). Genetic characterization of new Dobrava hantavirus isolate from Greece. J. Med. Virol. 69, 408–416. Netski, D., Thran, B. H., and St. Jeor, S. C. (1999). Sin Nombre virus pathogenesis in Peromyscus maniculatus. J. Virol. 73, 585–591. Ng, K. K., Arnold, J. J., and Cameron, C. E. (2008). Structure–function relationships among RNA-dependent RNA polymerases. Curr. Top. Microbiol. Immunol. 320, 137–156.


71

Nichol, S. T., Spiropoulou, C. F., Morzunov, S., Rollin, P. E., Ksiazek, T. G., Feldmann, H., Sanchez, A., Childs, J., Zaki, S., and Peters, C. J. (1993). Genetic identification of a hantavirus associated with an outbreak of acute respiratory illness. Science 262, 914–917. Nolte, K. B., Feddersen, R. M., Foucar, K., Zaki, S. R., Koster, F. T., Madar, D., Merlin, T. L., McFeeley, P. J., Umland, E. T., and Zumwalt, R. E. (1995). Hantavirus pulmonary syndrome in the United States: A pathological description of a disease caused by a new agent. Hum. Pathol. 26, 110–120. O’Reilly, E. K., and Kao, C. C. (1998). Analysis of RNA-dependent RNA polymerase structure and function as guided by known polymerase structures and computer predictions of secondary structure. Virology 252, 287–303. Obijeski, J. F., and Murphy, F. A. (1977). Bunyaviridae: Recent biochemical developments. J. Gen. Virol. 37, 1–14. Obijeski, J. F., Bishop, D. H., Palmer, E. L., and Murphy, F. A. (1976). Segmented genome and nucleocapsid of La Crosse virus. J. Virol. 20, 664–675. Obijeski, J. F., McCauley, J., and Skehel, J. J. (1980). Nucleotide sequences at the terminal of La Crosse virus RNAs. Nucleic Acids Res. 8, 2431–2438. Osborne, J. C., and Elliott, R. M. (2000). RNA binding properties of bunyamwera virus nucleocapsid protein and selective binding to an element in the 5’ terminus of the negative-sense S segment. J. Virol. 74, 9946–9952. Overby, A. K., Pettersson, R. F., and Neve, E. P. (2007). The glycoprotein cytoplasmic tail of Uukuniemi virus (Bunyaviridae) interacts with ribonucleoproteins and is critical for genome packaging. J. Virol. 81, 3198–3205. Overby, A. K., Pettersson, R. F., Grunewald, K., and Huiskonen, J. T. (2008). Insights into bunyavirus architecture from electron cryotomography of Uukuniemi virus. Proc. Natl. Acad. Sci. USA 105, 2375–2379. Padula, P. J., Edelstein, A., Miguel, S. D., Lopez, N. M., Rossi, C. M., and Rabinovich, R. D. (1998). Hantavirus pulmonary syndrome outbreak in Argentina: Molecular evidence for person-to-person transmission of Andes virus. Virology 241, 323–330. Padula, P. J., Sanchez, A. J., Edelstein, A., and Nichol, S. T. (2002). Complete nucleotide sequence of the M RNA segment of Andes virus and analysis of the variability of the termini of the virus S, M and L RNA segments. J. Gen. Virol. 83, 2117–2122. Panganiban, A. T., and Mir, M. A. (2009). Bunyavirus N: eIF4F surrogate and cap-guardian. Cell Cycle 8, 1332–1337. Patterson, J. L., Holloway, B., and Kolakofsky, D. (1984). La Crosse virions contain a primerstimulated RNA polymerase and a methylated cap-dependent endonuclease. J. Virol. 52, 215–222. Pensiero, M. N., and Hay, J. (1992). The Hantaan virus M-segment glycoproteins G1 and G2 can be expressed independently. J. Virol. 66, 1907–1914. Peters, C. J., Simpson, G. L., and Levy, H. (1999). Spectrum of hantavirus infection: Hemorrhagic fever with renal syndrome and hantavirus pulmonary syndrome. Annu. Rev. Med. 50, 531–545. Pettersson, R. F., and von Bonsdorff, C. H. (1975). Ribonucleoproteins of Uukuniemi virus are circular. J. Virol. 15, 386–392. Pinschewer, D. D., Perez, M., and de la Torre, J. C. (2003). Role of the virus nucleoprotein in the regulation of lymphocytic choriomeningitis virus transcription and RNA replication. J. Virol. 77, 3882–3887. Plotch, S. J., Bouloy, M., Ulmanen, I., and Krug, R. M. (1981). A unique cap(m7G pppXm)dependent influenza virion endonuclease cleaves capped RNAs to generate the primers that initiate viral RNA transcription. Cell 23, 847–858. Plyusnin, A. (2002). Genetics of hantaviruses: Implications to taxonomy. Arch. Virol. 147, 665–682.

72


Plyusnin, A., and Morzunov, S. P. (2001). Virus evolution and genetic diversity of hantaviruses and their rodent hosts. Curr. Top. Microbiol. Immunol. 256, 47–75. Plyusnin, A., Horling, J., Kanerva, M., Mustonen, J., Cheng, Y., Partanen, J., Vapalahti, O., Kukkonen, S. K., Niemimaa, J., Henttonen, H., Niklasson, B., Lundkvist, A., et al. (1997). Puumala hantavirus genome in patients with nephropathia epidemica: Correlation of PCR positivity with HLA haplotype and link to viral sequences in local rodents. J. Clin. Microbiol. 35, 1090–1096. Plyusnin, A., Kukkonen, S. K., Plyusnina, A., Vapalahti, O., and Vaheri, A. (2002). Transfection-mediated generation of functionally competent Tula hantavirus with recombinant S RNA segment. EMBO J. 21, 1497–1503. Poch, O., Sauvaget, I., Delarue, M., and Tordo, N. (1989). Identification of four conserved motifs among the RNA-dependent polymerase encoding elements. EMBO J. 8, 3867–3874. Raftery, M. J., Kraus, A. A., Ulrich, R., Kruger, D. H., and Schonrich, G. (2002). Hantavirus infection of dendritic cells. J. Virol. 76, 10724–10733. Raju, R., and Kolakofsky, D. (1987). Unusual transcripts in La Crosse virus-infected cells and the site for nucleocapsid assembly. J. Virol. 61, 667–672. Raju, R., and Kolakofsky, D. (1989). The ends of La Crosse virus genome and antigenome RNAs within nucleocapsids are base paired. J. Virol. 63, 122–128. Ramanathan, H. N., and Jonsson, C. B. (2008). New and Old World hantaviruses differentially utilize host cytoskeletal components during their life cycles. Virology 374, 138–150. Ramanathan, H. N., Chung, D. H., Plane, S. J., Sztul, E., Chu, Y. K., Guttieri, M. C., McDowell, M., Ali, G., and Jonsson, C. B. (2007). Dynein-dependent transport of the hantaan virus nucleocapsid protein to the endoplasmic reticulum-Golgi intermediate compartment. J. Virol. 81, 8634–8647. Ramsden, C., Melo, F. L., Figueiredo, L. M., Holmes, E. C., and Zanotto, P. M. (2008). High rates of molecular evolution in hantaviruses. Mol. Biol. Evol. 25, 1488–1492. Ravkov, E. V., and Compans, R. W. (2001). Hantavirus nucleocapsid protein is expressed as a membrane-associated protein in the perinuclear region. J. Virol. 75, 1808–1815. Ravkov, E. V., Nichol, S. T., and Compans, R. W. (1997). Polarized entry and release in epithelial cells of Black Creek Canal virus, a New World hantavirus. J. Virol. 71, 1147–1154. Ravkov, E. V., Nichol, S. T., Peters, C. J., and Compans, R. W. (1998). Role of actin microfilaments in Black Creek Canal virus morphogenesis. J. Virol. 72, 2865–2870. Rizvanov, A. A., van Geelen, A. G., Morzunov, S., Otteson, E. W., Bohlman, C., Pari, G. S., and St. Jeor, S. C. (2003). Generation of a recombinant cytomegalovirus for expression of a hantavirus glycoprotein. J. Virol. 77, 12203–12210. Rodriguez, L. L., Owens, J. H., Peters, C. J., and Nichol, S. T. (1998). Genetic reassortment among viruses causing hantavirus pulmonary syndrome. Virology 242, 99–106. Root, J. J., Black, W. C.t., Calisher, C. H., Wilson, K. R., and Beaty, B. J. (2004). Genetic relatedness of deer mice (Peromyscus maniculatus) infected with Sin Nombre virus. Vector Borne Zoonotic Dis. 4, 149–157. Rossier, C., Patterson, J., and Kolakofsky, D. (1986). La Crosse virus small genome mRNA is made in the cytoplasm. J. Virol. 58, 647–650. Rothwell, P. J., and Waksman, G. (2005). Structure and mechanism of DNA polymerases. Adv. Protein Chem. 71, 401–440. Ruo, S. L., Sanchez, A., Elliott, L. H., Brammer, L. S., McCormick, J. B., and Fisher-Hoch, S. P. (1991). Monoclonal antibodies to three strains of hantaviruses: Hantaan, R22, and Puumala. Arch. Virol. 119, 1–11. Rusnak, J. M., Byrne, W. R., Chung, K. N., Gibbs, P. H., Kim, T. T., Boudreau, E. F., Cosgriff, T., Pittman, P., Kim, K. Y., Erlichman, M. S., Rezvani, D. F., and


73

Huggins, J. W. (2009). Experience with intravenous ribavirin in the treatment of hemorrhagic fever with renal syndrome in Korea. Antiviral Res. 81, 68–76. Ruusala, A., Persson, R., Schmaljohn, C. S., and Pettersson, R. F. (1992). Coexpression of the membrane glycoproteins G1 and G2 of Hantaan virus is required for targeting to the Golgi complex. Virology 186, 53–64. Salonen, A., Ahola, T., and Kaariainen, L. (2005). Viral RNA replication in association with cellular membranes. Curr. Top. Microbiol. Immunol. 285, 139–173. Sanchez, A. B., and de la Torre, J. C. (2005). Genetic and biochemical evidence for an oligomeric structure of the functional L polymerase of the prototypic arenavirus lymphocytic choriomeningitis virus. J. Virol. 79, 7262–7268. Schmaljohn, C., and Hjelle, B. (1997). Hantaviruses: A global disease problem. Emerg. Infect. Dis. 3, 95–104. Schmaljohn, C. S., and Nichol, S. T. (2007). Bunyaviridae. In ‘‘Fields Virology’’ (D. M. Knipe and P. M. Howley, Eds.), pp. 1741–1789. Lippincott Williams & Wilkins, Philadelphia. Schmaljohn, C. S., Hasty, S. E., Dalrymple, J. M., LeDuc, J. W., Lee, H. W., von Bonsdorff, C. H., Brummer-Korvenkontio, M., Vaheri, A., Tsai, T. F., Regnery, H. L., et al. (1985). Antigenic and genetic properties of viruses linked to hemorrhagic fever with renal syndrome. Science 227, 1041–1044. Schmaljohn, C. S., Jennings, G. B., Hay, J., and Dalrymple, J. M. (1986). Coding strategy of the S genome segment of Hantaan virus. Virology 155, 633–643. Schmaljohn, C. S., Schmaljohn, A. L., and Dalrymple, J. M. (1987). Hantaan virus M RNA: Coding strategy, nucleotide sequence, and gene order. Virology 157, 31–39. Schmaljohn, C. S., Chu, Y. K., Schmaljohn, A. L., and Dalrymple, J. M. (1990). Antigenic subunits of Hantaan virus expressed by baculovirus and vaccinia virus recombinants. J. Virol. 64, 3162–3170. Schonrich, G., Rang, A., Lutteke, N., Raftery, M. J., Charbonnel, N., and Ulrich, R. G. (2008). Hantavirus-induced immunity in rodent reservoirs and humans. Immunol. Rev. 225, 163–189. Sen, N., Sen, A., and Mackow, E. R. (2007). Degrons at the C terminus of the pathogenic but not the nonpathogenic hantavirus G1 tail direct proteasomal degradation. J. Virol. 81, 4323–4330. Severson, W., Partin, L., Schmaljohn, C. S., and Jonsson, C. B. (1999). Characterization of the Hantaan nucleocapsid protein–ribonucleic acid interaction. J. Biol. Chem. 274, 33732–33739. Severson, W. E., Schmaljohn, C. S., Javadian, A., and Jonsson, C. B. (2003). Ribavirin causes error catastrophe during Hantaan virus replication. J. Virol. 77, 481–488. Severson, W., Xu, X., Kuhn, M., Senutovitch, N., Thokala, M., Ferron, F., Longhi, S., Canard, B., and Jonsson, C. B. (2005). Essential amino acids of the hantaan virus N protein in its interaction with RNA. J. Virol. 79, 10032–10039. Shi, X., and Elliott, R. M. (2002). Golgi localization of Hantaan virus glycoproteins requires coexpression of G1 and G2. Virology 300, 31–38. Shi, X., and Elliott, R. M. (2004). Analysis of N-linked glycosylation of hantaan virus glycoproteins and the role of oligosaccharide side chains in protein folding and intracellular trafficking. J. Virol. 78, 5414–5422. Shi, X., Lappin, D. F., and Elliott, R. M. (2004). Mapping the Golgi targeting and retention signal of Bunyamwera virus glycoproteins. J. Virol. 78, 10793–10802. Shi, X., Kohl, A., Li, P., and Elliott, R. M. (2007). Role of the cytoplasmic tail domains of Bunyamwera orthobunyavirus glycoproteins Gn and Gc in virus assembly and morphogenesis. J. Virol. 81, 10151–10160. Simons, J. F., and Pettersson, R. F. (1991). Host-derived 5’ ends and overlapping complementary 3’ ends of the two mRNAs transcribed from the ambisense S segment of Uukuniemi virus. J. Virol. 65, 4741–4748.

74


Simpson, S. Q., Spikes, L., Patel, S., and Faruqi, I. (2010). Hantavirus pulmonary syndrome. Infect. Dis. Clin. North Am. 24, 159–173. Sironen, T., Klingstrom, J., Vaheri, A., Andersson, L. C., Lundkvist, A., and Plyusnin, A. (2008). Pathology of Puumala hantavirus infection in macaques. PLoS ONE 3, e3035. Smallwood, S., and Moyer, S. A. (2004). The L polymerase protein of parainfluenza virus 3 forms an oligomer and can interact with the heterologous Sendai virus L, P and C proteins. Virology 318, 439–450. Smallwood, S., Cevik, B., and Moyer, S. A. (2002). Intragenic complementation and oligomerization of the L subunit of the sendai virus RNA polymerase. Virology 304, 235–245. Song, J. W., Baek, L. J., Schmaljohn, C. S., and Yanagihara, R. (2007a). Thottapalayam virus, a prototype shrewborne hantavirus. Emerg. Infect. Dis. 13, 980–985. Song, J. W., Kang, H. J., Song, K. J., Truong, T. T., Bennett, S. N., Arai, S., Truong, N. U., and Yanagihara, R. (2007b). Newfound hantavirus in Chinese mole shrew, Vietnam. Emerg. Infect. Dis. 13, 1784–1787. Song, K. J., Baek, L. J., Moon, S., Ha, S. J., Kim, S. H., Park, K. S., Klein, T. A., Sames, W., Kim, H. C., Lee, J. S., Yanagihara, R., and Song, J. W. (2007c). Muju virus, a novel hantavirus harboured by the arvicolid rodent Myodes regulus in Korea. J. Gen. Virol. 88, 3121–3129. Spiropoulou, C. F. (2001). Hantavirus maturation. In ‘‘Hantaviruses’’ (C. L. Schmaljohn and S. T. Nichol, Eds.), Springer-Verlag, Berlin, Heidelberg. Spiropoulou, C. F., Morzunov, S., Feldmann, H., Sanchez, A., Peters, C. J., and Nichol, S. T. (1994). Genome structure and variability of a virus causing hantavirus pulmonary syndrome. Virology 200, 715–723. Spiropoulou, C. F., Goldsmith, C. S., Shoemaker, T. R., Peters, C. J., and Compans, R. W. (2003). Sin Nombre virus glycoprotein trafficking. Virology 308, 48–63. Steitz, T. A., Smerdon, S. J., Jager, J., and Joyce, C. M. (1994). A unified polymerase mechanism for nonhomologous DNA and RNA polymerases. Science 266, 2022–2025. Stohwasser, R., Raab, K., Darai, G., and Bautz, E. K. (1991). Primary structure of the large (L) RNA segment of nephropathia epidemica virus strain Hallnas B1 coding for the viral RNA polymerase. Virology 183, 386–391. Sundstrom, J. B., McMullan, L. K., Spiropoulou, C. F., Hooper, W. C., Ansari, A. A., Peters, C. J., and Rollin, P. E. (2001). Hantavirus infection induces the expression of RANTES and IP-10 without causing increased permeability in human lung microvascular endothelial cells. J. Virol. 75, 6070–6085. Taruishi, M., Yoshimatsu, K., Hatsuse, R., Okumura, M., Nakamura, I., and Arikawa, J. (2008). Lack of vertical transmission of Hantaan virus from persistently infected dam to progeny in laboratory mice. Arch. Virol. 153, 1605–1609. Toh, H., Kikuno, R., Hayashida, H., Miyata, T., Kugimiya, W., Inouye, S., Yuki, S., and Saigo, K. (1985). Close structural resemblance between putative polymerase of a Drosophila transposable genetic element 17.6 and pol gene product of Moloney murine leukaemia virus. EMBO J. 4, 1267–1272. Tordo, N., Poch, O., Ermine, A., Keith, G., and Rougeon, F. (1988). Completion of the rabies virus genome sequence determination: Highly conserved domains among the L (polymerase) proteins of unsegmented negative-strand RNA viruses. Virology 165, 565–576. Ulmanen, I., Seppala, P., and Pettersson, R. F. (1981). In vitro translation of Uukuniemi virusspecific RNAs: Identification of a nonstructural protein and a precursor to the membrane glycoproteins. J. Virol. 37, 72–79. Ulrich, R., Hjelle, B., Pitra, C., and Kruger, D. H. (2002). Emerging viruses: The case ‘hantavirus’. Intervirology 45, 318–327. Van Epps, H. L., Schmaljohn, C. S., and Ennis, F. A. (1999). Human memory cytotoxic T-lymphocyte (CTL) responses to Hantaan virus infection: Identification of virus-specific and cross-reactive CD8(þ) CTL epitopes on nucleocapsid protein. J. Virol. 73, 5301–5308.


75

Vazquez, A. L., Alonso, J. M., and Parra, F. (2000). Mutation analysis of the GDD sequence motif of a calicivirus RNA-dependent RNA polymerase. J. Virol. 74, 3888–3891. Vincent, M. J., Quiroz, E., Gracia, F., Sanchez, A. J., Ksiazek, T. G., Kitsutani, P. T., Ruedas, L. A., Tinnin, D. S., Caceres, L., Garcia, A., Rollin, P. E., Mills, J. N., et al. (2000). Hantavirus pulmonary syndrome in Panama: Identification of novel hantaviruses and their likely reservoirs. Virology 277, 14–19. Walter, C. T., and Barr, J. N. (2010). Bunyamwera virus can repair both insertions and deletions during RNA replication. RNA 16, 1138–1145. Wang, H., Strandin, T., Hepojoki, J., Lankinen, H., and Vaheri, A. (2009). Degradation and aggresome formation of the Gn tail of the apathogenic Tula hantavirus. J. Gen. Virol. 90, 2995–3001. Wolf, E., Kim, P. S., and Berger, B. (1997). MultiCoil: A program for predicting two- and three-stranded coiled coils. Protein Sci. 6, 1179–1189. Worobey, M., and Holmes, E. C. (1999). Evolutionary aspects of recombination in RNA viruses. J. Gen. Virol. 80(Pt. 10), 2535–2543. Yamada, T., Hjelle, B., Lanzi, R., Morris, C., Anderson, B., and Jenison, S. (1995). Antibody responses to Four Corners hantavirus infections in the deer mouse (Peromyscus maniculatus): Identification of an immunodominant region of the viral nucleocapsid protein. J. Virol. 69, 1939–1943. Yang, X., Khosravi-Far, R., Chang, H. Y., and Baltimore, D. (1997). Daxx, a novel Fas-binding protein that activates JNK and apoptosis. Cell 89, 1067–1076. Yoneyama, M., Kikuchi, M., Natsukawa, T., Shinobu, N., Imaizumi, T., Miyagishi, M., Taira, K., Akira, S., and Fujita, T. (2004). The RNA helicase RIG-I has an essential function in double-stranded RNA-induced innate antiviral responses. Nat. Immunol. 5, 730–737. Yoshimatsu, K., Yoo, Y. C., Yoshida, R., Ishihara, C., Azuma, I., and Arikawa, J. (1993). Protective immunity of Hantaan virus nucleocapsid and envelope protein studied using baculovirus-expressed proteins. Arch. Virol. 130, 365–376. Yoshimatsu, K., Arikawa, J., Tamura, M., Yoshida, R., Lundkvist, A., Niklasson, B., Kariwa, H., and Azuma, I. (1996). Characterization of the nucleocapsid protein of Hantaan virus strain 76–118 using monoclonal antibodies. J. Gen. Virol. 77(Pt. 4), 695–704. Young, J. C., Hansen, G. R., Graves, T. K., Deasy, M. P., Humphreys, J. G., Fritz, C. L., Gorham, K. L., Khan, A. S., Ksiazek, T. G., Metzger, K. B., and Peters, C. J. (2000). The incubation period of hantavirus pulmonary syndrome. Am. J. Trop. Med. Hyg. 62, 714–717. Zaki, S. R., Greer, P. W., Coffield, L. M., Goldsmith, C. S., Nolte, K. B., Foucar, K., Feddersen, R. M., Zumwalt, R. E., Miller, G. L., Khan, A. S., et al. (1995). Hantavirus pulmonary syndrome. Pathogenesis of an emerging infectious disease. Am. J. Pathol. 146, 552–579. Zamoto-Niikura, A., Terasaki, K., Ikegami, T., Peters, C. J., and Makino, S. (2009). Rift valley fever virus L protein forms a biologically active oligomer. J. Virol. 83, 12779–12789. Zeier, M., Handermann, M., Bahr, U., Rensch, B., Muller, S., Kehm, R., Muranyi, W., and Darai, G. (2005). New ecological aspects of hantavirus infection: A change of a paradigm and a challenge of prevention—A review. Virus Genes 30, 157–180.

