ADVISORY BOARD DAVID BALTIMORE ROBERT M. CHANOCK PETER C. DOHERTY H. J. GROSS B. D. HARRISON BERNARD MOSS ERLING NORRBY J. J. SKEHEL M. H. V. VAN REGENMORTEL
Academic Press is an imprint of Elsevier 32 Jamestown Road, London, NW1 7BY, UK Radarweg 29, PO Box 211, 1000 AE Amsterdam, The Netherlands 30 Corporate Drive, Suite 400, Burlington, MA 01803, USA 525 B Street, Suite 1900, San Diego, CA 92101-4495, USA First edition 2008 Copyright # 2008 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: (þ44) (0) 1865 843830, fax: (þ44) (0) 1865 853333; e-mail:
[email protected]. Alternatively you can submit your request online by visiting the Elsevier web site at http://www.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 Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-0-12-374322-0 ISSN: 0065-3527 For information on all Academic Press publications visit our website at elsevierdirect.com Printed and bound in USA 08 09 10 11 12 10 9 8 7 6 5 4 3 2 1
CHAPTER
1 The History and Evolution of Human Dengue Emergence Nikos Vasilakis and Scott C. Weaver
Contents
Abstract
I. Introduction to Flaviviruses II. Dengue Viruses A. Classification of dengue viruses III. Dengue Epidemiology A. History of dengue virus as a human pathogen B. DENV transmission cycles IV. Evolution A. DENV evolutionary relationships—origins and emergence B. Rates of DENV evolution C. Evolution of virulence V. Potential for Sylvatic Denv Reemergence A. Epidemics and human contact B. The influence of natural immunity or vaccination on potential sylvatic DENV emergence C. Selection pressures D. Adaptation for urban transmission E. Conclusions and future work Acknowledgments References
2 5 5 8 8 21 29 29 35 37 42 42 44 47 49 54 55 55
Dengue viruses (DENV) are the most important human arboviral pathogens. Transmission in tropical and subtropical regions of the world includes a sylvatic, enzootic cycle between nonhuman primates and arboreal mosquitoes of the genus Aedes, and an urban,
Center for Tropical Diseases and Department of Pathology, University of Texas Medical Branch, Galveston, Texas 77555-0609 Advances in Virus Research, Volume 72 ISSN 0065-3527, DOI: 10.1016/S0065-3527(08)00401-6
#
2008 Elsevier Inc. All rights reserved.
1
2
Nikos Vasilakis and Scott C. Weaver
endemic/epidemic cycle principally between Aedes aegypti, a mosquito that exploits peridomestic water containers as its larval habitats, and human reservoir hosts that are preferred for blood feeding. Genetic studies suggest that all four serotypes of endemic/epidemic DENV evolved independently from ancestral, sylvatic viruses and subsequently became both ecologically and evolutionarily distinct. The independent evolution of these four serotypes was accompanied by the expansion of the sylvatic progenitors’ host range in Asia to new vectors and hosts, which probably occurred gradually over a period of several hundred years. Although many emerging viral pathogens adapt to human replication and transmission, the available evidence indicates that adaptation to humans is probably not a necessary component of sylvatic DENV emergence. These findings imply that the sylvatic DENV cycles in Asia and West Africa will remain a potential source of re-emergence. Sustained urban vector control programs and/or human vaccination will be required to control DEN because the enzootic vectors and primate reservoir hosts are not amenable to interventions.
I. INTRODUCTION TO FLAVIVIRUSES Dengue viruses (DENV) are members of the genus Flavivirus in the Family Flaviviridae. Flaviviruses are single-stranded RNA viruses of positive polarity and most, but not all, require hematophagous arthropods (mosquitoes or ticks) to complete their horizontal transmission cycle. They are responsible for a broad spectrum of pathogenic manifestations in humans, domestic animals, and birds (Heinz et al., 2000). Flaviviruses are widely distributed nearly throughout the world, except Antarctica. More than 50% of all known flaviviruses have been associated with human disease and include some of the most important human pathogens, such as Yellow fever virus (YFV), DENV, Japanese encephalitis virus ( JEV), and Tick-borne encephalitis virus (TBEV). For example, DENV is responsible for the highest incidence of human morbidity and mortality among all flaviviruses: ca. 100 million infections annually, resulting in approximately 500,000 cases of DEN hemorrhagic fever (DHF) with a case fatality rate of about 5% (Halstead, 1997). The majority of human infections with flaviviruses are asymptomatic, whereas symptomatic infections commonly manifest themselves as a flu-like disease that is characterized by sudden onset of fever, arthralgia, myalgia, retro-orbital headaches, maculopapular rash, leukopenia, vascular leakage, and/or encephalitis (Belov et al., 1995; Burke et al., 1988; Gritsun et al., 2003; Lumsden, 1958; Work et al.,