CHAPTER

3 Antigenic Variation and the Genetics and Epigenetics of the PfEMP1 Erythrocyte Surface Antigens in Plasmodium falciparum Malaria David E. Arnot*,†,‡,1 and Anja T. R. Jensen*,†

Contents

I. Introduction: Malaria Immunity, Vaccines, and Antigenic Variation II. The Interaction Between Antigenic Variation and Malaria Pathogenesis III. P. falciparum Genetics and Genomics IV. var Genes and the Structure of PfEMP1 Proteins V. var Gene Transcription and PfEMP1 Antigen Expression VI. Is MEE an Absolute Rule for var Genes? VII. Epigenetic Antigenic Variation VIII. Conclusions Acknowledgments References

78 80 82 85 86 87 89 90 91 91

* Centre for Medical Parasitology, Department of International Health, Immunology and Microbiology,

{

{

1

Faculty of Health Sciences, University of Copenhagen, CSS Oester Farimagsgade 5, Copenhagen K, Denmark Department of Infectious Diseases, Copenhagen University Hospital (Rigshospitalet), CSS Oester Farimagsgade 5, Copenhagen K, Denmark Institute of Immunology and Infection Research, School of Biology, University of Edinburgh, Edinburgh, Scotland, United Kingdom Corresponding author: Centre for Medical Parasitology, Department of International Health, Immunology and Microbiology, Faculty of Health Sciences, University of Copenhagen, CSS Oester Farimagsgade 5, Copenhagen K, Denmark e-mail address: [email protected]

Advances in Applied Microbiology, Volume 74 ISSN 0065-2164, DOI: 10.1016/B978-0-12-387022-3.00007-0

#


77

78

Abstract

David E. Arnot and Anja T. R. Jensen

How immunity to malaria develops remains one of the great unresolved issues in bio-medicine and resolution of its various paradoxes is likely to be the key to developing effective malaria vaccines. The basic epidemiological observations are; under conditions of intense natural transmission, humans do become immune to P. falciparum malaria, but this is a slow process requiring multiple disease episodes which many, particularly young children, do not survive. Adult survivors are immune to the symptoms of malaria, and unless pregnant, can control the growth of most or all new inoculations. Sterile immunity is not achieved and chronic parasitization of apparently healthy adults is the norm. In this article, we analyse the best understood malaria ‘‘antigenic variation’’ system, that based on Plasmodium falciparum’s PfEMP1-type cytoadhesion antigens, and critically review recent literature on the function and control of this multi-gene family of parasite variable surface antigens.

I. INTRODUCTION: MALARIA IMMUNITY, VACCINES, AND ANTIGENIC VARIATION Malaria remains a severe global public health problem which is unlikely to disappear without the introduction of a highly effective vaccine (Aguas et al., 2008). Many, but not all, malariologists consider this a realistic, albeit long-term, goal because repeated exposure to the bite of infected mosquitoes confers substantial immunity to the disease (Aponte et al., 2007; Doolan et al., 2009). However, despite the accumulation of a great deal of data on immune responses to malaria, an improved understanding of the epidemiology of malaria and development of a partially protective vaccine based on the circumsporozoite antigen (Cohen et al., 2010), it remains uncertain if highly effective vaccination can be achieved. Whether other parasite antigens can improve on the protection seen with current formulations is a subject of much current research. Explicitly or implicitly, malariological thinking is strongly marked by inclination toward one (Cohen et al., 1961), or the other (Brown and Brown, 1965), of two long-established schools of thought. What might be called the ‘‘antigenic variation’’ school attributes the slow acquisition of natural immunity to the need to accumulate immunological memory of many immunodominant variable surface antigens (VSA) expressed by the intraerythrocytic (IE) stages of the microorganism (Bull et al., 1998; Hviid, 2005; Fig. 3.1). This school has tended to view malaria vaccination as a formidably difficult problem due to the antigenic diversity of the malaria parasite, not susceptible to vaccine development by any established approach (Brown and Tanaka, 1975). The ‘‘strain-transcending immunity’’ school has been more optimistic about malaria vaccines because its adherents focus on data indicating that significant immune protection is being achieved after a few infections

Antigenic Variation and the Genetics and Epigenetics of PfEMP1 Antigens

79

B

A

5 μm

5 μm

FIGURE 3.1 Intraerythrocytic infection by Plasmodium falciparum. (A) The early intraerythrocytic stages of the protozoan malaria parasite P. falciparum, around 4 h after the invasion of a human red blood cell by the invasive merozoite stages. (B) After 48 h of intraerythrocytic growth and development of the parasites, the infected erythrocyte ruptures, releasing 8–20 daughter merozoites. The lower infected red blood cell is in the late stages of development and will shortly lyse, as has occurred in the cell above. Released merozoites will rapidly invade new erythrocytes and unchecked asexual growth causes the complex pathology, morbidity, and mortality, associated with P. falciparum malaria.

(Baird, 1994) or that immune protection can be conferred by transfused antibodies (Bouharoun-Tayoun et al., 1990; Cohen and Butcher, 1971). Both types of observation can support a view that significant immunity is acquired via pan-specific rather than variant-specific immune responses and that relatively simple malaria vaccines are conceivable—with the appropriate protective antigens (Doolan et al., 2009). Between these confines, many nuanced hypotheses have been proposed, although the experimental evaluation of conserved versus variable targets of immunity has been a slow process and this controversy is not close to being resolved. However, the genetics and genomics revolution undoubtedly offers some interesting new possibilities for faster evaluation of the predictions of competing theories of malaria immunity and uncovering the elusive immunological Achilles Heel of Plasmodium falciparum. Access to malaria parasite genome sequence data revealed that all human, simian, and rodent Plasmodium species have evolved large gene families encoding variable, but related protein antigens that vary in expression between isolates (Su et al., 1995). This was solid evidence for the existence, if not the function and importance, of malaria antigenic variation systems. The best characterized of these families are the PfEMP1 (P. falciparum erythrocyte membrane protein 1) proteins encoded by the var genes. A substantial body of evidence now indicates that these encode an immunodominant antigenic variation system of P. falciparum malaria (Marsh and Kinyanjui, 2006; Salanti et al., 2003). We will review the biology, genetics, and emerging principles underlying the control of expression of PfEMP1 genes.

80


II. THE INTERACTION BETWEEN ANTIGENIC VARIATION AND MALARIA PATHOGENESIS The seminal experiments demonstrating that recrudescent waves of malaria parasites differ in the antigens present on the Intra-erythrocytic (IE) surface were performed 50 years ago, using Plasmodium knowlesi infections of rhesus monkeys (Brown and Brown, 1965; Brown and Hills, 1974). Proof that the descendants of a single haploid clone can switch between the expression of alternative VSA during the course of an infection was provided a decade later (Barnwell et al., 1983). The discovery of antigenic variation in the P. knowlesi system in the 1960s strongly influenced the interpretation of the course of the therapeutic human malaria infections widely used in neurosyphillis treatment between the 1920s and early 1960s. These induced infections follow a course of recurrent waves of parasitaemia, although peaks decline in severity after the initial wave (Molineaux et al., 2001). The fact that repeated inoculations, with both Plasmodium vivax and P. falciparum, showed ‘‘a complete absence of ‘solid immunity,’ even when homologous strains were concerned’’ ( Jeffery, 1966) supported the view that it was antigenic variation which inhibited the acquisition of ‘‘solid immunity’’ and that changes in the antigenic composition of the parasite population led to the recurrent waves of parasitaemia (Kyes et al., 2001). Linkage between parasite erythrocyte surface proteins and serological antigenic variation was established by detection of variable, high-molecular weight P. falciparum proteins on the IE membrane (Leech et al., 1984) and the discovery of families of genes with features expected of candidate P. falciparum VSAs (Su et al., 1995). The discovery of the var genes provided a genetic basis for surface antigen switches. The realization that var-encoded PfEMP1 proteins bind to host endothelium proteins also provided a link to the pathology associated with IE sequestration (Smith et al., 1995) and further increased interest in the role of PfEMP1 as adhesion antigens. Among many interesting aspects of the role of PfEMP1 proteins as adhesins is their unusual topological distribution on the so-called ‘‘knob protuberances’’ at the IE surface (Aikawa et al., 1983). Antibodies against PfEMP1 domains can also be used to detect specific PfEMP1 antigens on the surface of live P. falciparum-infected erythrocytes (Staalsoe et al., 1999; Fig. 3.2). Antibody responses to PfEMP1 proteins are associated with the development of immunity to malaria (Bull et al., 1998; Khunrae et al., 2010), the most direct evidence for PfEMP1 being a protective antigen deriving from the study of pregnancy-associated malaria (PAM). Increasing levels of protection from the effects of PAM, such as low birth weight, correlate with antibody to the VAR2CSA PfEMP1 (Staalsoe et al., 2004) and these antibodies block the adherence of IE to placental chondroitin sulfate A (CSA), the host receptor for the PfEMP1 VAR2CSA protein (Salanti et al., 2003). That


A

B

81

C

FIGURE 3.2 Exported antigens of the parasite at the membrane of human red blood cells infected by P. falciparum. (A) Phase contrast microscopy of infected and uninfected erythrocytes. The dark granule is the optically dense, iron-containing hemozoin, a by-product of the parasite’s digestion of host hemoglobin which is sequestered in the digestive vacuole inside the growing parasite cell. (B) The same three infected and one uninfected erythrocytes, stained with an antibody detecting an exported parasite protein (KHARP) which binds to the submembranous actin–spectrin cytoskeleton of the erythrocyte. The immunofluorescently stained confocal micrograph shows the relatively abundant quantities of this antigen deposited at the periphery of the infected host cells. (C) Immunofluorescent live cell antibody staining (Bengtsson et al., 2008) of the VAR2CSA PfEMP1 (superimposed in the phase contrast image) on the outer surface of a live-stained cell. Fluorescent speckling represents PfEMP1 antigen concentrations at a surface knob structure ( Joergensen et al., 2010).

immunity to PAM is a special case of general malaria immunity, and that surviving childhood malaria involves progressive acquisition of immunity to different PfEMP1 antigens is the position of those who propose that antigenic variation plays a dominant role in malaria pathogenesis and acquired immunity (Chen, 2007; Hviid and Staalsoe, 2004; Kyes et al., 2001). PFEMP1 proteins enable IE to sequester to the host’s vascular endothelium (Smith et al., 1995) and certain binding motifs of PfEMP1 proteins also occur in otherwise unrelated proteins on P. vivax and P. knowlesi merozoites which interact with the red blood cell Duffy antigens (Chitnis and Miller, 1994). PfEMP1 was also quickly recognized to be the candidate VSA required for the role of the serum agglutination antigen discovered in serological studies in malaria endemic areas. African children’s plasma antibodies had a variable capacity to agglutinate diverse samples of parasitized erythrocytes but the capacity of children’s sera to recognize IE increased with the age and malaria infection experience of the child (Marsh and Howard, 1986). Large-scale serological studies in cohorts of African children confirmed this age and transmission intensity-related acquisition of VSAspecific antibodies and its strong correlation with the acquisition of clinical immunity (Bull et al., 1998, 1999, 2000; Nielsen et al., 2002). The finding that an IE’s capacity to be agglutinated by diverse plasma samples was associated with severe disease and young patient age was explained in terms of immune responses being first directed against ‘‘early-encountered’’ VSA, presumably those best adapted to growth in

82


that particular host (Bull et al., 2005). This implies that particular PfEMP1 adhesion receptors may be associated with the severe malaria episodes seen in young African children ( Jensen et al., 2004). That P. falciparum infections in individual malaria cases switch VSA expression was shown in the low-intensity transmission setting of Eastern Sudan (Staalsoe et al., 2002) and switching in PfEMP1 expression in experimental human infections has also been detected (Lavstsen et al., 2005; Peters et al., 2002).

III. P. FALCIPARUM GENETICS AND GENOMICS P. falciparum is a eukaryotic protozoan, with nucleus and intracellular organelles, and an obligate parasite of insects and humans. For most of its life cycle, it has a haploid complement of 14 chromosomes, in a single nucleus (Fig. 3.3). Following fusion of haploid ‘‘male’’ and 748

7G8

3D7

(752)

HB3

Mb –3.13 –2.70 –2.35

–1.81 –1.66 –1.37 –1.05

FIGURE 3.3 Pulsed field gel electrophoretic separation of the chromosomes of five P. falciparum clonal isolates. Aside from the small amounts of genetic information in the mitochondrion (6 kb) and the apicoplast (35 kb), the 23 Mb of P. falciparum genomic DNA are contained in the 14 chromosomes observed to be present in all isolates. Even with the limited resolution possible when separating such large (700–3500 kb) DNA molecules, the marked size differences between homologous chromosomes in different clonal isolates are apparent. PfEMP1 genes are present at the telomeres of almost all of these chromosomes, as well as in internal clusters.


83

‘‘female’’ gametes taken up in a blood meal from an infected host, there is a brief diploid phase in the mosquito midgut, followed by meiosis and a return to asexual multiplication of haploid forms in an oocyst formed in the mosquito stomach wall. Genome replication occurs repeatedly during sporogony in the mosquito midgut oocysts (producing 104 sporozoites/oocyst), during hepatocytic schizogony in the host liver (producing 104 hepatocytic merozoites), and during the 8- to 20-fold genome amplification occurring during intraerythrocytic replication (Arnot et al., 2011). Chromosomes present in the short-lived diploid zygote represent lineages that have completed 30–40 rounds of mitotic replication between meioses. Mitotic recombination between sister chromatids will not result in new allelic combinations unless mutations occur during growth. The rate of P. falciparum mitotic recombination has not been estimated, but in systems where mitotic recombination can be measured, it is thousands of times less frequent (104–105) than meiotic recombination (Barbera and Petes, 2006). However, relatively frequent accretions and deletions of telomeric DNA sequences occur during in vitro culture (Scherf and Mattei, 1992). Telomeric rearrangement of var genes during asexual growth (Duffy et al., 2009), as well as after meiosis (Freitas-Junior et al., 2000) are also documented. The relative contribution of mitotic and meiotic events to the generation of var gene diversity remains unclear, but the great majority of viable rearrangements of existing allele combinations, contributing to most of the evolution of the P. falciparum genome, are likely to occur at meiosis (Keightley and Otto, 2006; Petes and Pukkila, 1995). A limited number of laboratory genetic crosses between cloned P. falciparum lines have been carried out, which required chimpanzees to receive the sporozoite products of meiosis and develop the initial blood infection (Walliker et al., 1987; Wellems et al., 1990). The analysis of progeny, adapted to in vitro culture from the chimpanzee blood infections and cloned by limiting dilution, indicated that gametes fuse randomly and that meiotic recombination is significantly more frequent (30–50 times) than in higher eukaryotes, 1 cM in P. falciparum corresponding to 15–30 kb (Su et al., 1999). An excess of recombinant over parental genotypes was also found in the progeny of the cross, possibly indicating that during passage through the chimpanzee, or in in vitro culture, recombinant genotypes have been favoured by selection (Walliker et al., 1987). Intragenic recombination has been detected, although its origin and frequency are uncertain (Kerr et al., 1994). In nature, recombination frequencies have been shown to be roughly proportional to malaria transmission intensity (Babiker et al., 1997; Tanabe et al., 2007). P. falciparum chromosomes do not condense at mitosis, possibly because the parasite does not have detectable H1 linker histones (Miao

84


et al., 2006). Electron microscopic counting of spindle kinetochores (Prensier and Slomianny, 1986) and pulsed field gel (PFG) electrophoresis (Triglia et al., 1992) indicate that P. falciparum has 14 chromosomes, ranging from 0.6 to 3.5 Mb in size, in a total genome of 23 Mb (Fig. 3.3). P. falciparum has one of the most intensively studied genomes known (Gardner et al., 2002) and more than a dozen genomes have now been sequenced, with varying degrees of ‘‘finishing’’ of the repetitive and A þ T rich sequences of P. falciparum intergenic regions and telomeres. Progress has also been made in using genomic sequencing to clarify parasite population structure and evolutionary history (Mu et al., 2002, 2005). Combining genomic sequence information with other experimental data and genetic crossing results, some general observations on the genetics of P. falciparum can be made (Box 3.1).

BOX 3.1

General features of the genetics of P. Falciparum

1. P. falciparum replicates by mitotic division of a haploid genome in various vector and host cells. Recombination occurs during a brief diploid zygote phase after fusion of haploid gametes present in infected human blood meals, in the mosquito midgut. Reciprocal exchanges between parent genotypes, which segregate according to Mendelian principles, occur at meiosis in the zygote (Su et al., 1999; Walliker et al., 1987). 2. The position of most of the 5300 P. falciparum gene loci on 14 chromosomes is constant between isolates (Carlton, 2007) and there is considerable conservation of chromosome number and locus positions between different Plasmodium species (Carlton et al., 2008). 3. P. falciparum chromosome telomeres contain repetitive sequences and many hypervariable genes encoding polymorphic exported antigens (Templeton, 2009). The presence of variable genes and repetitive DNA at the telomeres explains the discrepancy between isolates having syntenic gene organization but variable PFG karyotypes showing homologous chromosome size differences (Fig. 3.3). 4. Unconventional elements of P. falciparum genetics include frequent ectopic recombination between nonhomologous chromosomes (Freitas-Junior et al., 2000), with nonreciprocal, gene conversionlike duplications of the donor sequence. Ectopic exchanges have been proposed to occur in telomere clusters located at the nuclear envelope (Freitas-Junior et al., 2000; Ralph et al., 2005b) rather than during chromatid pairing on the mitotic spindle.

85


IV. VAR GENES AND THE STRUCTURE OF PFEMP1 PROTEINS Almost all 28 P. falciparum telomeres have at least one var gene and additional intrachromosomal clusters of var genes are situated on chromosomes 4, 7, 8, and 12 of the 3D7 isolate (Gardner et al., 2002). Allelic, orthologous relationships between var genes are difficult to discern, with the exception of the var1csa and var2csa PfEMP1 genes, which are the most evolutionarily conserved var genes (Salanti et al., 2002; Sander et al., 2009). The paralogous relationships within gene families has been described as the hallmark of hypervariable malaria VSA, probably deriving from evolutionary expansion of gene families by gene duplication and immunity-driven divergence (Templeton, 2009). All var genes are organized into two exons separated by a 1–2 kb intron (Fig. 3.4). Exon 1 encodes the polymorphic extracellular domains, ending in a transmembrane sequence. Exon II encodes the more conserved 4–500 amino acid intracytoplasmic acidic terminal sequence (ATS). The ATS interacts with other exported proteins such as KHARP, which binds to actin and spectrin and connects PfEMP1 to the cytoskeleton. PfEMP1 gene sequences are essentially a mosaic of building blocks encoding amino acid segments combined into larger regions which loosely correspond to protein structural domains (Smith et al., 2001). This architecture is presumably a result of mutation and selection, followed by recombination between members of an evolving family undergoing gene duplications. Receptor-binding functions are combined with variation in the antigenic profile by tolerating mutations which alter antigenic properties, provided these do not compromise function. Structural analysis of the

Exon I

NTS

DBL1α

CIDR1α

DBL2β

Exon II

DBL3β

DBL4γ

DBL5δ

CIDR2β

TM

ATS

FIGURE 3.4 Schematic structure of a P. falciparum var gene and its encoded PfEMP1 protein. The N-terminal extracellular region and the short transmembrane (TM) sequence are encoded by Exon I and the conserved acidic terminal sequence are encoded by Exon II. The actual number and combinations of domains and domain subtypes vary considerably between the different, highly polymorphic, PfEMP1 molecules. NTS: N-terminal segment; DBL: Duffy binding-like; CIDR: cysteine-rich interdomain region; ATS: acidic terminal segment.

86


VAR2CSA PfEMP1 indicates that many amino acid substitutions are compatible with a conserved polypeptide backbone (Higgins, 2008).

V. VAR GENE TRANSCRIPTION AND PFEMP1 ANTIGEN EXPRESSION Intra-erythrocytic malaria parasites go through morphological changes (the ring, trophozoite, and schizont stages), and several mitotic divisions during the 48-h cycle between merozoite invasion and cell lysis and daughter merozoite release (Arnot and Gull, 1998; Arnot et al., 2011). These transitions are reflected in a stage-specific pattern of gene expression which may reflect a ‘‘just in time’’ system where translation immediately follows transcription (Bozdech et al., 2003). In synchronized in vitro cultures, PfEMP1 mRNA first appears around 3-5h post-merozoite invasion (PMI), steady-state var transcription peaks at around 18 h PMI, and declines to undetectable levels by 20–24 h PMI (Dahlba¨ck et al., 2007; Kyes et al., 2000). Export of PfEMP1 protein appears relatively slow, but surface-detectable PfEMP1 appears from 16 to 18 h PMI (Bengtsson et al., 2008; Kriek et al., 2003). Each var gene is associated with two distinct promoter regions, a 50 upstream (ups) promoter-regulating expression of the PfEMP1 mRNA and a second intronic promoter found in the relatively conserved intron (Calderwood et al., 2003). The intronic promoter drives expression of large, noncoding transcript whose role in var regulation is uncertain (Epp et al., 2009). Interestingly, subclasses of var genes show clearly conserved relationships between their ups promoter sequences and their positions on particular chromosomes (Kraemer and Smith, 2003; Lavstsen et al., 2003). Early studies of var transcription indicated that a proportion of the repertoire was transcribed in early ring stages (Smith et al., 1995). However, subsequent studies indicated this relaxed or loose pattern of polygenic transcription narrowed to an expression of a single locus by the trophozoite stages (Scherf et al., 1998). Transcription of many var genes in ring stages was also observed in individual IE using single-cell RT-PCR assays, but again, by the trophozoite stage of development, individual IE concentrated on the expression of one var gene (Chen et al., 1998). These studies detected only one PfEMP1 antigen on the surface of each infected erythrocyte and the proposal of a ‘‘mutually exclusive expression’’ (MEE) hypothesis that individual parasites activate only one gene at a time and that all paralogous var loci are in some way silenced. A variant of this hypothesis, where a dominant transcript is developmentally selected from initially polygenic transcripts, was also considered (Chen et al., 1998; Kyes et al., 2003). Observations that were difficult to accommodate in an MEE framework were that several experiments reported little or no


87

detectable tightening of relaxed transcription in trophozoites relative to the early ring stages, in both cultured populations and individual IE (Duffy et al., 2002; Noviyanti et al., 2001; Taylor et al., 2000). The publication of the genome sequence of the 3D7 clone of P. falciparum allowed primers to be designed which allowed quantitative RT-PCR to simultaneously measure transcription of all var genes in a population. This improved on the previous partial views of repertoire activity and analyzing the PfEMP1 genes transcribed in CSA adhesionselected parasites gave the striking result that only one gene, var2csa, was being transcribed at above background levels (Salanti et al., 2003). That repeated CSA adhesion selection establishes dominant var2csa expression was confirmed in several later studies (Dahlba¨ck et al., 2007; Duffy et al., 2005; Gamain et al., 2005). The MEE model was also supported by experiments which combined surveillance of the total var repertoire with transfection and gene knockouts to manipulate the context in which var promoters were placed. Disassociating a var ups promoter from its partner downstream intronic promoter was found to lead to constitutive activation of the unpaired ups promoter and simultaneous expression of two var promoters (Frank et al., 2006). However, forcing transfected P. falciparum cultures to express increasing numbers of episomal upsC-type promoters led to silencing of all the endogenous chromosomal var promoters (Dzikowski and Deitsch, 2008). When parasites were released from episome-mediated endogenous var promoter silencing, they reverted to exclusive expression of one var gene, from several different endogenous chromosomal promoters. Previously silenced var genes appeared to be activated randomly and the episome-mediated silencing episode appeared to have deleted any prior ‘‘memory’’ or epigenetic marking of formerly expressed genes. These experiments were interpreted as indicating the existence of a mechanism in which continuous transcription of a promoter established exclusive expression of that var gene, and that this state could be inherited without recombination in an epigenetic rather than genetic fashion (Dzikowski and Deitsch, 2008).

VI. IS MEE AN ABSOLUTE RULE FOR VAR GENES? While the analysis of transfected var promoters and CSA adhesion selection gave results which supported MEE models of var transcription, mRNA assays (Northern blots, nuclear run-on, and single-cell RT-PCR) nonetheless only sometimes (Chen et al., 1998; Horrocks et al., 2004; Scherf et al., 1998), but not always (Duffy et al., 2002; Noviyanti et al., 2001; Taylor et al., 2000), showed a narrowing of initially polygenic transcription during differentiation of untransfected Intra-erythrocytic P. falciparum. These anomalies were generally overlooked until two recent studies reopened this question.

88


That the expression of duplicated copies of the var2csa genes, which appear quite common (Sander et al., 2009), did not to appear to be governed by MEE was reported in a recent publication (Brolin et al., 2009). The linked question of whether both var transcripts are translated and exported to the erythrocyte surface was not analyzed. However ‘‘double expression’’ of two different PFMP1 antigens on a single IE has now been closely investigated and using P. falciparum selected using different, specific anti-PfEMP1 antisera, 3D7 sublines were recently shown to simultaneously express two different var mRNAs and to export both antigens to the surface of the same IE ( Joergensen et al., 2010; PLOS Pathogens). Immunofluorescence microscopy and flow cytometric detection of dual antibody binding was reinforced by RNA FISH detection of two different var transcripts in single individual cells. A functional rationale for the dual expression phenomenon was provided by showing that coexpressors, but not monoexpressors, were able to bind synergistically to two host receptors, CD54/ICAM and CD31/PECAM ( Joergensen et al., 2010; PLOS Pathogens). Whether this type of exception to the MEE type of var gene transcription situation is common and occurs in malaria infections, as well in antibodyselected in vitro P. falciparum cultures remains to be seen. Detecting dual expression depends on identifying individual parasites which are expressing two particular PfEMP1 antigen specificities which can be detected by antibody reagents. Given the enormous diversity of naturally expressed PfEMP1, there is a low probability of detecting dual expressor parasite clones with the small number of characterized antisera currently available. However, if the expression of more than one PfEMP1/IE, rather than the absolute imposition of MEE, is within the capacity of parasites in natural infections, this could be advantageous during the posthepatocytic establishment phase. The first generation posthepatocytic parasite genomes lack ‘‘memory’’ of any var gene transcription and the data indicate that MEE-based silencing of the repertoire requires prior transcription of a var gene (Chookajorn et al., 2007; Dzikowski and Deitsch, 2008). If expression of > 1 PfEMP1 by the first wave of posthepatic IE increases the likelihood of successful sequestration, this will promote clonal survival. ‘‘Successful’’ genes would be marked for expression in their descendent genomes, while unexpressed genes remain unmarked and silenced. From initially loose transcription, progressively more exclusive transcription might then be selected from the most effectively sequestering parasites. An optimally exclusive expression pattern, perhaps unigenic, will emerge. In real infections, considerable heterogeneity of surface PfEMP1 expression by the population is predicted, as variants wax and wane in the face of immunity (Dietz et al., 2006). Although this prediction has not been systematically investigated in natural infections, with specific typing sera, a narrowing of initially loose var gene


89

transcription has been observed in the earliest stages of blood stage infections induced in human volunteers (Wang et al., 2009).

VII. EPIGENETIC ANTIGENIC VARIATION Epigenetics, in the sense of heritable changes in phenotype that occur in the absence of mating or genomic rearrangement, has been invoked to explain malaria antigenic variation as transcriptional switching of PfEMP1 expression appears to leave the participant genes in situ and unaltered (Dzikowski and Deitsch, 2008; Scherf et al., 1998). Although many var switches do not involve genetic rearrangements, these have been observed. For example, in P. falciparum clone Dd2, switching from the expression of one chromosome 12 var gene to one of two adjacent var genes was accompanied by the deletion of the formerly expressed locus (Deitsch et al., 1999). In the IT/FCR3 P. falciparum lineage, a switch in expression from the A4 var to the R29 var was accompanied by translocation of the 50 end of the A4 var to another chromosome (Horrocks et al., 2004). In the CS2 line, a var gene on chromosome 4 was translocated into an existing var gene’s intron on chromosome 12, where the translocated gene became active (Duffy et al., 2009). Current data are thus compatible with the possibility that P. falciparum, in an inversion of the Trypanosoma bruceii situation where recombinational switching predominates (Morrison et al., 2005), combines some recombination-based VSA activation beside a predominantly in situ switching system. Epigenetic influences on var transcription include several different observations of expression-associated chromatin modifications and nuclear position effects on transcriptional activation (Box 3.2). As in the broader field of gene regulation, there are controversies about prime causes and secondary effects (Ptashne, 2007). Histone modifications are not known to be heritable and must be reestablished after each DNA synthesis phase. They are also generally associated with the passage of the complex transcription apparatus over a locus, or the absence of such effectors, and can be seen as an effect of gene activation rather than a primary actor. There are indications that soluble transcription factors/ promoter regulatory elements bind to var promoters and that their absence will silence ‘‘unbound’’ promoters (Dzikowski and Deitsch, 2008). This may indicate that some form of self-perpetuating (i.e., epigenetic) positive feedback loop ensures continuous transcription similar to the classical phage lambda repressor switch, which is activated by external, nongenetic influences (Ptashne, 2009). Alternatively, it may be more closely related to other eukaryotic systems where one or a small number of genes in a multigene family are selected, by some poorly understood process, for expression by individual cells, such as the Drosophila odor receptor genes in olfactory neurons (Vosshall and Stocker, 2007).

90


BOX 3.2 Epigenetic features of P. Falciparum var gene regulation

1. Trans-acting factors, presumably regulatory proteins, bind var gene promoters (Frank et al., 2006; Voss et al., 2005), and the titration of such factors by transfected episomal var promoters leads to a shutdown of endogenous var transcription (Chookajorn et al., 2007; Dzikowski and Deitsch, 2008). 2. Epigenetic marks, that is, the characteristic histone posttranslational modifications (PTM) associated with downregulated genes, including H3K9me3 and histone hypoacetylation have been found to be associated with inactive var genes (Duraisingh et al., 2005; FreitasJunior et al., 2005). 3. P. falciparum sirtuins, homologous to key components of eukaryotic chromatin modification complexes, have been shown to affect var gene regulation, sirtuin gene deletion tending to relax silencing (Merrick et al., 2010; Tonkin et al., 2009). 4. The positioning of transcriptionally active var genes at particular sites underneath the nuclear envelope membrane has been proposed to influence var gene transcription status (Duraisingh et al., 2005; Ralph et al., 2005b). 5. Noncoding RNA (ncRNA) molecules are transcribed by P. falciparum (Li et al., 2008) but no functioning RNAi regulatory system appears to exit (Baum et al., 2009). Whether or not ncRNA is involved in epigenetic silencing is unclear (Epp et al., 2009; Ralph et al., 2005a).

VIII. CONCLUSIONS Despite considerable progress in the analysis of antigenic variation in human P. falciparum malaria, at present we do not understand how the parasite establishes this system in the early stages of infection or how it breaks an established pattern of var gene expression to express a new PfEMP1 antigen. Both genetic and epigenetic features of the system are apparent but relative rates and causes and effects remain uncertain (Cui and Miao, 2010; Horrocks et al., 2009; Merrick and Duraisingh, 2010). With the establishment and characterization of genetically unmodified parasite isolates that show exceptions to the active promoter counting/MEE situation (Brolin et al., 2009; Joergensen et al 2010; PLOS Pathogens), it may be possible to further analyze what permits and/or limits multiple var gene expression. Although there are many difficulties in analyzing in vivo human infections, more data on var gene expression and switching during clinical P. falciparum malaria are needed.


91

ACKNOWLEDGMENTS We thank Dominique Bengtsson and Adam Sander for comments and unpublished micrographs and Thor Theander, Lars Hviid, Ali Salanti, Thomas Lavstsen, Morten Nielsen, Lars Joergensen, Elena Ronander, and Louise Joergensen at the Centre for Medical Parasitology, University of Copenhagen, for many interesting discussions. ‘‘We thank the Niels Bohr Foundation and Danmarks Grundforskningsfonds and the Howard Hughes Medical Foundation for support’’.

REFERENCES Aguas, R., White, L. J., Snow, R. W., and Gomes, M. G. (2008). Prospects for malaria eradication in sub-Saharan Africa. PLoS ONE 3, e1767. Aikawa, M., Rabbege, J. R., Udeinya, I., and Miller, L. H. (1983). Electron microscopy of knobs in Plasmodium falciparum-infected erythrocytes. J. Parasitol. 69, 435–437. Aponte, J. J., Menendez, C., Schellenberg, D., Kahigwa, E., Mshinda, H., Vountasou, P., Tanner, M., and Alonso, P. L. (2007). Age interactions in the development of naturally acquired immunity to Plasmodium falciparum and its clinical presentation. PLoS Med. 4, e242. Arnot, D. E., and Gull, K. (1998). The Plasmodium cell-cycle: Facts and questions. Ann. Trop. Med. Parasitol. 92, 361–365. Arnot, D. E., Ronander, E., and Bengtsson, D. C. (2011). The progression of the intraerythrocytic cell cycle of Plasmodium falciparum and the role of the centriolar plaques in asynchronous mitotic division during schizogony. Int. J. Parasitol. 41, 71–80. Babiker, H. A., Lines, J., Hill, W. G., and Walliker, D. (1997). Population structure of Plasmodium falciparum in villages with different malaria endemicity in east Africa. Am. J. Trop. Med. Hyg. 56, 141–147. Baird, J. K. (1994). Age and ‘strain-transcending’ immunity to Plasmodium falciparum. Parasitol. Today 10, 302. Barbera, M. A., and Petes, T. D. (2006). Selection and analysis of spontaneous reciprocal mitotic cross-overs in Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. USA 103, 12819–12824. Barnwell, J. W., Howard, R. J., Coon, H. G., and Miller, L. H. (1983). Splenic requirement for antigenic variation and expression of the variant antigen on the erythrocyte membrane in cloned Plasmodium knowlesi malaria. Infect. Immun. 40, 985–994. Baum, J., Papenfuss, A. T., Mair, G. R., Janse, C. J., Vlachou, D., Waters, A. P., Cowman, A. F., Crabb, B. S., and De Koning-Ward, T. F. (2009). Molecular genetics and comparative genomics reveal RNAi is not functional in malaria parasites. Nucleic Acids Res. 37, 3788–3798. Bengtsson, D., Sowa, K. M., Salanti, A., Jensen, A. T., Joergensen, L., Turner, L., Theander, T. G., and Arnot, D. E. (2008). A method for visualizing surface-exposed and internal PfEMP1 adhesion antigens in Plasmodium falciparum infected erythrocytes. Malar. J. 7, 101. Bouharoun-Tayoun, H., Attanath, P., Sabchareon, A., Chongsuphajaisiddhi, T., and Druilhe, P. (1990). Antibodies that protect humans against Plasmodium falciparum blood stages do not on their own inhibit parasite growth and invasion in vitro, but act in cooperation with monocytes. J. Exp. Med. 172, 1633–1641. Bozdech, Z., Llinas, M., Pulliam, B. L., Wong, E. D., Zhu, J., and Derisi, J. L. (2003). The transcriptome of the intraerythrocytic developmental cycle of Plasmodium falciparum. PLoS Biol. 1, 5. Brolin, K. J., Ribacke, U., Nilsson, S., Ankarklev, J., Moll, K., Wahlgren, M., and Chen, Q. (2009). Simultaneous transcription of duplicated var2csa gene copies in individual Plasmodium falciparum parasites. Genome Biol. 10, R117. Brown, K. N., and Brown, I. N. (1965). Immunity to malaria: Antigenic variation in chronic infections of Plasmodium knowlesi. Nature 208, 1286–1288.

92


Brown, K. N., and Hills, L. A. (1974). Antigenic variation and immunity to Plasmodium knowlesi: antibodies which induce antigenic variation and antibodies which destroy parasites. Trans. R. Soc. Trop. Med. Hyg. 68, 139–142. Brown, K. N., and Tanaka, A. (1975). Vaccination against Plasmodium knowlesi malaria. Trans. R. Soc. Trop. Med. Hyg. 69, 350–353. Bull, P. C., Lowe, B. S., Kortok, M., Molyneux, C. S., Newbold, C. I., and Marsh, K. (1998). Parasite antigens on the infected red cell are targets for naturally acquired immunity to malaria. Nat. Med. 4, 358–360. Bull, P. C., Lowe, B. S., Kortok, M., and Marsh, K. (1999). Antibody recognition of Plasmodium falciparum erythrocyte surface antigens in Kenya: Evidence for rare and prevalent variants. Infect. Immun. 67, 733–739. Bull, P. C., Kortok, M., Kai, O., Ndungu, F., Ross, A., Lowe, B. S., Newbold, C. I., and Marsh, K. (2000). Plasmodium falciparum-infected erythrocytes: Agglutination by diverse Kenyan plasma is associated with severe disease and young host age. J. Infect. Dis. 182, 252–259. Bull, P. C., Berriman, M., Kyes, S., Quail, M. A., Hall, N., Kortok, M. M., Marsh, K., and Newbold, C. I. (2005). Plasmodium falciparum variant surface antigen expression patterns during malaria. PLoS Pathog. 1, e26. Calderwood, M. S., Gannoun-Zaki, L., Wellems, T. E., and Deitsch, K. W. (2003). Plasmodium falciparum var genes are regulated by two regions with separate promoters, one upstream of the coding region and a second within the intron. J. Biol. Chem. 278, 34125–34132. Carlton, J. M. (2007). Toward a malaria haplotype map. Nat. Genet. 39, 5–6. Carlton, J. M., Escalante, A. A., Neafsey, D., and Volkman, S. K. (2008). Comparative evolutionary genomics of human malaria parasites. Trends Parasitol. 24, 545–550. Chen, Q. (2007). The naturally acquired immunity in severe malaria and its implication for a PfEMP-1 based vaccine. Microbes Infect. 9, 777–783. Chen, Q., Fernandez, V., Sundstro¨m, A., Schlichtherle, M., Datta, S., Hagblom, P., and Wahlgren, M. (1998). Developmental selection of var gene expression in Plasmodium falciparum. Nature 394, 392–395. Chitnis, C. E., and Miller, L. H. (1994). Identification of the erythrocyte binding domains of Plasmodium vivax and Plasmodium knowlesi proteins involved in erythrocyte invasion. J. Exp. Med. 180, 497–506. Chookajorn, T., Dzikowski, R., Frank, M., Li, F., Jiwani, A. Z., Hartl, D. L., and Deitsch, K. W. (2007). Epigenetic memory at malaria virulence genes. Proc. Natl. Acad. Sci. USA 104, 899–902. Cohen, S., and Butcher, G. A. (1971). Serum antibody in acquired malarial immunity. Trans. R. Soc. Trop. Med. Hyg. 65, 125–135. Cohen, S., McGregor, I. A., and Carrington, S. (1961). Gammaglobulin and acquired immunity to human malaria. Nature 192, 733–737. Cohen, J., Nussenzweig, V., Nussenzweig, R., Vekemans, J., and Leach, A. (2010). From the circumsporozoite protein to the RTS, S/AS candidate vaccine. Hum. Vaccin. 6, 90–96. Cui, L., and Miao, J. (2010). Chromatin-mediated epigenetic regulation in the malaria parasite Plasmodium falciparum. Eukaryot. Cell 9, 1138–1149. Dahlba¨ck, M., Lavstsen, T., Salanti, A., Hviid, L., Arnot, D. E., Theander, T., and Nielsen, M. A. (2007). Changes in var gene mRNA levels during erythrocytic development in two phenotypically distinct Plasmodium falciparum parasites. Malar. J. 6, 78. Deitsch, K. W., Del Pinal, A., and Wellems, T. E. (1999). Intra-cluster recombination and var transcription switches in the antigenic variation of Plasmodium falciparum. Mol. Biochem. Parasitol. 101, 107–116. Dietz, K., Raddatz, G., and Molineaux, L. (2006). Mathematical model of the first wave of Plasmodium falciparum asexual parasitemia in non-immune and vaccinated individuals. Am. J. Trop. Med. Hyg. 75, 46–55. Doolan, D. L., Dobano, C., and Baird, J. K. (2009). Acquired immunity to malaria. Clin. Microbiol. Rev. 22, 13–36.


93

Duffy, M. F., Byrne, T. J., Carret, C., Ivens, A., and Brown, G. V. (2009). Ectopic recombination of a malaria var gene during mitosis associated with an altered var switch rate. J. Mol. Biol. 389, 453–469. Duffy, M. F., Brown, G. V., Basuki, W., Krejany, E. O., Noviyanti, R., Cowman, A. F., and Reeder, J. C. (2002). Transcription of multiple var genes by individual, trophozoite-stage Plasmodium falciparum cells expressing a chondroitin sulphate A binding phenotype. Mol. Microbiol. 43, 1285–1293. Duffy, M. F., Byrne, T. J., Elliott, S. R., Wilson, D. W., Rogerson, S. J., Beeson, J. G., Noviyanti, R., and Brown, G. V. (2005). Broad analysis reveals a consistent pattern of var gene transcription in Plasmodium falciparum repeatedly selected for a defined adhesion phenotype. Mol. Microbiol. 56, 774–788. Duraisingh, M. T., Voss, T. S., Marty, A. J., Duffy, M. F., Good, R. T., Thompson, J. K., FreitasJunior, L. H., Scherf, A., Crabb, B. S., and Cowman, A. F. (2005). Heterochromatin silencing and locus repositioning linked to regulation of virulence genes in Plasmodium falciparum. Cell 121, 13–24. Dzikowski, R., and Deitsch, K. W. (2008). Active transcription is required for maintenance of epigenetic memory in the malaria parasite Plasmodium falciparum. J. Mol. Biol. 382, 288–297. Epp, C., Li, F., Howitt, C. A., Chookajorn, T., and Deitsch, K. W. (2009). Chromatin associated sense and antisense noncoding RNAs are transcribed from the var gene family of virulence genes of the malaria parasite Plasmodium falciparum. RNA 15, 116–127. Frank, M., Dzikowski, R., Constantini, D., Amulic, B., Berdougo, E., and Deitsch, K. (2006). Strict pairing of var promoters and introns is required for var gene silencing in the malaria parasite Plasmodium falciparum. J. Biol. Chem. 281, 9942–9952. Freitas-Junior, L. H., Bottius, E., Pirrit, L. A., Deitsch, K. W., Scheidig, C., Guinet, F., Nehrbass, U., Wellems, T. E., and Scherf, A. (2000). Frequent ectopic recombination of virulence factor genes in telomeric chromosome clusters of P. falciparum. Nature 407, 1018–1022. Freitas-Junior, L. H., Hernandez-Rivas, R., Ralph, S. A., Montiel-Condado, D., RuvalcabaSalazar, O. K., Rojas-Meza, A. P., Mancio-Silva, L., Leal-Silvestre, R. J., Gontijo, A. M., Shorte, S., and Scherf, A. (2005). Telomeric heterochromatin propagation and histone acetylation control mutually exclusive expression of antigenic variation genes in malaria parasites. Cell 121, 25–36. Gamain, B., Trimnell, A. R., Scheidig, C., Scherf, A., Miller, L. H., and Smith, J. D. (2005). Identification of multiple chondroitin sulfate A (CSA)-binding domains in the var2CSA gene transcribed in CSA-binding parasites. J. Infect. Dis. 191, 1010–1013. Gardner, M. J., Hall, N., Fung, E., White, O., Berriman, M., Hyman, R. W., Carlton, J. M., Pain, A., Nelson, K. E., Bowman, S., Paulsen, I. T., James, K., et al. (2002). Genome sequence of the human malaria parasite Plasmodium falciparum. Nature 419, 498–511. Higgins, M. K. (2008). The structure of a chondroitin sulfate-binding domain important in placental malaria. J. Biol. Chem. 283, 21842–21846. Horrocks, P., Kyes, S., Pinches, R., Christodoulou, Z., and Newbold, C. (2004). Transcription of subtelomerically located var gene variant in Plasmodium falciparum appears to require the truncation of an adjacent var gene. Mol. Biochem. Parasitol. 134, 193–199. Horrocks, P., Wong, E., Russell, K., and Emes, R. D. (2009). Control of gene expression in Plasmodium falciparum—Ten years on. Mol. Biochem. Parasitol. 164, 9–25. Hviid, L. (2005). Naturally acquired immunity to Plasmodium falciparum malaria in Africa. Acta Trop. 95, 270–275. Hviid, L., and Staalsoe, T. (2004). Malaria immunity in infants: A special case of a general phenomenon? Trends Parasitol. 20, 66–72. Jeffery, G. M. (1966). Epidemiological significance of repeated infections with homologous and heterologous strains and species of Plasmodium. Bull. World Health Organ. 35, 873–882. Jensen, A. T. R., Magistrado, P. A., Sharp, S., Joergensen, L., Lavstsen, T., Chiucchiuini, A., Salanti, A., Vestergaard, L. S., Lusingu, J. P., Hermsen, R., Sauerwein, R., Christensen, J.,

94


et al. (2004). Plasmodium falciparum associated with severe childhood malaria preferentially expresses PfEMP1 encoded by Group A var genes. J. Exp. Med. 199, 1179–1190. Joergensen, L. M., Salanti, A., Dobrilovic, T., Barfod, L., Hassenkam, T., Theander, T. G., Hviid, L., and Arnot, D. E. (2010). The kinetics of antibody binding to Plasmodium falciparum VAR2CSA PfEMP1 antigen and modelling of PfEMP1 antigen packing on the membrane knobs. Malar. J. 9, 100. Joergensen, L., Bengtsson, D. C., Bengtsson, A., Ronander, E., Berger, S. S., Turner, L., Dalgaard, M. B., Cham, G. K., Victor, M. E., Lavstsen, T., Theander, T. G., Arnot, D. E., et al. (2010). Surface co-expression of two different PfEMP1 antigens on single Plasmodium falciparum-infected erythrocytes facilitates binding to ICAM1 and PECAM1. PLoS Pathogens. 6(9), e1001083. Keightley, P. D., and Otto, S. P. (2006). Interference among deleterious mutations favours sex and recombination in finite populations. Nature 443, 89–92. Kerr, P. J., Ranford-Cartwright, L. C., and Walliker, D. (1994). Proof of intragenic recombination in Plasmodium falciparum. Mol. Biochem. Parasitol. 66, 241–248. Khunrae, P., Dahlback, M., Nielsen, M. A., Andersen, G., Ditlev, S. B., Resende, M., Pinto, V. V., Theander, T. G., Higgins, M. K., and Salanti, A. (2010). Full-length recombinant Plasmodium falciparum VAR2CSA binds specifically to CSPG and induces potent parasite adhesion-blocking antibodies. J. Mol. Biol. 397, 826–834. Kraemer, S. M., and Smith, J. D. (2003). Evidence for the importance of genetic structuring to the structural and functional specialization of the Plasmodium falciparum var gene family. Mol. Microbiol. 50, 1527–1538. Kriek, N., Tilley, L., Horrocks, P., Pinches, R., Elford, B. C., Ferguson, D. J. P., Lingelbach, K., and Newbold, C. I. (2003). Characterization of the pathway for transport of the cytoadherence-mediating protein, PfEMP1, to the host cell surface in malaria parasite-infected erythrocytes. Mol. Microbiol. 50, 1215–1227. Kyes, S., Pinches, R., and Newbold, C. (2000). A simple RNA analysis method shows var and rif multigene family expression patterns in Plasmodium falciparum. Mol. Biochem. Parasitol. 105, 311–315. Kyes, S., Horrocks, P., and Newbold, C. (2001). Antigenic variation at the infected red cell surface in malaria. Annu. Rev. Microbiol. 55, 673–707. Kyes, S. A., Christodoulou, Z., Raza, A., Horrocks, P., Pinches, R., Rowe, J. A., and Newbold, C. I. (2003). A well-conserved Plasmodium falciparum var gene shows an unusual stage-specific transcript pattern. Mol. Microbiol. 48, 1339–1348. Lavstsen, T., Salanti, A., Jensen, A. T. R., Arnot, D. E., and Theander, T. G. (2003). Subgrouping of Plasmodium falciparum 3D7 var genes based on sequence analysis of coding and non-coding regions. Malar. J. 2, 27. Lavstsen, T., Magistrado, P., Hermsen, C. C., Salanti, A., Jensen, A. T. R., Sauerwein, R., Hviid, L., Theander, T. G., and Staalsoe, T. (2005). Expression of Plasmodium falciparum erythrocyte membrane protein 1 in experimentally infected humans. Malar. J. 4, 21. Leech, J. H., Barnwell, J. W., Miller, L. H., and Howard, R. J. (1984). Identification of a strainspecific malarial antigen exposed on the surface of Plasmodium falciparum-infected erythrocytes. J. Exp. Med. 159, 1567–1575. Li, F., Sonbuchner, L., Kyes, S. A., Epp, C., and Deitsch, K. W. (2008). Nuclear non-coding RNAs are transcribed from the centromeres of Plasmodium falciparum and are associated with centromeric chromatin. J. Biol. Chem. 283, 5692–5698. Marsh, K., and Howard, R. J. (1986). Antigens induced on erythrocytes by P. falciparum: Expression of diverse and conserved determinants. Science 231, 150–153. Marsh, K., and Kinyanjui, S. (2006). Immune effector mechanisms in malaria. Parasite Immunol. 28, 51–60. Merrick, C. J., and Duraisingh, M. T. (2010). Epigenetics in Plasmodium: What do we really know? Eukaryot. Cell 9, 1150–1158.


95

Merrick, C. J., Dzikowski, R., Imamura, H., Chuang, J., Deitsch, K., and Duraisingh, M. T. (2010). The effect of Plasmodium falciparum Sir2a histone deacetylase on clonal and longitudinal variation in expression of the var family of virulence genes. Int. J. Parasitol. 40, 35–43. Miao, J., Fan, Q., Cui, L., Li, J., Li, J., and Cui, L. (2006). The malaria parasite Plasmodium falciparum histones: Organization, expression, and acetylation. Gene 369, 53–65. Molineaux, L., Diebner, H. H., Eichner, M., Collins, W. E., Jeffery, G. M., and Dietz, K. (2001). Plasmodium falciparum parasitaemia described by a new mathematical model. Parasitology 122, 379–391. Morrison, L. J., Majiwa, P., Read, A. F., and Barry, J. D. (2005). Probabilistic order in antigenic variation of Trypanosoma brucei. Int. J. Parasitol. 35, 961–972. Mu, J., Duan, J., Makova, K. D., Joy, D. A., Huynh, C. Q., Branch, O. H., Li, W. H., and Su, X. Z. (2002). Chromosome-wide SNPs reveal an ancient origin for Plasmodium falciparum. Nature 418, 323–324. Mu, J., Awadalla, P., Duan, J., McGee, K. M., Joy, D. A., McVean, G. A., and Su, X. Z. (2005). Recombination hotspots and population structure in Plasmodium falciparum. PLoS Biol. 3, e335. Nielsen, M. A., Staalsoe, T., Kurtzhals, J. A. L., Goka, B. Q., Dodoo, D., Alifrangis, M., Theander, T. G., Akanmori, B. D., and Hviid, L. (2002). Plasmodium falciparum variant surface antigen expression varies between isolates causing severe and non-severe malaria and is modified by acquired immunity. J. Immunol. 168, 3444–3450. Noviyanti, R., Brown, G. V., Wickham, M. E., Duffy, M. F., Cowman, A. F., and Reeder, J. C. (2001). Multiple var gene transcripts are expressed in Plasmodium falciparum infected erythrocytes selected for adhesion. Mol. Biochem. Parasitol. 114, 227–237. Peters, J., Fowler, E., Gatton, M., Chen, N., Saul, A., and Cheng, Q. (2002). High diversity and rapid changeover of expressed var genes during the acute phase of Plasmodium falciparum infections in human volunteers. Proc. Natl. Acad. Sci. USA 99, 10689–10694. Petes, T. D., and Pukkila, P. J. (1995). Meiotic sister chromatid recombination. Adv. Genet. 33, 41–62. Prensier, G., and Slomianny, C. (1986). The karyotype of Plasmodium falciparum determined by ultrastructural serial sectioning and 3D reconstruction. J. Parasitol. 72, 731–736. Ptashne, M. (2007). On the use of the word ‘epigenetic’. Curr. Biol. 17, R233–R236. Ptashne, M. (2009). Binding reactions: Epigenetic switches, signal transduction and cancer. Curr. Biol. 19, R234–R241. Ralph, S. A., Bischoff, E., Mattei, D., Sismeiro, O., Dillies, M. A., Guigon, G., Coppee, J. Y., David, P. H., and Scherf, A. (2005a). Transcriptome analysis of antigenic variation in Plasmodium falciparum—var silencing is not dependent on antisense RNA. Genome Biol. 6, R93. Ralph, S. A., Scheidig-Benatar, C., and Scherf, A. (2005b). Antigenic variation in Plasmodium falciparum is associated with movement of var loci between subnuclear locations. Proc. Natl. Acad. Sci. USA 102, 5414–5419. Salanti, A., Jensen, A. T. R., Zornig, H. D., Staalsoe, T., Joergensen, L., Nielsen, M. A., Khattab, A., Arnot, D. E., Klinkert, M. Q., Hviid, L., and Theander, T. G. (2002). A subfamily of common and highly conserved var genes expressed by CSA-adhering Plasmodium falciparum. Mol. Biochem. Parasitol. 122, 111–115. Salanti, A., Staalsoe, T., Lavstsen, T., Jensen, A. T. R., Sowa, M. P. K., Arnot, D. E., Hviid, L., and Theander, T. G. (2003). Selective upregulation of a single distinctly structured var gene in CSA-adhering Plasmodium falciparum involved in pregnancy-associated malaria. Mol. Microbiol. 49, 179–191. Sander, A. F., Salanti, A., Lavstsen, T., Nielsen, M. A., Magistrado, P., Lusingu, J., Ndam, N. T., and Arnot, D. E. (2009). Multiple var2csa-type PfEMP1 genes located at different chromosomal loci occur in many Plasmodium falciparum isolates. PLoS ONE 4, e6667. Scherf, A., and Mattei, D. (1992). Cloning and characterization of chromosome breakpoints of Plasmodium falciparum: Breakage and new telomere formation occurs frequently and randomly in subtelomeric genes. Nucleic Acids Res. 20, 1491–1496.

96


Scherf, A., Hernandez-Rivas, R., Buffet, P., Bottius, E., Benatar, C., Pouvelle, B., Gysin, J., and Lanzer, M. (1998). Antigenic variation in malaria: In situ switching, relaxed and mutually exclusive transcription of var genes during intra-erythrocytic development in Plasmodium falciparum. EMBO J. 17, 5418–5428. Smith, J. D., Chitnis, C. E., Craig, A. G., Roberts, D. J., Hudson-Taylor, D. E., Peterson, D. S., Pinches, R., Newbold, C. I., and Miller, L. H. (1995). Switches in expression of Plasmodium falciparum var genes correlate with changes in antigenic and cytoadherent phenotypes of infected erythrocytes. Cell 82, 101–110. Smith, J. D., Gamain, B., Baruch, D. I., and Kyes, S. (2001). Decoding the language of var genes and Plasmodium falciparum sequestration. Trends Parasitol. 17, 538–545. Staalsoe, T., Giha, H. A., Dodoo, D., Theander, T. G., and Hviid, L. (1999). Detection of antibodies to variant antigens on Plasmodium falciparum infected erythrocytes by flow cytometry. Cytometry 35, 329–336. Staalsoe, T., Hamad, A. A., Hviid, L., Elhassan, I. M., Arnot, D. E., and Theander, T. G. (2002). In vivo switching between variant surface antigens in human Plasmodium falciparum infection. J. Infect. Dis. 186, 719–722. Staalsoe, T., Shulman, C. E., Bulmer, J. N., Kawuondo, K., Marsh, K., and Hviid, L. (2004). Variant surface antigen-specific IgG and protection against the clinical consequences of pregnancy-associated Plasmodium falciparum malaria. Lancet 363, 283–289. Su, X., Heatwole, V. M., Wertheimer, S. P., Guinet, F., Herrfeldt, J. A., Peterson, D. S., Ravetch, J. A., and Wellems, T. E. (1995). The large diverse gene family var encodes proteins involved in cytoadherence and antigenic variation of Plasmodium falciparuminfected erythrocytes. Cell 82, 89–100. Su, X., Ferdig, M. T., Huang, Y., Huynh, C. Q., Liu, A., You, J., Wootton, J. C., and Wellems, T. E. (1999). A genetic map and recombination parameters of the human malaria parasite Plasmodium falciparum. Science 286, 1351–1353. Tanabe, K., Sakihama, N., Rooth, I., Bjorkman, A., and Farnert, A. (2007). High frequency of recombination-driven allelic diversity and temporal variation of Plasmodium falciparum msp1 in Tanzania. Am. J. Trop. Med. Hyg. 76, 1037–1045. Taylor, H. M., Kyes, S. A., Harris, D., Kriek, N., and Newbold, C. I. (2000). A study of var gene transcription in vitro using universal var gene primers. Mol. Biochem. Parasitol. 105, 13–23. Templeton, T. J. (2009). The varieties of gene amplification, diversification and hypervariability in the human malaria parasite, Plasmodium falciparum. Mol. Biochem. Parasitol. 166, 109–116. Tonkin, C. J., Carret, C. K., Duraisingh, M. T., Voss, T. S., Ralph, S. A., Hommel, M., Duffy, M. F., Silva, L. M., Scherf, A., Ivens, A., Speed, T. P., Beeson, J. G., et al. (2009). Sir2 paralogues cooperate to regulate virulence genes and antigenic variation in Plasmodium falciparum. PLoS Biol. 7, e84. Triglia, T., Wellems, T. E., and Kemp, D. J. (1992). Towards a high-resolution map of the Plasmodium falciparum genome. Parasitol. Today 8, 225–229. Voss, T. S., Healer, J., Marty, A. J., Duffy, M. F., Thompson, J. K., Beeson, J. G., Reeder, J. C., Crabb, B. S., and Cowman, A. F. (2005). A var gene promoter controls allelic exclusion of virulence genes in Plasmodium falciparum malaria. Nature 439, 1004–1008. Vosshall, L. B., and Stocker, R. F. (2007). Molecular architecture of smell and taste in Drosophila. Annu. Rev. Neurosci. 30, 505–533. Walliker, D., Quakyi, I. A., Wellems, T. E., McCutchan, T. F., Szarfman, A., London, W. T., Corcoran, L. M., Burkot, T. R., and Carter, R. (1987). Genetic analysis of the human malaria parasite Plasmodium falciparum. Science 236, 1661–1666. Wang, C. W., Hermsen, C. C., Sauerwein, R. W., Arnot, D. E., Theander, T. G., and Lavstsen, T. (2009). The Plasmodium falciparum var gene transcription strategy at the onset of blood stage infection in a human volunteer. Parasitol. Int. 58, 478–480. Wellems, T. E., Panton, L. J., Gluzman, I. Y., Do, R. V., Gwadz, R. W., Walker-Jonah, A., and Krogstad, D. J. (1990). Chloroquine resistance not linked to mdr-like genes in a Plasmodium falciparum cross. Nature 345, 253–255.

CHAPTER

4 Biological Warfare of the Spiny Plant: Introducing Pathogenic Microorganisms into Herbivore’s Tissues Malka Halpern,1 Avivit Waissler, Adi Dror, and Simcha Lev-Yadun

Contents

I. Introduction II. Pathogenic Bacteria and Thorns A. The microflora of spines versus photosynthetic leaf parts B. Pathogenic potential of species identified on spines and thorns III. Silica Needles and Raphids made of Calcium Oxalate A. Silica bodies (phytoliths) B. Raphids C. Biological warfare of plants through raphids and silica needles IV. Cases of Thorn Injuries Reported in Medical Literature A. Bacteria B. Fungi V. Aposematism in Spiny Animals VI. Concluding Remarks References

98 99 102 103 107 108 108 108 108 109 110 110 111 112

Department of Science Education—Biology, Faculty of Natural Sciences, University of Haifa-Oranim, Tivon, Israel 1 Corresponding author: e-mail address: [email protected] Advances in Applied Microbiology, Volume 74 ISSN 0065-2164, DOI: 10.1016/B978-0-12-387022-3.00008-2

#


97

98

Abstract

Malka Halpern et al.

Recently, it has been proposed that plants which have spines, thorns, and prickles use pathogenic aerobic and anaerobic bacteria, as well as pathogenic fungi, for defense against herbivores, especially vertebrates. Their sharp defensive appendages may inject various pathogenic agents into the body of the herbivores by piercing the outer defensive layer of the skin in a type of biological warfare. Here, we review data regarding the various bacterial taxa found on spines, as well as the medical literature regarding infections by bacteria and fungi related to spine injuries. We also present new evidence that, concerning the microbial flora, spines belonging to the palm tree Washingtonia filifera are probably a different habitat than the nondefensive green photosynthetic leaf surfaces. In addition, many plant species have microscopic internal and external spines (raphids and silica needles) which can also wound large herbivores as well as insects and other small invertebrate herbivores that usually attack in between large spines, prickles, and thorns. The large spines and sharp microscopic structures may inject not only the microorganisms that inhabit them into the herbivore’s tissues, but also those preexisting on the skin surface or inside the digestive system of the herbivores and on the surface of nonspiny plant parts. A majority of the spiny plants visually advertise their spiny nature, a characteristic known as aposematism (warning coloration). The pathogenic microorganisms may sometimes be much more dangerous than the physical wounds inflicted by the spines. In accordance, we suggest that the possible cooperation or even just the random association of spines with pathogenic microorganisms contributed to the evolution of aposematism in spiny plants and animals. The role of these sharp defensive structures in inserting pathogenic viruses into the tissues of herbivores was never studied systematically and deserves special attention.

I. INTRODUCTION Plants suffer from herbivory impact by various types of herbivores (Crawley, 1983), resulting in a permanent evolutionary arms race between plants and the herbivores that feed on them. Spines, thorns, and prickles are a common means of antiherbivory physical defense in thousands of plant species (and animals), especially in arid regions (Cooper and OwenSmith, 1986; Gowda, 1996; Grubb, 1992; Janzen, 1986; Janzen and Martin, 1982; Lev-Yadun, 2009a,b; Myers and Bazely, 1991; Rebollo et al., 2002; Ronel et al., 2009, 2010). English botanical usage distinguishes between a prickle (a sharp emergence from the cortex), a spine (a sharp part of a leaf), and a thorn (a sharp branch). They provide physical protection against herbivory by wounding the mouth, the digestive system, and

Biological Warfare of the Spiny Plant

99

other body parts of herbivores ( Janzen, 1986; Janzen and Martin, 1982). Many of these spiny plants are aposematic, that is, the unpalatability of these plants is associated with various types of conspicuous coloration. Recently, it has been proposed that this coloration deters large herbivores (e.g., Lev-Yadun, 2001, 2003a,b, 2006, 2009a,b,c; Lev-Yadun and Neèman, 2004, 2006; Midgley, 2004; Rubino and McCarthy, 2004; Ruxton et al., 2004; Speed and Ruxton, 2005). Phyllosphere is a term used to refer to leaf surfaces or total aboveground surfaces of a plant as a habitat for microorganisms, including filamentous fungi, yeasts, and bacteria (Lindow and Brandl, 2003; Lindow and Leveau, 2002). Recently, it has been proposed that plants use pathogenic aerobic and anaerobic bacteria, as well as pathogenic fungiinhabiting spines, thorns, and prickles for defense against herbivores, especially vertebrates (Halpern et al., 2007a,b). In addition, Lev-Yadun and Halpern (2008) proposed that many plant species have microscopic internal or even external spines which can wound large herbivores, as well as insects and small herbivores that usually attack in between spines and thorns. Here, we review data regarding the various bacterial taxa found on spines, as well as the medical and veterinary literature regarding infections by bacteria and fungi related to spine injuries. We also present new and unpublished evidence that spine phyllosphere may, in certain cases, be a different habitat for bacteria than the nondefensive green photosynthetic leaf phyllosphere. The large spines, thorns, prickles, and sharp microscopic structures may insert not only the microorganisms that inhabit them, but also those that preexist on the skin or inside the digestive system of the herbivores. This review also addresses the potential defensive nature of this phenomenon.

II. PATHOGENIC BACTERIA AND THORNS Halpern et al. (2007a,b) showed recently that spines of Phoenix dactylifera (date palm) and thorns of Crataegus aronia (common hawthorn), Sarcopoterium spinosum (thorny burnet), and Alhagi graecorum (manna tree) harbor an array of pathogenic bacteria. In another (unpublished) study, we demonstrated that spines of the palm tree Washingtonia filifera Wendl. (cotton palm) also harbor pathogenic bacteria (for more details see Section II.A). Every typical mature individual of the tree species carries thousands of conspicuous aposematic thorns or spines (Figs. 4.1A, B and 4.2A, C). Although only a small fraction of thorns or spines found on each tree was sampled, it sufficiently enabled us to isolate and identify various pathogenic bacterial species from these thorns. The bacteria proved to be a common phenomenon (Table 4.1). Twenty-seven different species of the bacterial isolates that were identified from the thorny and spiny trees and

100


A

B

FIGURE 4.1 Thorns and spines of the tree species Phoenix dactylifera (date palm) and Crataegus aronia (common hawthorn). (A) Date palm spines are sharp leaflets that develop along the proximal parts of the large leaves. Their tips are usually conspicuously yellow, red, and black. (B) Common hawthorn thorns are hard branches that end in a sharp thorn and are found all over the tree. They are red when young and turn gray with time.

A

B

C

FIGURE 4.2 (A) Spines and leaves of the tree species Washingtonia filifera (cotton palm). The canopy of a young American palm Washingtonia filifera. The large leaves with their typical ca. 1 m long spiny petioles and fan-shaped photosynthetic area. (B) A closeup of the central photosynthetic leaf area of Washingtonia filifera. (C) Aposematic colorful spines along the petiole of the cotton palm Washingtonia filifera.

TABLE 4.1 Pathogens and opportunistic pathogens species that were isolated from thorns of common hawthorn, date palm, cotton palm, thorny burnet, and manna tree and from leaves of cotton palm Thorny burnet, Common manna tree hawthorn

Date palm

Closest relative in EZtaxon database

Thorn Thorn isolates isolates

Spine Spine Leaf isolates isolates isolates

Reference

Halpern et al. Halpern et al. (2007b) (2007a) Proteobacteria

Escherichia vulneris Shigella boydii Rahnella aquatilis Pantoea agglomerans P. brenneri P. dispersa Pseudomonas stutzeri Acinetobacter johnsonii A. lwoffii Clostridium perfringens C. sordellii C. sardiniense Bacillus anthracis B. thuringiensis B. cereus B. licheniformis B. megaterium B. circulans B. pumilus Exiguobacterium indicum Staphylococcus cohnii S. epidermidis S. equorum S. gallinarum S. hominis S. pasteuri Enterococcus faecalis E. faecium E. mundtii

þ þ þ þ þ þ þ

þ þ þ þ

Current study

þ

þ þ þ

Firmicutes þ

Cotton palm

þ þ þ þ þ

þ

þ

þ þ

þ

þ þ þ þ þ

þ þ

þ þ þ þ þ

þ þ

þ þ

þ þ

þ þ þ þ þ þ þ þ

Actinobacteria Micrococcus luteus Kocuria rosea

þ þ

þ

The identification of the species was based on partial sequences similarities of the 16S rRNA gene to the database in the NCBI gene bank. Similarities to the type strains of the listed species were more than 99%.

102


shrubs were species known to be pathogenic or opportunistic pathogenic to animals or humans (Halpern et al., 2007a,b). The pathogenic or opportunistic pathogenic bacteria that were isolated from date palm and common palm spines and from thorns of common hawthorn, thorny burnet, and manna tree are listed in Table 4.1, columns 2–5 (Halpern et al., 2007a,b and unpublished data).

A. The microflora of spines versus photosynthetic leaf parts To understand whether spines are a different habitat than green photosynthetic leaf surfaces when it comes to the microbial flora, we compared the microbial communities of the phyllosphere on spines to the photosynthetic leaf surfaces of the palm tree W. filifera (unpublished study). Cotton palm trees have a single straight stem that can reach more than 20 m in height and are usually 40–70 cm in diameter (Fig. 4.2A). The petioles, which may reach more than 1 m in length, usually have two rows of sharp spines along their right and left margins (Fig. 4.2C). Every cotton palm tree carries thousands of colorful aposematic spines. The spines may be straight, curved, or recurved and are yellow, orange, or brown in color depending on leaf age and individual tree characteristics, and a panel of the same color connects them along the edge of the petioles, enhancing spine conspicuousness. Spines and green photosynthetic leaf surfaces (Fig. 4.1B and C) were sampled from different trees in Northern Israel in December 2008, March 2009, May 2009, and June 2009 (unpublished). Using culturable methods, we isolated and identified a total of 97 bacterial isolates (accession numbers HM163472–HM163568), which were then classified as pathogens or nonpathogens according to the scientific literature records. Fifteen pathogenic species were identified from spines, and 18 pathogenic species were identified from the green photosynthetic leaf surfaces (Table 4.1, columns 5 and 6). The following pathogenic or opportunistic pathogenic bacteria— Pantoea brenneri, Pseudomonas stutzeri, Acinetobacter johnsonii, Acinetobacter lwoffii, Clostridium sordellii, Bacillus anthracis, Staphylococcus hominis subsp. hominis, and Kocuria rosea—were isolated only from W. filifera spines and were not identified on green photosynthetic leaf surfaces (Table 4.1). In sum (including the nonpathogenic species), 28 bacterial species were found on the spines and 36 species on the green photosynthetic leaf surfaces. Out of these, 11 species were found on both the spines and green photosynthetic leaf surfaces. This data was used to calculate Sorensen’s relatedness index (values are between 0 and 1.0; Wolda, 1981). The similarity between the bacterial flora on green photosynthetic leaf surfaces and spines according to Sorensen’s index was 0.343, indicating that the microflora of the spines in cotton palms is different from that of the green photosynthetic leaf surfaces. Thus, the microbe–spine


103

combination seems to be an important factor in the common evolution of the aposematic coloration of spiny, thorny, and prickly plants. Hence, bacteria that inhabit spines or thorns in plants seem to have enhanced the common, convergent evolution of aposematism in these organisms. When specific pathogenic bacterial species are more common or more diverse on spines, thorns, and prickles than on photosynthetic surfaces, this is a good indication for an evolutionary development in which specific plant surfaces change to host pathogenic bacteria on its sharp defensive appendages. Even if such differences in the composition of pathogenic bacterial species will be found, the coevolution of spiny plants and bacteria will not be proven yet: The selection of surface characteristics that enhance the ability of specific bacteria species to exist may have occurred only in the plant partner, thus not establishing coevolutionary relations. However, if variations that cause a better adaptation of the bacteria to the thorn habitat than to the photosynthetic leaf surfaces can be observed in specific pathogenic bacterial species, this will indicate an established coevolutionary relation (e.g., production of molecules that result in a better adhesion of the bacteria to the thorns than to leaf surfaces). Theoretically, sharp appendages provide better opportunities for bacterial insertion into live animal tissues. Thus, bacteria that are able to exploit live internal animal tissues are expected to have evolved adaptations toward inhabiting sharp appendages (in plants and animals). For instance, Pantoea agglomerans was isolated from thorns (Table 4.1) and at the same time has been proven to cause infections via thorns, spines, or prickles (Table 4.2). This bacterium may have coevolved with palm spines to deter herbivores. However, the whole issue of surface characteristics (chemical and physical) of defensive sharp appendages as a potential advantageous habitat for pathogenic bacteria, fungi, and viruses has never been addressed systematically.

B. Pathogenic potential of species identified on spines and thorns The pathogenic or opportunistic pathogenic bacteria that were isolated from spines of date and cotton palms and from thorns of common hawthorn, thorny burnet, and manna tree are listed in Table 4.1, columns 2–5 (Halpern et al., 2007a,b and unpublished data). Some of the important pathogens that were found to inhabit the aposematic spines and thorns, and which can cause infectious wounds, are discussed here.

1. Clostridium Among the pathogenic isolates from spines and thorns were members of the genus Clostridium. These Gram-positive, endospore-forming obligate anaerobic bacteria cause infections associated with wounds. The species

104


TABLE 4.2 Evidence from the literature indicating microbial infections after thorn injury Microorganism species

Pantoea agglomerans

Enterobacter cloacae Serratia fonticola Pseudomonas aeroginosa C. tetani

Examples for thorn source

References

Bacteria Palm spine, Rose Cruz et al. (2007), De Champs prickle, Lemon et al. (2000), Duerinckx tree thorn, (2008), Flatauer and Khan Hawthorn thorn (1978), Harris (2010), Kratz et al., 2003, Vincent and Szabo (1988), Ulloa-Gutierrez et al. (2004) Rose prickle, Cengiz et al. (2005), Harris Hawthorn thorn (2010) Hawthorn thorn Gorret et al. (2009) Vidyadhara and Rao (2006) Ergonul et al. (2003), Hodes and Teferedegne (1990), Pascual et al. (2003) Cahill and King (1984), Vidyadhara and Rao (2006) McManigal and Henderson (1986) Bakker et al. (2004)

Rose prickle

Staphylococcus aureus Mycobacterium marinum Gordona terrae

Cactus spine

Fusarium solani Fonsecaea pedrosoi

Rose prickle Mimosa pudica spine

Palm spine

NA Fungi

Cladophialophora NA carrionii Sporothrix Rose prickle schenckii Candida Rose prickle parapsilosis

Kantarcioglu et al. (2010) Lo´pez-Martıńez and MeńdezTovar (2007), Salgado et al. (2004) Lo´pez-Martıńez and MeńdezTovar (2007), Son et al. (2010) Engle et al. (2007), Haldar et al. (2007), Ware et al. (1999) Turkal and Baumgardner (1995)

NA, The source of the thorn injury is not available.

that were identified from the thorns and spines were Clostridium perfringens, which is known as a flesh eater (Shimizu et al., 2002) and can produce a necrotizing infection of the skeletal muscle called gas gangrene (Shimizu et al., 2002), and C. sordellii and Clostridium sardiniense, which are


105

considered C. perfringens-like strains and have been isolated from infected tissues in cases of gas gangrene (el Sanousi and Musa, 1989; Masaki et al., 1988; Fig. 4.3, Table 4.1). These species were not identified on green photosynthetic leaf surfaces. Microorganisms can grow on plant surfaces in biofilms (assemblages of bacterial cells attached to a surface and enclosed in adhesive polysaccharides excreted by the cells). Biofilm environment exhibits remarkable heterogeneity. For example, within biofilms, highly aerated zones can border anaerobic zones separated by distances of only tens of microns (Hall-Stoodley and Stoodley, 2005). Members of

100

DQ298076 Clostridium perfringens ATCC 13124T DQ298077 100 DQ298079 DQ298090 DQ298089 98 DQ298091 DQ298078 Clostridium sardiniense DSM 2632T 99 DQ298123 100 DQ298122 Clostridium botulinum ATCC 25763T HM163536

91 100

100

HM163534 Clostridium sulfidigenes SGB2T

Clostridium difficile ATCC 9689T Clostridium sordellii ATCC 9714T 100 HM163489 DQ298113 DQ298112 DQ298111 Escherichia vulneris ATCC 33821T

0.01

FIGURE 4.3 A phylogenetic tree of Clostridium isolates from cotton palm spines (HM163489) and from date palm spines and common hawthorn thorns (names of isolates are accession numbers starting with DQ). The nonpathogenic C. sulfidigenes was isolated from green photosynthetic leaf surfaces (accession numbers HM163536 and HM163534). The tree shows the relationship based on partial sequences of the 16S ribosomal RNA gene of selected isolates. The sequence alignment was performed by use of the CLUSTAL W program and the tree was generated by the neighbor-joining method in MEGA 4 software (Tamura et al., 2007). Bootstrap values (from 1000 replicates) greater than 50% are shown at the branch points. The bar indicates 1% sequence divergence. Escherichia vulneris ATCC 33821T was used as an outgroup.

106


the genus Clostridium, even though they are obligate anaerobes, were found to be common viable inhabitants of the thorns and spines (Halpern et al., 2007a,b and unpublished data; Table 4.1). This can be explained by the fact that they live on the thorn surfaces in biofilms.

2. Bacillus anthracis Species of the Bacillus cereus sensu lato group, which comprises three species—B. anthracis, Bacillus thuringiensis, and B. cereus sensu stricto ( Jensen et al., 2003)—were also isolated from the thorns and spines of all the sampled tree species (Table 4.1). B. anthracis is the etiological agent of anthrax, a notorious acute fatal disease in animals (domesticated and wild, particularly herbivorous) and humans ( Jensen et al., 2003). The cutaneous form of the disease is usually acquired through injured skin or mucous membranes, a typical spine and thorn injury. B. anthracis was isolated and identified from spines of the cotton palm (Table 4.1) and was not isolated from green photosynthetic leaf surfaces. It was also identified on thorns of two thorny shrub species—S. spinosum (thorny burnet) and A. graecorum (manna tree; Halpern et al., 2007b)—as well as from spines of the date palm and thorns of the common hawthorn (Halpern et al., 2007a; Table 4.1). Sudden deaths of wild chimpanzees caused by B. anthracis in a tropical rainforest, Tay National Park in the Ivory Coast, were reported by Leendertz et al. (2004). The source of the chimpanzee’s infection remained a mystery. It is possible that plant thorns were the cause of that incident and other similar enigmatic cases.

3. Pantoea Species of the genus Pantoea, a member of the Enterobacteriaceae, were identified on thorns of common hawthorn and cotton palm spines (Table 4.1, Fig. 4.4). Pantoea species are ubiquitous in nature and occasionally associated with infections. P. agglomerans has been reported in the medical literature as the cause of septic arthritis after palm spine injury (Kratz et al., 2003), as well as the cause of osteomyelitis and synovitis after rose prickle injury (Duerinckx, 2008; Vincent and Szabo, 1988). P. brenneri has recently been identified as a novel species in human clinical samples (Brady et al., 2010). Pantoea dispersa, known formally as Erwinia herbicola, was also reported as pathogenic to humans (Schmid et al., 2003).

4. The pathogenic potential of other bacterial species Enterococcus faecalis and Enterococcus faecium (Table 4.1) can cause various complicated infections (abdominal, skin, urinary tract, and blood; Ruoff et al., 1990). Rahnella aquatilis infections have involved episodes of bacteraemia, urinary tract infections, postsurgical wound infections, and endocarditis. The sources for most Rahnella-related human illnesses remain unknown ( Janda, 2006; Oh and Tay, 1995; Table 4.1). Shigella


99 74

72

107

HM163549 HM163568 Pantoea dispersa LMG 2603T HM163551 Pantoea brenneri LMG 5343T HM163514

98

81

DQ298110

Pantoea agglomerans DSM 3493T Pseudomonas stutzeri CCUG 11256T (U26262)

0.02

FIGURE 4.4 A phylogenetic tree of Pantoea isolates from cotton palm spines (accession numbers HM163549, HM163551, and HM163514) and green photosynthetic leaf surfaces (HM163568) and from common hawthorn (DQ298110). See also Table 4.1. The tree shows the relationship based on partial sequences of the 16S ribosomal RNA gene of selected isolates. The sequence alignment was performed by use of the CLUSTAL W program and the tree was generated by the neighbor-joining method in MEGA 4 software (Tamura et al., 2007). Bootstrap values (from 1000 replicates) greater than 50% are shown at the branch points. The bar indicates 1% sequence divergence. Pseudomonas stutzeri CCUG 11256T was used as an outgroup.

(Table 4.1) is a common cause of diarrheal illnesses and foodborne disease. Shigella boydii was implicated in an outbreak of foodborne illness in the United States (Chan and Blaschek, 2005) and identified as a cause of necrotizing enterocolitis (a serious gastrointestinal disease in neonates; Sawardekar, 2005). Kocuria rosea and Micrococcus luteus (Table 4.1) are members of the family Micrococcaceae (Actinobacteria). These species may cause intracranial abscesses, meningitis, pneumonia, and septic arthritis in immunosuppressed or immunocompetent hosts (Altuntas et al., 2004; Seifert et al., 1995).

III. SILICA NEEDLES AND RAPHIDS MADE OF CALCIUM OXALATE Lev-Yadun and Halpern (2008) proposed that many spiny and nonspiny plant species which have microscopic, sharp internal defensive structures, such as raphids and sharp silica bodies, may also insert pathogenic microorganisms into the tissues of herbivores through the microscopic wounds that these tiny, sharp defensive structures induce, especially in the mouth and digestive system. Small invertebrates may escape most spines, thorns, and prickles by moving between them. However, they may not escape these minute, sharp defensive structures.

108


A. Silica bodies (phytoliths) Silica bodies in plants are formed by an ordered biological deposition of silicon that enters the plant via the roots (Liang et al., 2005; Richmond and Sussman, 2003). Phytoliths are chemically and physically similar to sandpaper and act like it. Silica bodies in plants are found in a rich repertoire of shapes (Meunier and Colin, 2001; Prychild et al., 2004; Rapp and Mulholland, 1992), some of which are needle shaped or otherwise sharp. One of the functions of silica bodies is defense from herbivory by causing teeth wear in mammals and mouth parts deterioration in insects (Baker et al., 1959; Massey and Hartley, 2006; Massey et al., 2007; McNaughton and Tarrants, 1983; McNaughton et al., 1985; Walker et al., 1978) or by wounding animal tissues (e.g., Bhatt et al., 1984; Hodson et al., 1994; Newman, 1986; O’Neill et al., 1980).

B. Raphids Thousands of plant species belonging to many families produce raphids made of calcium oxalate (Franceschi and Horner, 1980). Raphids are always elongated, needle shaped, and have two sharp pointed ends. Focused studies on field behavior of herbivores, such as dorcas gazelles (Gazelle dorcas), have shown that they avoid plant tissues with raphids (Salts and Ward, 2000; Ward et al., 1997).

C. Biological warfare of plants through raphids and silica needles Lev-Yadun and Halpern (2008) hypothesized that both types of microscopic internal spines serve another function in addition to the ability to inject plant toxins into the wounded tissue of herbivores by causing a mechanical irritation: At the same time, they might also be able to insert pathogenic microorganisms. Raphids and silica needles can internally wound the mouth and digestive system of insects and other small herbivores that manage to escape from thorns by passing between them. Through these wounds, microorganisms found on the plant surface or in the mouth and digestive tract of the herbivore may cause infection.

IV. CASES OF THORN INJURIES REPORTED IN MEDICAL LITERATURE There are indications from medical case reports that injuries from plant thorns can result in septic inflammation. None of the published data concerning this phenomenon has been related to ecological or evolutionary


109

issues, or aposematism; the relevant articles were published for medical practice. However, the information showed that plant thorns may regularly harbor various toxic or pathogenic microorganisms (either bacteria or fungi; e.g., Cahill and King, 1984; De Champs et al., 2000; Engle et al., 2007; Ergonul et al., 2003; Freiberg et al., 1993; Haldar et al., 2007; Hodes and Teferedegne, 1990; Kratz et al., 2003; Pascual et al., 2003; Sugarman et al., 1977; Vincent and Szabo, 1988). In Israel, for instance, orchard workers who were wounded by palm spines have experienced infections of such severity and frequency that the costly practice of removing all the millions of spines from palm trees by mechanical saws was adopted in many orchards (Halpern et al., 2007a). Cases reporting plant thorn injuries that have resulted in septic inflammation are summarized in Table 4.2.

A. Bacteria Here, we review indications from medical case reports that plant thorn injuries can result in septic inflammation caused by bacteria (Table 4.2). Clostridium tetani, for instance, (a Gram-positive, spore-forming anaerobic bacterium), is the etiological agent of tetanus, a serious disease in humans and animals that can be fatal when untreated. The national tetanus surveillance in the United States reported that 26 (31%) cases of tetanus in 1998– 2000 were caused by injuries sustained by the patient while farming or gardening. Rose bush prickles were one of the causes of puncture wounds that resulted in tetanus (Pascual et al., 2003). In Ethiopia, thorn injuries were the known cause of 5 out of 55 cases of tetanus (Hodes and Teferedegne, 1990). Thorn injury was also reported to be the cause of tetanus in Turkey (Ergonul et al., 2003). P. agglomerans, which has been isolated from common hawthorn thorns (Halpern et al., 2007a; Table 4.1), has been reported in the medical literature as the cause of septic arthritis after palm spine injury (Kratz et al., 2003), as well as the cause of osteomyelitis after rose prickle injury (Vincent and Szabo, 1988). Enterobacter cloacae was reported as the cause of a prolonged cellulitis in a 5-year-old healthy boy due to plant thorn injury (Cengiz et al., 2005). In another medical case, a deep infection caused by E. cloacae and P. agglomerans was associated with a small piece of retained hawthorn thorn (Harris, 2010). Serratia fonticola infection was reported in a child with septic arthritis, who had fallen off his bicycle and sustained an infection with hawthorn thorns (Gorret et al., 2009). The infection was eradicated only after surgical drainage and removal of a remaining thorn. Pseudomonas aeruginosa and Staphylococcus aureus were reported to be the cause of thorn prick osteomyelitis of the foot in barefoot walkers in India (Vidyadhara and Rao, 2006). Gordona terrae (actinomyces) was isolated and identified as the cause of mycetoma of the hand in a 15-year-old boy from Sierra Leone who had pricked the base of his thumb with a thorn while in the jungle (Bakker et al., 2004). Other inflammatory

110


states of the musculoskeletal system have similarly been associated with specific pathogens introduced into animal tissues by various thorn injuries, such as Staphylococcus aureus, S. albus, Streptococcus b hemolyticus, and Nocardia pyarthrosis (Cahill and King, 1984; De Champs et al., 2000; Freiberg et al., 1993; Kratz et al., 2003; Sugarman et al., 1977; Table 4.2).

B. Fungi Not only bacteria can cause septic inflammation through plant thorn injury, but also there are strong indications in the medical literature that thorns, spines, or prickles can introduce pathogenic fungi into animals or humans (Halpern et al., 2007b; Table 4.2). Dermatophytes that cause subcutaneous mycoses are unable to penetrate the skin. They must be inserted into the subcutaneous tissue by a puncture wound (Willey et al., 2008). One type of subcutaneous mycosis is chromoblastomycosis, caused by pigmented or dematiaceous saprophytic molds. The most common etiologic agents are Fonsecaea pedrosoi and Cladophialophora carrionii, both of which can be isolated from plants. Infection is acquired by inoculation of the etiologic agent into the subcutaneous tissues of the subject by penetrating thorns or spines of diverse plants (Lo´pez-Martıńez and Meńdez-Tovar, 2007; Son et al., 2010). Salgado et al. (2004) reported the isolation of F. pedrosoi from spines of the plant Mimosa pudica at the place of infection identified by one of their patients. Another subcutaneous mycosis is sporotrichosis, caused by the fungus Sporothrix schenckii. This disease occurs throughout the world and is the most common subcutaneous mycotic disease in the United States. The disease is an occupational hazard for florists, gardeners, and forestry workers; it is also known as the rose-gardener’s disease as it is commonly transmitted through a prick from rose prickles (Engle et al., 2007; Haldar et al., 2007; Ware et al., 1999). Mycetoma is a chronic, specific, granulomatous, progressive subcutaneous inflammatory disease with a global distribution. Abscesses in the skin can spread to the bones and muscles. The disease is caused by true fungi or by filamentous bacteria and hence it is classified as Eumycetoma and Actinomycetoma, respectively (Fahal, 2004). These fungi or bacteria gain access to the tissues via wooden splinters or thorns.

V. APOSEMATISM IN SPINY ANIMALS The phenomenon of biological defense through spines that harbor pathogenic bacteria is not confined to plants. Various bacteria defend animals from predation, and this defense is sometimes associated with spines. For example, Vibrio vulnificus causes severe and often fatal infections in humans through contamination of wounds, typically in patients injured


111

while handling pond-cultivated fish (Bisharat et al., 1999). Bacteria producing tetrodotoxin (a strong neurotoxin) was isolated from the spine apparatus of the Caribbean sea urchin, Meoma ventricosa (Ritchie et al., 2000). Pathogenic bacteria-harboring animal spines may also be associated with the evolution of aposematism in spiny animals (e.g., Inbar and Lev-Yadun, 2005; Ruxton et al., 2004; Speed and Ruxton, 2005).

VI. CONCLUDING REMARKS The physical defense against herbivores provided by thorns, spines, prickles, silica needles, and raphids might be only the tip of the iceberg in a much more complicated story. The various sharp external defensive plant structures act by wounding and inserting pathogenic microorganisms (bacteria and fungi) into the body of the herbivores as a sort of natural injection. These may cause severe infections in the herbivores that are much more dangerous and painful than the mechanical wounding itself. These thorn-inhabiting microorganisms may have uniquely contributed to the defensive function of sharp plant appendages and internal structures and thus to the common evolution of aposematism (warning coloration) in thorny, spiny, and prickly plants or of plants that have internal microscopic spines. Moreover, there is a possibility that microorganisms-inhabiting plant thorns secrete toxins while multiplying on the plant and that those toxins can also potentially harm herbivores. While the pain from contacting thorns is immediate, the effect of the microorganisms is delayed. This delay between initial contact and wounding and the microorganism’s effect may give rise to the question of how efficient this protection process can be. Yet, the same is true for the delayed action of poisons in the numerous known aposematic poisonous organisms. Nevertheless, there is a general agreement that many poisonous and colorful organisms are aposematic (e.g., Cott, 1940; Edmunds, 1974; Gittleman and Harvey, 1980; Harvey and Paxton, 1981; Lev-Yadun, 2009a; Ruxton et al., 2004). Therefore, there is no reason to view the contamination by microorganisms and its delayed effect any differently. The microbe–thorn/spine/prickle combination seems to be an important factor in the common evolution of the aposematic coloration of thorny/ spiny/prickly plants (Halpern et al., 2007a,b; Lev-Yadun and Halpern, 2008). Hence, bacteria that inhabit thorns or spines in plants and animals alike seem to have enhanced the common, convergent evolution of aposematism in these spiny organisms. In order to provide more evidence regarding the hypothesis of a common evolution of microbes and plants, numerous studies on the bacterial and fungi population dynamics on young and mature thorns, spines, and prickles of many plant and animal species should be performed using molecular tools such as pyrosequencing. At the same time,

112


the thorns, spines, and prickles should be compared with the smooth surfaces of these organisms. The nature of biofilms on thorns, spines, and prickles is practically unknown and careful studies of such biofilms are also needed. The possibility that bacteria secrete toxins against herbivores on plant surfaces also deserves special attention. If certain bacteria use plant surfaces as a habitat, they may have evolved mechanisms to conserve it by damaging herbivores in various ways to defend their habitat, a phenomenon known from ants that protect plants (e.g., Huxley and Cutler, 1991; Jolivet, 1998) and mutualistic fungi (Clay, 1990; Lev-Yadun and Halpern, 2007). The role of sharp defensive structures in inserting pathogenic viruses into the tissues of herbivores was never studied in an ecological–evolutionary context and also deserves special attention.

REFERENCES Altuntas, F., Yildiz, O., Eser, B., Gu¨ndogan, K., Sumerkan, B., and Cetin, M. (2004). Catheterrelated bacteremia due to Kocuria rosea in a patient undergoing peripheral blood stem cell transplantation. J. Infect. Dis. 22, 62. Baker, G., Jones, L. H. P., and Wardrop, I. D. (1959). Cause of wear in sheep’s teeth. Nature 184, 1583–1584. Bakker, X. R., Spauwen, P. H., and Dolmans, W. M. (2004). Mycetoma of the hand caused by Gordona Terrae: A case report. J. Hand Surg. Br. 29, 188–190. Bhatt, T., Coombs, M., and O’Neill, C. (1984). Biogenic silica fiber promotes carcinogenesis in mouse skin. Int. J. Cancer 34, 519–528. Bisharat, N., Agmon, V., Finkelstein, R., Raz, R., Ben-Dror, G., Lerner, L., Soboh, S., Colodner, R., Cameron, D. N., Wykstra, D. L., Swerdlow, D., and Farmer, J. J., III (1999). Clinical, epidemiological, and microbiological features of Vibrio vulnificus biogroup 3 causing outbreaks of wound infection and bacteraemia in Israel. Lancet 354, 1421–1424. Brady, C. L., Cleenwerck, I., Venter, S. N., Engelbeen, K., de Vos, P., and Coutinho, T. A. (2010). Emended description of the genus Pantoea, description of four species from human clinical samples, Pantoea septica sp. nov., Pantoea eucrina sp. nov., Pantoea brenneri sp. nov. and Pantoea conspicua sp. nov., and transfer of Pectobacterium cypripedii (Hori, 1911) Brenner et al. 1973 emend. Hauben et al. 1998 to the genus as Pantoea cypripedii comb. nov. Int. Syst. Evol. Microbiol. 60, 2430–2440. Cahill, N., and King, J. D. (1984). Palm thorn synovitis. J. Pediatr. Orthop. 4, 175–179. Cengiz, A. B., Kanra, G., Celik, T., Co¨mert, B., Devrim, I., Ozen, M., and Kara, A. (2005). Prolonged cellulitis due to plant thorn. Turk. J. Pediatr. 47, 393–396. Chan, Y. C., and Blaschek, H. P. (2005). Comparative analysis of Shigella boydii 18 foodborne outbreak isolate and related enteric bacteria: Role of rpoS and adiA in acid stress response. J. Food Prot. 68, 521–527. Clay, K. (1990). Fungal endophytes of grasses. Annu. Rev. Ecol. Syst. 21, 275–297. Cooper, S. M., and Owen-Smith, N. (1986). Effects of plant spinescence on large mammalian herbivores. Oecologia 68, 446–455. Cott, H. B. (1940). Adaptive Coloration in Animals. Methuen and Co Ltd, London. Crawley, M. J. (1983). Herbivory: The Dynamics of Animal-Plant Interactions. Blackwell Scientific Publications, Oxford. Cruz, A. T., Cazacu, A. C., and Allen, C. H. (2007). Pantoea agglomerans, a plant pathogen causing human disease. J. Clin. Microbiol. 45, 1989–1992.


113

De Champs, C., Le Seaux, S., Dubost, J. J., Boisgard, S., Sauvezie, B., and Sirot, J. (2000). Isolation of Pantoea agglomerans in two cases of septic monoarthritis after plant thorn and wood sliver injuries. J. Clin. Microbiol. 38, 460–461. Duerinckx, J. F. H. (2008). Subacute Synovitis of the knee after a rose thorn injury: Unusual clinical picture. Clin. Orthop. Relat. Res. 466, 3138–3142. Edmunds, M. (1974). Defence in Animals: A Survey of Anti-Predator Defences. Longman Group Ltd, Harlow. el Sanousi, S. M., and Musa, M. T. (1989). Note on an association of Clostridium novyi type A and Clostridium sordellii with a case of gas-gangrene in a Zebu cow. Rev. E´lev. Me´d. Ve´t. Pays Trop. 42, 391–392. Engle, J., Desir, J., and Bernstein, J. M. (2007). A rose by any other name. Skinmed 6, 139–141. Ergonul, O., Erbay, A., Eren, S., and Dokuzoguz, B. (2003). Analysis of the case fatality rate of tetanus among adults in a tertiary hospital in Turkey. Eur. J. Clin. Microbiol. Infect. Dis. 22, 188–190. Fahal, A. H. (2004). Mycetoma: A thorn in the flesh. Trans. R. Soc. Trop. Med. Hyg. 98, 3–11. Flatauer, F. E., and Khan, M. A. (1978). Septic arthritis caused by Enterobacter agglomerans. Arch. Intern. Med. 138, 788. Franceschi, V. R., and Horner, H. T., Jr. (1980). Calcium oxalate crystals in plants. Bot. Rev. 46, 361–427. Freiberg, A. A., Herzenberg, J. E., and Sangeorzan, J. A. (1993). Thorn synovitis of the knee joint with Nocardia pyarthrosis. Clin. Orthop. 287, 233–236. Gittleman, J. L., and Harvey, P. H. (1980). Why are distasteful prey not cryptic? Nature 286, 149–150. Gorret, J., Chevalier, J., Gaschet, A., Fraisse, B., Violas, P., Chapuis, M., and JolivetGougeon, A. (2009). Childhood delayed septic arthritis of the knee caused by Serratia fonticola. Knee 16, 512–514. Gowda, J. H. (1996). Spines of Acacia tortilis: What do they defend and how? Oikos 77, 279–284. Grubb, P. J. (1992). A positive distrust in simplicity—Lessons from plant defences and from competition among plants and among animals. J. Ecol. 80, 585–610. Haldar, N., Sharma, M. K., and Gugnani, H. C. (2007). Sporotrichosis in north-east India. Mycoses 50, 201–204. Hall-Stoodley, L., and Stoodley, P. (2005). Biofilm formation and dispersal and the transmission of human pathogens. Trends Microbiol. 13, 7–10. Halpern, M., Raats, D., and Lev-Yadun, S. (2007a). Plant biological warfare: Infecting pathogenic bacteria into herbivores by thorns. Environ. Microbiol. 9, 584–592. Halpern, M., Raats, D., and Lev-Yadun, S. (2007b). The potential anti-herbivory role of microorganisms on plant thorns. Plant Signal. Behav. 2, 503–504. Harris, E. J. (2010). Retained hawthorn fragment in a child’s foot complicated by infection: Diagnosis and excision aided by localization with ultrasound. J. Foot Ankle Surg. 49, 161–165. Harvey, P. H., and Paxton, R. J. (1981). The evolution of aposematic coloration. Oikos 37, 391–396. Hodes, R. M., and Teferedegne, B. (1990). Tetanus in Ethiopia: Analysis of 55 cases from Addis Ababa. East Afr. Med. J. 67, 887–893. Hodson, M. J., Smith, R. J., van Blaaderen, A., Crafton, T., and O’Neill, C. H. (1994). Detecting plant silica fibres in animal tissue by confocal fluorescence microscopy. Ann. Occup. Hyg. 38, 149–160. Huxley, C. R., and Cutler, D. F. (1991). Ant-Plant Interactions. Oxford University Press, London. Inbar, M., and Lev-Yadun, S. (2005). Conspicuous and aposematic spines in the animal kingdom. Naturwissenschaften 92, 170–172.

114


Janda, M. J. (2006). New members of the family Enterobacteriaceae. In ‘‘The Prokaryotes’’ (M. Dworkin, S. Falkow, E. Rosenberg, K. H. Schleifer and E. Stackebrandt, Eds.), Vol. 6, pp. 5–40. Springer, Singapore. Janzen, D. H. (1986). Chihuahuan Desert nopaleras: Defaunated big mammal vegetation. Annu. Rev. Ecol. Syst. 17, 595–636. Janzen, D. H., and Martin, P. S. (1982). Neotropical anachronisms: The fruits the gomphotheres ate. Science 215, 19–27. Jensen, G. B., Hansen, B. M., Eilenberg, J., and Mahillon, J. (2003). The hidden lifestyles of Bacillus cereus and relatives. Environ. Microbiol. 5, 631–640. Jolivet, P. (1998). Interrelationship Between Insects and Plants. CRC Press, Boca Raton. Kantarcioglu, A. S., Summerbell, R. C., Sutton, D. A., Yu¨cell, A., Sarikaya, E., Kaner, G., Iscimen, A., and Altas, K. (2010). A dark strain in the Fusarium solani species complex isolated from primary subcutaneous sporotrichioid lesions associated with traumatic inoculation via a rose bush thorn. Med. Mycol. 48, 103–109. Kratz, A., Greenberg, D., Barki, Y., Cohen, E., and Lifshitz, M. (2003). Pantoea agglomerans as a cause of septic arthritis after palm tree thorn injury; case report and literature review. Arch. Dis. Child. 88, 542–544. Leendertz, F. H., Ellerbrok, H., Boesch, C., Couacy-Hymann, E., Matz-Rensing, K., Hakenbeck, R., Bergmann, C., Abaza, P., Junglen, S., Moebius, Y., Vigilant, L., Formenty, P., et al. (2004). Anthrax kills wild chimpanzees in a tropical rainforest. Nature 430, 451–452. Lev-Yadun, S. (2001). Aposematic (warning) coloration associated with thorns in higher plants. J. Theor. Biol. 210, 385–388. Lev-Yadun, S. (2003a). Why do some thorny plants resemble green zebras? J. Theor. Biol. 244, 483–489. Lev-Yadun, S. (2003b). Weapon (thorn) automimicry and mimicry of aposematic colorful thorns in plants. J. Theor. Biol. 244, 183–188. Lev-Yadun, S. (2006). Defensive coloration in plants: A review of current ideas about antiherbivore coloration strategies. In ‘‘Floriculture, Ornamental and Plant Biotechnology: Advances and Topical Issues’’ ( J. A. Teixeira da Silva, Ed.), Vol. 4, pp. 292–299. Global Science Books, London. Lev-Yadun, S. (2009a). Aposematic (warning) coloration in plants. In ‘‘Plant-Environment Interactions. From Sensory Plant Biology to Active Plant Behavior’’ (F. Baluska, Ed.), pp. 167–202. Springer-Verlag, Berlin. Lev-Yadun, S. (2009b). Mu¨llerian and Batesian mimicry rings of white-variegated aposematic spiny and thorny plants: A hypothesis. Isr. J. Plant Sci. 57, 107–116. Lev-Yadun, S. (2009c). Mu¨llerian mimicry in aposematic spiny plants. Plant Signal. Behav. 4, 482–483. Lev-Yadun, S., and Halpern, M. (2007). Ergot (Claviceps purpurea)—An aposematic fungus. Symbiosis J. 43, 105–108. Lev-Yadun, S., and Halpern, M. (2008). External and internal spines in plants insert pathogenic microorganisms into herbivore’s tissues for defense. In ‘‘Microbial Ecology Research Trends’’ (T. Van Dijk, Ed.), pp. 155–168. Nova Scientific Publishers, Inc., New York. Lev-Yadun, S., and Neèman, G. (2004). When may green plants be aposematic? Biol. J. Linn. Soc. 81, 413–416. Lev-Yadun, S., and Neèman, G. (2006). Color changes in old aposematic thorns, spines, and prickles. Isr. J. Plant Sci. 54, 327–333. Liang, Y., Si, J., and Ro¨mheld, V. (2005). Silicon uptake and transport is an active process in Cucumis sativus. New Phytol. 167, 797–804. Lindow, S. E., and Brandl, M. T. (2003). Microbiology of the Phyllosphere. Appl. Environ. Microbiol. 69, 1875–1883.


115

Lindow, S. E., and Leveau, J. H. (2002). Phyllosphere microbiology. Curr. Opin. Biotechnol. 13, 238–243. Lo´pez-Martıńez, R., and Meńdez-Tovar, L. J. (2007). Chromoblastomycosis. Clin. Dermatol. 25, 188–194. Masaki, T., Umehashi, H., Miyazaki, H., Takano, M., Yamakawa, K., and Nakamura, S. (1988). Clostridium absonum from gas gangrene. Jpn. J. Med. Sci. Biol. 41, 27–30. Massey, F. P., and Hartley, S. E. (2006). Experimental demonstration of the antiherbivore effects of silica in grasses: Impacts on foliage digestibility and vole growth rates. Proc. R. Soc. Lond. B 273, 2299–2304. Massey, F. P., Ennos, A. R., and Hartley, S. E. (2007). Grasses and the resource availability hypothesis: The importance of silica-based defenses. J. Ecol. 95, 414–424. McManigal, S. A., and Henderson, J. C. (1986). Mycobacterium marinum infection associated with cactus spine injury. J. Med. Technol. 3, 235–236. McNaughton, S. J., and Tarrants, J. L. (1983). Grass leaf silicification: Natural selection for an inducible defense against herbivores. Proc. Natl. Acad. Sci. USA 80, 790–791. McNaughton, S. J., Tarrants, J. L., McNaughton, M. M., and Davis, R. H. (1985). Silica as a defense against herbivory and a growth promotor in African grasses. Ecology 66, 528–535. Meunier, J. D., and Colin, F. (2001). Phytoliths: Application in Earth Sciences and Human History. A. A. Balkema Publishers, Liege. Midgley, J. J. (2004). Why are spines of African Acacia species white? Afr. J. Range For. Sci. 21, 211–212. Myers, J. H., and Bazely, D. (1991). Thorns, spines, prickles, and hairs: Are they stimulated by herbivory and do they deter herbivores? In ‘‘Phytochemical Induction by Herbivores’’ (D. W. Tallamy and M. J. Raupp, Eds.), pp. 325–344. John Wiley and Sons, Inc., New York. Newman, R. H. (1986). Fine biogenic silica fibres in sugar cane: A possible hazard. Ann. Occup. Hyg. 30, 365–370. O’Neill, C. H., Hodges, G. M., Riddle, P. N., Jordan, P. W., Newman, R. H., Flood, R. J., and Toulson, E. C. (1980). A fine fibrous silica contaminant of flour in the high oesophageal cancer area of north-east Iran. Int. J. Cancer 26, 617–628. Oh, H. M. L., and Tay, L. (1995). Bacteraemia caused by Rahnella aquatilis: Report of two cases and review. Scand. J. Infect. Dis. 27, 79–80. Pascual, F. B., McGinley, E. L., Zanardi, L. R., Cortese, M. M., and Murphy, T. V. (2003). Tetanus surveillance, United States, 1998–2000. MMWR Surveill. Summ. 52(SS03), 1–8, http://www.cdc.gov/mmwr/preview/mmwrhtml/ss5203a1.htm. Prychild, C. J., Rudall, P. J., and Gregory, M. (2004). Systematics and biology of silica bodies in monocotyledons. Bot. Rev. 69, 377–440. Rapp, G., Jr., and Mulholland, S. C. (1992). Phytolith systematics. Emerging issues. Plenum Press, New York. Rebollo, S., Milchunas, D. G., Noy-Meir, I., and Chapman, P. L. (2002). The role of spiny plant refuge in structuring grazed shortgrass steppe plant communities. Oikos 98, 53–64. Richmond, K. E., and Sussman, M. (2003). Got silicon? The non-essential beneficial plant nutrient Curr. Opin. Plant Biol. 6, 268–272. Ritchie, K. B., Nagelkerken, I., James, S., and Smith, G. W. (2000). A tetrodotoxin-producing marine pathogen. Nature 404, 354. Ronel, M., Khateeb, S., and Lev-Yadun, S. (2009). Protective spiny modules in thistles of the Asteraceae in Israel. J. Torrey Bot. Soc. 136, 46–56. Ronel, M., Neèman, G., and Lev-Yadun, S. (2010). Spiny east-Mediterranean plant species flower later and in a drier season than non-spiny species. Flora 205, 276–281. Rubino, D. L., and McCarthy, B. C. (2004). Presence of aposematic (warning) coloration in vascular plants of southeastern Ohio. J. Torrey Bot. Soc. 131, 252–256. Ruoff, K. L., de la Maza, L., Murtagh, M. J., Spargo, J. D., and Ferraro, M. J. (1990). Species identities of enterococci isolated from clinical specimens. J. Clin. Microbiol. 28, 435–437.

116


Ruxton, G. D., Sherratt, T. N., and Speed, M. P. (2004). Avoiding Attack. The Evolutionary Ecology of Crypsis Warning Signals, and Mimicry. Oxford University Press, Oxford. Salgado, C. G., da Silva, J. P., Diniz, J. A., da Silva, M. B., da Costa, P. F., Teixeira, C., and Salgado, U. I. (2004). Isolation of Fonsecaea pedrosoi from thorns of Mimosa pudica, a probable natural source of chromoblastomycosis. Rev. Inst. Med. Trop. Saõ Paulo 46, 33–36. Salts, D., and Ward, D. (2000). Responding to a three-pronged attack: Desert lilies subject to herbivory by dorcas gazelles. Plant Ecol. 148, 127–138. Sawardekar, K. P. (2005). Shigellosis caused by Shigella boydii in a preterm neonate, masquerading as necrotizing enterocolitis. J. Pediatr. Infect. Dis. 24, 184–185. Schmid, H., Schubert, S., Weber, C., and Bogner, J. R. (2003). Isolation of a Pantoea dispersalike strain fron a 71-year-old woman with acute myeloid leukemia and multiple myeloma. Infection 31, 66–67. Seifert, H., Kaltheuner, M., and Perdreau-Remington, F. (1995). Micrococcus luteus endocarditis: Case report and review of the literature. Zentralbl. Bakteriol. 282, 431–435. Shimizu, T., Ohtani, K., Hirakawa, H., Ohshima, K., Yamashita, A., Shiba, T., Ogasawara, N., Hattori, M., Kuhara, S., and Hayashi, H. (2002). Complete genome sequence of Clostridium perfringens, an anaerobic flesh-eater. Proc. Natl. Acad. Sci. USA 99, 996–1001. Son, Y.-M., Kang, H.-K., Na, S.-Y., Lee, H.-Y., Baek, J.-O., Lee, J.-R., Roh, J.-Y., and Seo, Y.-H. (2010). Chromoblastomycosis Caused by Phialophora richardsiae. Ann. Dermatol. 22, 362–366. Speed, M. P., and Ruxton, G. D. (2005). Warning displays in spiny animals: One (more) evolutionary route to aposematism. Evolution 59, 2499–2508. Sugarman, M., Stobie, D. G., Quismorio, F. P., Terry, R., and Hanson, V. (1977). Plant thorn synovitis. Arthritis Rheum. 20, 1125–1128. Tamura, K., Dudley, J., Nei, M., and Kumar, S. (2007). MEGA4: Molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24, 1596–1599. Turkal, N. W., and Baumgardner, D. J. (1995). Candida parapsilosis infection in a rose thorn wound. J. Am. Board Fam. Pract. 8, 484–485. Ulloa-Gutierrez, R., Moya, T., and Avila-Aguero, M. L. (2004). Pantoea agglomerans and thornassociated suppurative arthritis. Pediatr. Infect. Dis. J. 23, 690. Vidyadhara, S., and Rao, S. K. (2006). Thorn prick osteomyelitis of the foot in barefoot walkers: A report of four cases. J. Orthop. Surg. 14, 222–224. Vincent, K., and Szabo, R. M. (1988). Enterobacter agglomerans osteomyelitis of the hand from a rose thorn. A case report. Orthopedics 11, 465–467. Walker, A., Hoeck, H. N., and Perez, L. (1978). Microwear of mammalian teeth as an indicator of diet. Science 201, 908–910. Ward, D., Spiegel, M., and Saltz, D. (1997). Gazelle herbivory and interpopulation differences in calcium oxalate content of leaves of a desert lily. J. Chem. Ecol. 23, 333–346. Ware, A. J., Cockerell, C. J., and Skiest, D. J. (1999). Kussman HM disseminated sporotrichosis with extensive cutaneous involvement in a patient with AIDS. J. Am. Acad. Dermatol. 40, 350–355. Willey, J. M., Sherwood, L. M., and Woolverton, C. J. (2008). Prescot, Harley, and Klein’s Microbiology. 7th edn. McGraw Hill, New York. Wolda, H. (1981). Similarity indices, sample size and diversity. Oecologia 50, 296–302.

INDEX A Aerobic aromatic degradation C23O inactivation catecholic compounds, 7 cellular toxicity, 6 kinetic properties, enzymes, 8 ROS, 7 genetic organization and regulation, 8–9 lower pathway meta-cleavage, 6 meta-fission, 6–7 ortho-cleavage, 5 schematic representation, 4 upper pathway, 5 Aromatic compounds, bacterial growth aerobic degradation (see Aerobic aromatic degradation) catabolic genes (see Catabolic genes) enzyme substrate range/kinetics base pair mutations, 20 C23O, 20 catabolic enzyme inactivation, 21–22 catechol, 22–23 TDO, 22–24 repair enzymes cellular stress reduction, 18–20 enzymes reactivation, 17–18 B Bacillus anthracis, 106 Base pair mutations, 20–21 Biological warfare. See Spiny plant C C23O inactivation catecholic compounds, 7 cellular toxicity, 6 kinetic properties, enzymes, 8 ROS, 7 Catabolic enzyme inactivation, 21–22 Catabolic genes horizontal recruitment

conjugative transposons and genomic islands, 16–17 plasmid transfer, 15–16 vertical recruitment Comamonas testosteroi, 11 CymR binding, 11 gene dosage, 12–13 genetic rearrangement and gene capture, 13–14 mutations, 10 phenol degradation operon, 11–12 Pseudomonas azelaica, 10–11 Clostridium biofilm environment, 105 C. perfringens, 104 flesh eater, 104 gas gangrene, 104 phylogenetic tree, 105 Comamonas testosteroi, 11 E Enterococcus faecalis, 106 Enterococcus faecium, 106 Enzymes reactivation, 17–18 G Glutathoine S-transferases (GSTs), 19 Glycoprotein precursor (GPC) domain I and II, 50–51 Golgi complex, 51 N-linked glycosylation, 50 plasma membrane, 51 schematic illustration, 50 virus biology host–pathogen interactions, 52 receptor-mediated endocytosis, 51 viral assembly, 52 virulence, 52–53 H Hantavirus antiviral therapeutic agents, 60 Bunyaviridae family, 37

117

118

Index

Hantavirus (cont.) classification of, 37 disease diagnosis of, 57–58 epidemiology, 53 in human, 55–56 pathogenesis, 56–57 reservoir hosts, 54–55 therapy, 58 vaccines, 58–59 genetic system, 60–61 glycoproteins Gn and Gc (see Glycoprotein precursor) H1N1, 59 HCPS, 36–38, 56 HFRS, 36–37 HPS, 36 molecular biology nucleocapsid protein (see Nucleocapsid protein) replication cycle, 39–41 structure, 38–39 RdRp (see RNA-dependent RNA polymerase) rodent hosts, 60 translation initiation mechanism, 60 Hantavirus cardiopulmonary syndrome (HCPS), 36–38, 56 Hantavirus pulmonary syndrome (HPS), 36 Hemorrhagic fever with renal syndrome (HFRS), 55. See also Hantavirus M Malaria epigenetics, 89–90 immunity protection, 78–79 intraerythrocytic infection, 78–79 pathogenesis human red blood cells, 80–81 IE’s capacity, 81 Plasmodium knowlesi, 80 Meta-cleavage, 6 Meta-fission, 6–7 mRNA synthesis, 48 N N-linked glycosylation, 50 Nucleocapsid protein cellular proteins interaction, 44–45 transcription and translation, 43–44 viral RNA encapsidation and assembly, 42–43

O Ortho-cleavage, 5 P Plasmodium falciparum erythrocyte membrane protein 1 (PfEMP1). See also Malaria antigen expression, 86–87 epigenetics, 89–90 genetics and genomics chimpanzee blood infections, 83 features of, 84 genome replication, 83 haploid forms, 82–83 mitotic recombination, 83 MEE models, 87–89 pathogenesis human red blood cells, 80–81 IE’s capacity, 81 Plasmodium knowlesi, 80 pulsed field gel (PFG) electrophoresis, 84 structure of, 85–86 transcription, 86–87 var genes, 85–86 (see also var genes) Pantoea, 106–107 Phoenix dactylifera, 99–100 Phyllosphere, 99 Phytoliths, 108 Plasmodium knowlesi, 80 Pseudomonas azelaica, 10–11 Pulsed field gel (PFG) electrophoresis, 84 R Raphids, 108 Reactive oxygen species (ROS), 7 Repair enzymes cellular stress reduction, 18–20 enzymes reactivation, 17–18 Replication cycle b1 and b3, 40 mRNAs, 40–41 viral RdRp, 40–41 virus particles attachment, 39 RNA-dependent RNA polymerase (RdRp) Asp residues, 47 conserved structural motifs, 46 localization, 47 mechanism of action, 47–48 mRNA synthesis, 48 polymerases structure, 46 polymerization, 49

119

Index

RNA recombination, 49–50 RNA replication, 49 schematic representation, 45 S Shigella boydii, 106–107 Sillica needles biological warfare of, 108 phytoliths, 108 raphids, 108 Spiny animals, 110–111 Spiny plant pathogenic bacteria and thorns aposematic thorns, 99–100 Bacillus anthracis, 106 Clostridium (see Clostridium) Enterococcus faecalis, 106 Enterococcus faecium, 106 microflora vs. photosynthetic leaf parts, 102–103 palm spines, 102 Pantoea, 106–107 Phoenix dactylifera, 99–100 Shigella boydii, 106–107 Washingtonia filifera, 99–100 phyllosphere, 99 sillica needles biological warfare of, 108 phytoliths, 108

raphids, 108 thorn injury bacteria, 109–110 fungi, 110 septic inflammation, 108 T Thorn injury bacteria, 109–110 fungi, 110 septic inflammation, 108 Toluene dioxygenase (TDO), 22–24 V var genes MEE models, 87–89 P. falciparum, 85–86 transcription, 86–87 Virus assembly, 52 hantavirus (see Hantavirus) host–pathogen interactions, 52 receptor-mediated endocytosis, 51 replication cycle, 39–41 virulence, 52–53 W Washingtonia filifera, 99–100

CONTENTS OF PREVIOUS VOLUMES Volume 40 Microbial Cellulases: Protein Architecture, Molecular Properties, and Biosynthesis Ajay Singh and Kiyoshi Hayashi Factors Inhibiting and Stimulating Bacterial Growth in Milk: An Historical Perspective D. K. O’Toole Challenges in Commercial Biotechnology. Part I. Product, Process, and Market Discovery Alesˇ Prokop Challenges in Commercial Biotechnology. Part II. Product, Process, and Market Development Alesˇ Prokop Effects of Genetically Engineered Microorganisms on Microbial Populations and Processes in Natural Habitats Jack D. Doyle, Guenther Stotzky, Gwendolyn McClung, and Charles W. Hendricks Detection, Isolation, and Stability of Megaplasmid-Encoded Chloroaromatic Herbicide-Degrading Genes within Pseudomonas Species Douglas J. Cork and Amjad Khalil

Improving Productivity of Heterologous Proteins in Recombinant Saccharomyces cerevisiae Fermentations Amit Vasavada Manipulations of Catabolic Genes for the Degradation and Detoxification of Xenobiotics Rup Lal, Sukanya Lal, P. S. Dhanaraj, and D. M. Saxena Aqueous Two-Phase Extraction for Downstream Processing of Enzymes/Proteins K. S. M. S. Raghava Rao, N. K. Rastogi, M. K. Gowthaman, and N. G. Karanth Biotechnological Potentials of Anoxygenic Phototrophic Bacteria. Part I. Production of Single Cell Protein, Vitamins, Ubiquinones, Hormones, and Enzymes and Use in Waste Treatment Ch. Sasikala and Ch. V. Ramana Biotechnological Potentials of Anoxygenic Phototrophic Bacteria. Part II. Biopolyesters, Biopesticide, Biofuel, and Biofertilizer Ch. Sasikala and Ch. V. Ramana Index

Volume 42

Volume 41

The Insecticidal Proteins of Bacillus thuringiensis P. Ananda Kumar, R. P. Sharma, and V. S. Malik

Microbial Oxidation of Unsaturated Fatty Acids Ching T. Hou

Microbiological Production of Lactic Acid John H. Litchfield

Index

121

122

Contents of Previous Volumes

Biodegradable Polyesters Ch. Sasikala The Utility of Strains of Morphological Group II Bacillus Samuel Singer

Investigation of the Carbon- and Sulfur-Oxidizing Capabilities of Microorganisms by Active-Site Modeling Herbert L. Holland

Phytase Rudy J. Wodzinski and A. H. J. Ullah

Microbial Synthesis of D-Ribose: Metabolic Deregulation and Fermentation Process P. de Wulf and E. J. Vandamme

Index

Production and Application of Tannin Acyl Hydrolase: State of the Art P. K. Lekha and B. K. Lonsane

Volume 43 Production of Acetic Acid by Clostridium thermoaceticum Munir Cheryan, Sarad Parekh, Minish Shah, and Kusuma Witjitra Contact Lenses, Disinfectants, and Acanthamoeba Keratitis Donald G. Ahearn and Manal M. Gabriel Marine Microorganisms as a Source of New Natural Products V. S. Bernan, M. Greenstein, and W. M. Maiese Stereoselective Biotransformations in Synthesis of Some Pharmaceutical Intermediates Ramesh N. Patel Microbial Xylanolytic Enzyme System: Properties and Applications Pratima Bajpai Oleaginous Microorganisms: An Assessment of the Potential Jacek Leman Index

Volume 44 Biologically Active Fungal Metabolites Cedric Pearce Old and New Synthetic Capacities of Baker’s Yeast P. D’Arrigo, G. Pedrocchi-Fantoni, and S. Servi

Ethanol Production from Agricultural Biomass Substrates Rodney J. Bothast and Badal C. Saha Thermal Processing of Foods, A Retrospective, Part I: Uncertainties in Thermal Processing and Statistical Analysis M. N. Ramesh, S. G. Prapulla, M. A. Kumar, and M. Mahadevaiah Thermal Processing of Foods, A Retrospective, Part II: On-Line Methods for Ensuring Commercial Sterility M. N. Ramesh, M. A. Kumar, S. G. Prapulla, and M. Mahadevaiah Index

Volume 45 One Gene to Whole Pathway: The Role of Norsolorinic Acid in Aflatoxin Research J. W. Bennett, P.-K. Chang, and D. Bhatnagar Formation of Flavor Compounds in Cheese P. F. Fox and J. M. Wallace The Role of Microorganisms in Soy Sauce Production Desmond K. O’Toole Gene Transfer Among Bacteria in Natural Environments Xiaoming Yin and G. Stotzky


Breathing Manganese and Iron: Solid-State Respiration Kenneth H. Nealson and Brenda Little

Microbial Production of Oligosaccharides: A Review S. G. Prapulla, V. Subhaprada, and N. G. Karanth

Enzymatic Deinking Pratima Bajpai

Index

Microbial Production of Docosahexaenoic Acid (DHA, C22:6) Ajay Singh and Owen P. Word Index

Volume 46 Cumulative Subject Index

Volume 47 Seeing Red: The Story of Prodigiosin J. W. Bennett and Ronald Bentley Microbial/Enzymatic Synthesis of Chiral Drug Intermediates Ramesh N. Patel Recent Developments in the Molecular Genetics of the Erythromycin-Producing Organism Saccharopolyspora erythraea Thomas J. Vanden Boom Bioactive Products from Streptomyces Vladisalv Behal Advances in Phytase Research Edward J. Mullaney, Catherine B. Daly, and Abdul H. J. Ullah Biotransformation of Unsaturated Fatty Acids of industrial Products Ching T. Hou Ethanol and Thermotolerance in the Bioconversion of Xylose by Yeasts Thomas W. Jeffries and Yong-Su Jin Microbial Degradation of the Pesticide Lindane (g-Hexachlorocyclohexane) Brajesh Kumar Singh, Ramesh Chander Kuhad, Ajay Singh, K. K. Tripathi, and P. K. Ghosh

123

Volume 48 Biodegredation of Nitro-Substituted Explosives by White-Rot Fungi: A Mechanistic Approach Benoit Van Aken and Spiros N. Agathos Microbial Degredation of Pollutants in Pulp Mill Effluents Pratima Bajpai Bioremediation Technologies for Metal-Containing Wastewaters Using Metabolically Active Microorganisms Thomas Pumpel and Kishorel M. Paknikar The Role of Microorganisms in Ecological Risk Assessment of Hydrophobic Organic Contaminants in Soils C. J. A. MacLeod, A. W. J. Morriss, and K. T. Semple The Development of Fungi: A New Concept Introduced By Anton de Bary Gerhart Drews Bartolomeo Gosio, 1863–1944: An Appreciation Ronald Bentley Index

Volume 49 Biodegredation of Explosives Susan J. Rosser, Amrik Basran, Emmal R. Travis, Christopher E. French, and Neil C. Bruce Biodiversity of Acidophilic Prokaryotes Kevin B. Hallberg and D. Barrie Johnson

124


Laboratory Birproduction of Paralytic Shellfish Toxins in Dinoflagellates Dennis P. H. Hsieh, Dazhi Wang, and Garry H. Chang Metal Toxicity in Yeasts and the Role of Oxidative Stress S. V. Avery Foodbourne Microbial Pathogens and the Food Research Institute M. Ellin Doyle and Michael W. Pariza Alexander Flemin and the Discovery of Penicillin J. W. Bennett and King-Thom Chung Index

Volume 50 Paleobiology of the Archean Sherry L. Cady A Comparative Genomics Approach for Studying Ancestral Proteins and Evolution Ping Liang and Monica Riley Chromosome Packaging by Archaeal Histones Kathleen Sandman and John N. Reeve DNA Recombination and Repair in the Archaea Erica M. Seitz, Cynthia A. Haseltine, and Stephen C. Kowalczykowski Basal and Regulated Transcription in Archaea Jo¨rg Soppa Protein Folding and Molecular Chaperones in Archaea Michel R. Leroux Archaeal Proteasomes: Proteolytic Nanocompartments of the Cell Julie A. Maupin-Furlow, Steven J. Kaczowka, Mark S. Ou, and Heather L. Wilson Archaeal Catabolite Repression: A Gene Regulatory Paradigm Elisabetta Bini and Paul Blum Index

Volume 51 The Biochemistry and Molecular Biology of Lipid Accumulation in Oleaginous Microorganisms Colin Ratledge and James P. Wynn Bioethanol Technology: Developments and Perspectives Owen P. Ward and Ajay Singh Progress of Aspergillus oryzae Genomics Masayuki Machida Transmission Genetics of Microbotryum violaceum (Ustilago violacea): A Case History E. D. Garber and M. Ruddat Molecular Biology of the Koji Molds Katsuhiko Kitamoto Noninvasive Methods for the Investigation of Organisms at Low Oxygen Levels David Lloyd The Development of the Penicillin Production Process in Delft, The Netherlands, During World War II Under Nazi Occupation Marlene Burns and Piet W. M. van Dijck Genomics for Applied Microbiology William C. Nierman and Karen E. Nelson Index

Volume 52 Soil-Based Gene Discovery: A New Technology to Accelerate and Broaden Biocatalytic Applications Kevin A. Gray, Toby H. Richardson, Dan E. Robertson, Paul E. Swanson, and Mani V. Subramanian The Potential of Site-Specific Recombinases as Novel Reporters in Whole-Cell Biosensors of Pollution Paul Hinde, Jane Meadows, Jon Saunders, and Clive Edwards


Microbial Phosphate Removal and Polyphosphate Production from Wastewaters John W. McGrath and John P. Quinn Biosurfactants: Evolution and Diversity in Bacteria Raina M. Maier Comparative Biology of Mesophilic and Thermophilic Nitrile Hydratases Don A. Cowan, Rory A. Cameron, and Tsepo L. Tsekoa From Enzyme Adaptation to Gene Regulation William C. Summers Acid Resistance in Escherichia coli Hope T. Richard and John W. Foster Iron Chelation in Chemotherapy Eugene D. Weinberg Angular Leaf Spot: A Disease Caused by the Fungus Phaeoisariopsis griseola (Sacc.) Ferraris on Phaseolus vulgaris L. Sebastian Stenglein, L. Daniel Ploper, Oscar Vizgarra, and Pedro Balatti The Fungal Genetics Stock Center: From Molds to Molecules Kevin McCluskey Adaptation by Phase Variation in Pathogenic Bacteria Laurence Salau¨n, Lori A. S. Snyder, and Nigel J. Saunders What Is an Antibiotic? Revisited Ronald Bentley and J. W. Bennett An Alternative View of the Early History of Microbiology Milton Wainwright The Delft School of Microbiology, from the Nineteenth to the Twenty-first Century Lesley A. Robertson

125

Anaerobic Dehalogenation of Organohalide Contaminants in the Marine Environment Max M. Ha¨ggblom, Young-Boem Ahn, Donna E. Fennell, Lee J. Kerkhof, and Sung-Keun Rhee Biotechnological Application of Metal-Reducing Microorganisms Jonathan R. Lloyd, Derek R. Lovley, and Lynne E. Macaskie Determinants of Freeze Tolerance in Microorganisms, Physiological Importance, and Biotechnological Applications An Tanghe, Patrick Van Dijck, and Johan M. Thevelein Fungal Osmotolerance P. Hooley, D. A. Fincham, M. P. Whitehead, and N. J. W. Clipson Mycotoxin Research in South Africa M. F. Dutton Electrophoretic Karyotype Analysis in Fungi J. Beadle, M. Wright, L. McNeely, and J. W. Bennett Tissue Infection and Site-Specific Gene Expression in Candida albicans Chantal Fradin and Bernard Hube LuxS and Autoinducer-2: Their Contribution to Quorum Sensing and Metabolism in Bacteria Klaus Winzer, Kim R. Hardie, and Paul Williams Microbiological Contributions to the Search of Extraterrestrial Life Brendlyn D. Faison Index

Volume 54

Volume 53

Metarhizium spp.: Cosmopolitan InsectPathogenic Fungi – Mycological Aspects Donald W. Roberts and Raymond J. St. Leger

Biodegradation of Organic Pollutants in the Rhizosphere Liz J. Shaw and Richard G. Burns

Molecular Biology of the Burkholderia cepacia Complex Jimmy S. H. Tsang

Index

126


Non-Culturable Bacteria in Complex Commensal Populations William G. Wade l Red-Mediated Genetic Manipulation of Antibiotic-Producing Streptomyces Bertolt Gust, Govind Chandra, Dagmara Jakimowicz, Tian Yuqing, Celia J. Bruton, and Keith F. Chater Colicins and Microcins: The Next Generation Antimicrobials Osnat Gillor, Benjamin C. Kirkup, and Margaret A. Riley Mannose-Binding Quinone Glycoside, MBQ: Potential Utility and Action Mechanism Yasuhiro Igarashi and Toshikazu Oki Protozoan Grazing of Freshwater Biofilms Jacqueline Dawn Parry Metals in Yeast Fermentation Processes Graeme M. Walker Interactions between Lactobacilli and Antibiotic-Associated Diarrhea Paul Naaber and Marika Mikelsaar Bacterial Diversity in the Human Gut Sandra MacFarlane and George T. MacFarlane Interpreting the Host-Pathogen Dialogue Through Microarrays Brian K. Coombes, Philip R. Hardwidge, and B. Brett Finlay The Inactivation of Microbes by Sunlight: Solar Disinfection as a Water Treatment Process Robert H. Reed Index

Volume 55 Fungi and the Indoor Environment: Their Impact on Human Health

J. D. Cooley, W. C. Wong, C. A. Jumper, and D. C. Straus Fungal Contamination as a Major Contributor to Sick Building Syndrome De-Wei LI and Chin S. Yang Indoor Moulds and Their Associations with Air Distribution Systems Donald G. Ahearn, Daniel L. Price, Robert Simmons, Judith Noble-Wang, and Sidney A. Crow, Jr. Microbial Cell Wall Agents and Sick Building Syndrome Ragnar Rylander The Role of Stachybotrys in the Phenomenon Known as Sick Building Syndrome Eeva-Liisa Hintikka Moisture-Problem Buildings with Molds Causing Work-Related Diseases Kari Reijula Possible Role of Fungal Hemolysins in Sick Building Syndrome Stephen J. Vesper and Mary Jo Vesper The Roles of Penicillium and Aspergillus in Sick Building Syndrome (SBS) Christopher J. Schwab and David C. Straus Pulmonary Effects of Stachybotrys chartarum in Animal Studies Iwona Yike and Dorr G. Dearborn Toxic Mold Syndrome Michael B. Levy and Jordan N. Fink Fungal Hypersensitivity: Pathophysiology, Diagnosis, Therapy Vincent A. Marinkovich Indoor Molds and Asthma in Adults Maritta S. Jaakkola and Jouni J. K. Jaakkola Role of Molds and Mycotoxins in Being Sick in Buildings: Neurobehavioral and Pulmonary Impairment Kaye H. Kilburn


The Diagnosis of Cognitive Impairment Associated with Exposure to Mold Wayne A. Gordon and Joshua B. Cantor Mold and Mycotoxins: Effects on the Neurological and Immune Systems in Humans Andrew W. Campbell, Jack D. Thrasher, Michael R. Gray, and Aristo Vojdani Identification, Remediation, and Monitoring Processes Used in a Mold-Contaminated High School S. C. Wilson, W. H. Holder, K. V. Easterwood, G. D. Hubbard, R. F. Johnson, J. D. Cooley, and D. C. Straus The Microbial Status and Remediation of Contents in Mold-Contaminated Structures Stephen C. Wilson and Robert C. Layton Specific Detection of Fungi Associated With SBS When Using Quantitative Polymerase Chain Reaction Patricia Cruz and Linda D. Stetzenbach Index

Volume 56 Potential and Opportunities for Use of Recombinant Lactic Acid Bacteria in Human Health Sean Hanniffy, Ursula Wiedermann, Andreas Repa, Annick Mercenier, Catherine Daniel, Jean Fioramonti, Helena Tlaskolova, Hana Kozakova, Hans Israelsen, Sren Madsen, Astrid Vrang, Pascal Hols, Jean Delcour, Peter Bron, Michiel Kleerebezem, and Jerry Wells Novel Aspects of Signaling in Streptomyces Development Gilles P. van Wezel and Erik Vijgenboom Polysaccharide Breakdown by Anaerobic Microorganisms Inhabiting the Mammalian Gut Harry J. Flint Lincosamides: Chemical Structure, Biosynthesis, Mechanism of Action, Resistance, and Applications

127

Jaroslav Spı´zˇek, Jitka Novotna´, and Toma´sˇ Rˇezanka Ribosome Engineering and Secondary Metabolite Production Kozo Ochi, Susumu Okamoto, Yuzuru Tozawa, Takashi Inaoka, Takeshi Hosaka, Jun Xu, and Kazuhiko Kurosawa Developments in Microbial Methods for the Treatment of Dye Effluents R. C. Kuhad, N. Sood, K. K. Tripathi, A. Singh, and O. P. Ward Extracellular Glycosyl Hydrolases from Clostridia Wolfgang H. Schwarz, Vladimir V. Zverlov, and Hubert Bahl Kernel Knowledge: Smut of Corn Marıá D. Garcıá-Pedrajas and Scott E. Gold Bacterial ACC Deaminase and the Alleviation of Plant Stress Bernard R. Glick Uses of Trichoderma spp. to Alleviate or Remediate Soil and Water Pollution G. E. Harman, M. Lorito, and J. M. Lynch Bacteriophage Defense Systems and Strategies for Lactic Acid Bacteria Joseph M. Sturino and Todd R. Klaenhammer Current Issues in Genetic Toxicology Testing for Microbiologists Kristien Mortelmans and Doppalapudi S. Rupa Index

Volume 57 Microbial Transformations of Mercury: Potentials, Challenges, and Achievements in Controlling Mercury Toxicity in the Environment Tamar Barkay and Irene Wagner-Do¨bler

128


Interactions Between Nematodes and Microorganisms: Bridging Ecological and Molecular Approaches Keith G. Davies Biofilm Development in Bacteria Katharine Kierek-Pearson and Ece Karatan Microbial Biogeochemistry of Uranium Mill Tailings Edward R. Landa Yeast Modulation of Wine Flavor Jan H. Swiegers and Isak S. Pretorius Moving Toward a Systems Biology Approach to the Study of Fungal Pathogenesis in the Rice Blast Fungus Magnaporthe grisea Claire Veneault-Fourrey and Nicholas J. Talbot

Richard ffrench-Constant and Nicholas Waterfield Engineering Antibodies for Biosensor Technologies Sarah Goodchild, Tracey Love, Neal Hopkins, and Carl Mayers Molecular Characterization of Ochratoxin A Biosynthesis and Producing Fungi J. O’Callaghan and A. D. W. Dobson Index

Volume 59 Biodegradation by Members of the Genus Rhodococcus: Biochemistry, Physiology, and Genetic Adaptation Michael J. Larkin, Leonid A. Kulakov, and Christopher C. R. Allen

The Biotrophic Stages of Oomycete–Plant Interactions Laura J. Grenville-Briggs and Pieter van West

Genomes as Resources for Biocatalysis Jon D. Stewart

Contribution of Nanosized Bacteria to the Total Biomass and Activity of a Soil Microbial Community Nicolai S. Panikov

Process and Catalyst Design Objectives for Specific Redox Biocatalysis Daniel Meyer, Bruno Bu¨hler, and Andreas Schmid

Index

Volume 58 Physiology and Biotechnology of Aspergillus O. P. Ward, W. M. Qin, J. Dhanjoon, J. Ye, and A. Singh Conjugative Gene Transfer in the Gastrointestinal Environment Tine Rask Licht and Andrea Wilcks Force Measurements Between a Bacterium and Another Surface In Situ Ruchirej Yongsunthon and Steven K. Lower

The Biosynthesis of Polyketide Metabolites by Dinoflagellates Kathleen S. Rein and Richard V. Snyder Biological Halogenation has Moved far Beyond Haloperoxidases Karl-Heinz van Peé, Changjiang Dong, Silvana Flecks, Jim Naismith, Eugenio P. Patallo, and Tobias Wage Phage for Rapid Detection and Control of Bacterial Pathogens in Food Catherine E. D. Rees and Christine E. R. Dodd Gastrointestinal Microflora: Probiotics S. Kolida, D. M. Saulnier, and G. R. Gibson

Actinomycetes and Lignin Degradation Ralph Kirby

The Role of Helen Purdy Beale in the Early Development of Plant Serology and Virology Karen-Beth G. Scholthof and Paul D. Peterson

An ABC Guide to the Bacterial Toxin Complexes

Index


Volume 60 Microbial Biocatalytic Processes and Their Development John M. Woodley Occurrence and Biocatalytic Potential of Carbohydrate Oxidases Erik W. van Hellemond, Nicole G. H. Leferink, Dominic P. H. M. Heuts, Marco W. Fraaije, and Willem J. H. van Berkel Microbial Interactions with Humic Substances J. Ian Van Trump, Yvonne Sun, and John D. Coates Significance of Microbial Interactions in the Mycorrhizosphere Gary D. Bending, Thomas J. Aspray, and John M. Whipps Escherich and Escherichia Herbert C. Friedmann Index

Volume 61 Unusual Two-Component Signal Transduction Pathways in the Actinobacteria Matthew I. Hutchings Acyl-HSL Signal Decay: Intrinsic to Bacterial Cell–Cell Communications Ya-Juan Wang, Jean Jing Huang, and Jared Renton Leadbetter Microbial Exoenzyme Production in Food Peggy G. Braun Biogenetic Diversity of Cyanobacterial Metabolites Ryan M. Van Wagoner, Allison K. Drummond, and Jeffrey L. C. Wright Pathways to Discovering New Microbial Metabolism for Functional Genomics and Biotechnology Lawrence P. Wackett

129

Biocatalysis by Dehalogenating Enzymes Dick B. Janssen Lipases from Extremophiles and Potential for Industrial Applications Moh’d Salameh and Juergen Wiegel In Situ Bioremediation Kirsten S. Jrgensen Bacterial Cycling of Methyl Halides Hendrik Scha¨fer, Laurence G. Miller, Ronald S. Oremland, and J. Colin Murrell Index

Volume 62 Anaerobic Biodegradation of Methyl tert-Butyl Ether (MTBE) and Related Fuel Oxygenates Max M. Ha¨ggblom, Laura K. G. Youngster, Piyapawn Somsamak, and Hans H. Richnow Controlled Biomineralization by and Applications of Magnetotactic Bacteria Dennis A. Bazylinski and Sabrina Schu¨bbe The Distribution and Diversity of Euryarchaeota in Termite Guts Kevin J. Purdy Understanding Microbially Active Biogeochemical Environments Deirdre Gleeson, Frank McDermott, and Nicholas Clipson The Scale-Up of Microbial Batch and Fed-Batch Fermentation Processes Christopher J. Hewitt and Alvin W. Neinow Production of Recombinant Proteins in Bacillus subtilis Wolfgang Schumann

130


Quorum Sensing: Fact, Fiction, and Everything in Between Yevgeniy Turovskiy, Dimitri Kashtanov, Boris Paskhover, and Michael L. Chikindas Rhizobacteria and Plant Sulfur Supply Michael A. Kertesz, Emma Fellows, and Achim Schmalenberger Antibiotics and Resistance Genes: Influencing the Microbial Ecosystem in the Gut Katarzyna A. Kazimierczak and Karen P. Scott Index

Volume 63 A Ferment of Fermentations: Reflections on the Production of Commodity Chemicals Using Microorganisms Ronald Bentley and Joan W. Bennett Submerged Culture Fermentation of ‘‘Higher Fungi’’: The Macrofungi Mariana L. Fazenda, Robert Seviour, Brian McNeil, and Linda M. Harvey Bioprocessing Using Novel Cell Culture Systems Sarad Parekh, Venkatesh Srinivasan, and Michael Horn Nanotechnology in the Detection and Control of Microorganisms Pengju G. Luo and Fred J. Stutzenberger Metabolic Aspects of Aerobic Obligate Methanotrophy Yuri A. Trotsenko and John Colin Murrell Bacterial Efflux Transport in Biotechnology Tina K. Van Dyk Antibiotic Resistance in the Environment, with Particular Reference to MRSA William Gaze, Colette O’Neill, Elizabeth Wellington, and Peter Hawkey Host Defense Peptides in the Oral Cavity Deirdre A. Devine and Celine Cosseau Index

Volume 64 Diversity of Microbial Toluene Degradation Pathways R. E. Parales, J. V. Parales, D. A. Pelletier, and J. L. Ditty Microbial Endocrinology: Experimental Design Issues in the Study of Interkingdom Signalling in Infectious Disease Primrose P. E. Freestone and Mark Lyte Molecular Genetics of Selenate Reduction by Enterobacter cloacae SLD1a-1 Nathan Yee and Donald Y. Kobayashi Metagenomics of Dental Biofilms Peter Mullany, Stephanie Hunter, and Elaine Allan Biosensors for Ligand Detection Alison K. East, Tim H. Mauchline, and Philip S. Poole Islands Shaping Thought in Microbial Ecology Christopher J. van der Gast Human Pathogens and the Phyllosphere John M. Whipps, Paul Hand, David A. C. Pink, and Gary D. Bending Microbial Retention on Open Food Contact Surfaces and Implications for Food Contamination Joanna Verran, Paul Airey, Adele Packer, and Kathryn A. Whitehead Index

Volume 65 Capsular Polysaccharides in Escherichia coli David Corbett and Ian S. Roberts Microbial PAH Degradation Evelyn Doyle, Lorraine Muckian, Anne Marie Hickey, and Nicholas Clipson Acid Stress Responses in Listeria monocytogenes Sheila Ryan, Colin Hill, and Cormac G. M. Gahan


Global Regulators of Transcription in Escherichia coli: Mechanisms of Action and Methods for Study David C. Grainger and Stephen J. W. Busby The Role of Sigma B (sB) in the Stress Adaptations of Listeria monocytogenes: Overlaps Between Stress Adaptation and Virulence Conor P. O’ Byrne and Kimon A. G. Karatzas Protein Secretion and Membrane Insertion Systems in Bacteria and Eukaryotic Organelles Milton H. Saier, Chin Hong Ma, Loren Rodgers, Dorjee G. Tamang, and Ming Ren Yen Metabolic Behavior of Bacterial Biological Control Agents in Soil and Plant Rhizospheres Cynthia A. Pielach, Daniel P. Roberts, and Donald Y. Kobayashi Copper Homeostasis in Bacteria Deenah Osman and Jennifer S. Cavet Pathogen Surveillance Through Monitoring of Sewer Systems Ryan G. Sinclair, Christopher Y. Choi, Mark R. Riley, and Charles P. Gerba Index

131

Cutinases: Properties and Industrial Applications Tatiana Fontes Pio and Gabriela Alves Macedo Microbial Deterioration of Stone Monuments—An Updated Overview Stefanie Scheerer, Otto Ortega-Morales, and Christine Gaylarde Microbial Processes in Oil Fields: Culprits, Problems, and Opportunities Noha Youssef, Mostafa S. Elshahed, and Michael J. McInerney Index

Volume 67 Phage Evolution and Ecology Stephen T. Abedon Nucleoid-Associated Proteins and Bacterial Physiology Charles J. Dorman Biodegradation of Pharmaceutical and Personal Care Products Jeanne Kagle, Abigail W. Porter, Robert W. Murdoch, Giomar Rivera-Cancel, and Anthony G. Hay Bioremediation of Cyanotoxins Christine Edwards and Linda A. Lawton Virulence in Cryptococcus Species Hansong Ma and Robin C. May

Volume 66 Multiple Effector Mechanisms Induced by Recombinant Listeria monocytogenes Anticancer Immunotherapeutics Anu Wallecha, Kyla Driscoll Carroll, Paulo Cesar Maciag, Sandra Rivera, Vafa Shahabi, and Yvonne Paterson Diagnosis of Clinically Relevant Fungi in Medicine and Veterinary Sciences Olivier Sparagano and Sam Foggett Diversity in Bacterial Chemotactic Responses and Niche Adaptation Lance D. Miller, Matthew H. Russell, and Gladys Alexandre

Molecular Networks in the Fungal Pathogen Candida albicans Rebecca A. Hall, Fabien Cottier, and Fritz A. Mu¨hlschlegel Temperature Sensors of Eubacteria Wolfgang Schumann Deciphering Bacterial Flagellar Gene Regulatory Networks in the Genomic Era Todd G. Smith and Timothy R. Hoover Genetic Tools to Study Gene Expression During Bacterial Pathogen Infection Ansel Hsiao and Jun Zhu Index

132


Volume 68 Bacterial L-Forms E. J. Allan, C. Hoischen, and J. Gumpert Biochemistry, Physiology and Biotechnology of Sulfate-Reducing Bacteria Larry L. Barton and Guy D. Fauque Biotechnological Applications of Recombinant Microbial Prolidases Casey M. Theriot, Sherry R. Tove, and Amy M. Grunden The Capsule of the Fungal Pathogen Cryptococcus neoformans Oscar Zaragoza, Marcio L. Rodrigues, Magdia De Jesus, Susana Frases, Ekaterina Dadachova, and Arturo Casadevall Baculovirus Interactions In Vitro and In Vivo Xiao-Wen Cheng and Dwight E. Lynn Posttranscriptional Gene Regulation in Kaposi’s Sarcoma-Associated Herpesvirus Nicholas K. Conrad Index

Volume 69 Variation in Form and Function: The Helix-Turn-Helix Regulators of the GntR Superfamily Paul A. Hoskisson and Se´bastien Rigali Biogenesis of the Cell Wall and Other Glycoconjugates of Mycobacterium tuberculosis Devinder Kaur, Marcelo E. Guerin, Henrieta Sˇkovierova´, Patrick J. Brennan, and Mary Jackson Antimicrobial Properties of Hydroxyxanthenes Joy G. Waite and Ahmed E. Yousef In Vitro Biofilm Models: An Overview Andrew J. McBain

Zones of Inhibition? The Transfer of Information Relating to Penicillin in Europe during World War II Gilbert Shama The Genomes of Lager Yeasts Ursula Bond Index

Volume 70 Thermostable Enzymes as Biocatalysts in the Biofuel Industry Carl J. Yeoman, Yejun Han, Dylan Dodd, Charles M. Schroeder, Roderick I. Mackie, and Isaac K. O. Cann Production of Biofuels from Synthesis Gas Using Microbial Catalysts Oscar Tirado-Acevedo, Mari S. Chinn, and Amy M. Grunden Microbial Naphthenic Acid Degradation Corinne Whitby Surface and Adhesion Properties of Lactobacilli G. Deepika and D. Charalampopoulos Shining Light on the Microbial World: The Application of Raman Microspectroscopy Wei E. Huang, Mengqiu Li, Roger M. Jarvis, Royston Goodacre, and Steven A. Banwart Detection of Invasive Aspergillosis Christopher R. Thornton Bacteriophage Host Range and Bacterial Resistance Paul Hyman and Stephen T. Abedon Index

Volume 71 Influence of Escherichia coli Shiga Toxin on the Mammalian Central Nervous System Fumiko Obata


Natural Products for Type II Diabetes Treatment Amruta Bedekar, Karan Shah, and Mattheos Koffas Experimental Models Used to Study Human Tuberculosis Ronan O’Toole Biosynthesis of Peptide Signals in Gram-Positive Bacteria Matthew Thoendel and Alexander R. Horswill Cell Immobilization for Production of Lactic Acid: Biofilms Do It Naturally Suzanne F. Dagher, Alicia L. Ragout, Faustino Sin˜eriz, and Jose´ M. Bruno-Ba´rcena Microbial Fingerprinting using Matrix-Assisted Laser Desorption Ionization Time-Of-Flight Mass Spectrometry (MALDI-TOF MS): Applications and Challenges R. Giebel, C. Worden, S. M. Rust, G. T. Kleinheinz, M. Robbins, and T. R. Sandrin

Index

Volume 72 Evolution of the Probiotic Concept: From Conception to Validation and Acceptance in Medical Science Walter J. Dobrogosz, Trent J. Peacock, and Hosni M. Hassan Prokaryotic and Eukaryotic Diversity of the Human Gut Julian R. Marchesi

133

Oxalate-Degrading Bacteria of the Human Gut as Probiotics in the Management of Kidney Stone Disease Valerie R. Abratt and Sharon J. Reid Morphology and Rheology in Filamentous Cultivations T. Wucherpfennig, K. A. Kiep, H. Driouch, C. Wittmann, and R. Krull Methanogenic Degradation of Petroleum Hydrocarbons in Subsurface Environments: Remediation, Heavy Oil Formation, and Energy Recovery N. D. Gray, A. Sherry, C. Hubert, J. Dolfing, and I. M. Head Index

Volume 73 Heterologous Protein Secretion by Bacillus Species: From the Cradle to the Grave Susanne Pohl and Colin R. Harwood Function of Protein Phosphatase-1, Glc7, in Saccharomyces cerevisiae John F. Cannon Milliliter-Scale Stirred Tank Reactors for the Cultivation of Microorganisms Ralf Hortsch and Dirk Weuster-Botz Type I Interferon Modulates the Battle of Host Immune System Against Viruses Young-Jin Seo and Bumsuk Hahm Index

Advances in Applied Microbiology, Volume 74

Advances in applied microbiology

Advances in Applied Microbiology, Volume 26

Advances in Applied Microbiology, Volume 42

Advances in Applied Microbiology, Volume 37

Advances in Applied Microbiology, Volume 49

Advances in Applied Microbiology, Volume 73

Advances in Applied Microbiology, Volume 14

Advances in Applied Microbiology, Volume 60

Advances in Applied Microbiology, Volume 45

Advances in Applied Microbiology, Volume 10

Advances in Applied Microbiology, Volume 17

Advances in Applied Microbiology, Volume 9

Advances in Applied Microbiology, Volume 18

Advances in Applied Microbiology, Volume 60

Advances in Applied Microbiology, Volume 39

Advances in Applied Microbiology, Volume 11

Advances in Applied Microbiology, Volume 49

Advances in Applied Microbiology, Volume 68

Advances in Applied Microbiology, Volume 56

Advances in Applied Microbiology, Volume 6

Advances in Applied Microbiology, Volume 36

Advances in Applied Microbiology, Volume 72

Advances in Applied Microbiology, Volume 12

Advances in Applied Microbiology, Volume 63

Advances in Applied Microbiology, Volume 47

Advances in Applied Microbiology, Volume 75

Advances in Applied Microbiology, Volume 8

Advances in Applied Microbiology, Volume 65

Advances in Applied Microbiology, Volume 48

Advances in Applied Microbiology, Volume 43

Advances in Applied Microbiology, Volume 74