History and Evolution of Human Dengue Emergence
3
1957). Depending on the flavivirus, the infection may also cause severe encephalitis with lifelong neurologic sequelae (Brinker and Monath, 1980; Charrel et al., 2004), persistent disease (Ravi et al., 1993; Sharma et al., 1991), or even death (McLean and Donohue, 1959; Tsai and Mitchell, 1989; Work et al., 1957). In animals, flavivirus infections occur in a wide range of animals including sheep, cattle, equids, monkeys, muskrats, rodents, bats, birds, and seabirds (Autorino et al., 2002; Clifford et al., 1971; Gritsun et al., 2003; Lanciotti et al., 1999; Lvov et al., 1971; Malkinson and Banet, 2002; Shope, 2003; St George et al., 1977; Swanepoel, 1994; Swanepoel and Coetzer, 1994; Varelas-Wesley and Calisher, 1982). Infection of animals, as in humans, varies from asymptomatic to lethal. The name flaviviruses comes from the Latin word ‘‘flavus,’’ meaning yellow that signifies jaundice, a common sign of infection with the prototypic Yellow fever virus. The genus Flavivirus includes 56 species (Heinz et al., 2000). Yellow fever virus was among the first filterable agents shown to cause human diseases (along with DENV) (Ashburn and Craig, 1907; Reed and Carroll, 1902), the first virus isolated whose transmission involves the mosquito vector Aedes (Stegomyia) aegypti (Reed and Carroll, 1902; Reed et al., 1900), and the first flavivirus to be cultivated in vitro (Lloyd et al., 1936). Viruses in the Flavivirus genus are grouped taxonomically into three groups with regard to their vector association and antigenic relationships: (1) tick-borne, (2) mosquitoborne, and (3) viruses with no known arthropod vector (NKV). Within the tick-borne group are two antigenically distinct clades: mammalian and seabird virus groups (Fig. 1). The mammalian group includes several important human pathogens, such as Kyasanur Forest disease (KFDV), Powassan (POWV), Omsk hemorrhagic fever (OHFV), TBEV, and Louping ill viruses (LIV). The mosquito-borne viruses can be divided into groups that principally use Aedes spp. or Culex spp. mosquito vectors. The DENV belong to the former group (Fig. 1). Although the closest relative of the DENV as depicted in this tree is an African virus, Kedougou virus from Senegal, this grouping is not robust and a large group of mosquito-borne viruses from Africa, Asia, and the New World group with DENV with equivalently poor statistical support. Therefore, the current flavivirus tree is not informative as to the probable origins of the DENV. Several other members of the mosquito-borne group are the causative agents of severe diseases in humans: St. Louis encephalitis (SLEV), West Nile (WNV), Ilheus (ILHV), Zika (ZIKV), Wesselsbron (WESSV), and YFV. Lastly, within the viruses with no known arthropod vector, there are three antigenically distinct groups: Entebbe bat, Yokose, and Sokoluk viruses group phylogenetically with the mosquito-borne clade, while Modoc-virus and Rio Bravo-like viruses are phylogenetically
4
Nikos Vasilakis and Scott C. Weaver
5% amino acid sequence divergence
100
100
100
Cell fusing agent No known vertebrate host Kamiti river Alkhurma Kyasanur forest Tick-borne encephalitis (Neudoerfl) Tick-borne encephalitis (Vasilchenko) Greek goat Turkish sheep Louping III Negishi Mammals Spanish sheep Tick-borne encephalitis (Sofjin) Tick-borne Langat Powassan (deer tick) Powassan Royalfarm Karshi Gadgets gully Kadam Saumarez reef Seabirds Meaban Tyuleniy Bussu Naranjal Iguape Aroa Stratford Kokobera Cacipacore Alfuy Murray Valley encephalitis Japanese encephalitis Culex spp. Usutu Koutango vectors West Nile (Kunjin) West Nile Yaounde Ntaya Israel Turkey Bagaza Tembusu Mosquito-borne Rocio IIheus St. Louis encephalitis Spondweni Zika Kedougou Dengue-1 Dengue-3 Dengue-2 Dengue-4 Aedes spp. Yellow fever Sepik vectors EdgeHill Bouboui Banzi Uganda S Jugra Saboya Potiskum Yokose Bats Entebbebat Sokuluk Apoi Carey Island Phnom-penh bat BatuCave Bats Dakar RioBravo Montana Myotis leucoencephalitis No known vector Bukalasa Sal Vieja Cowbone Ridge Rodents Modoc San Perlita Jutiapa
FIGURE 1 Phylogenetic tree of the flaviviruses derived from partial NS5 sequences from the GenBank library. Subtypes are written in parentheses after virus names. New World viruses are printed in bold and underlined. The tree was drawn using neighbor joining, and similar topologies were produced using Bayesian methods and maximum parsimony. Numbers indicate bootstrap values for major clades to the right. Reproduced from Hanley & Weaver, 2009, with permission.
distinct from the vector-borne groups. Members of these groups have been isolated mainly from bats or rodents and some have been associated with establishment of persistent infections (Baer and Woodall, 1966; Constantine and Woodall, 1964). A handful of these viruses, such as Dakar bat, Modoc, and Rio Bravo, have been isolated from humans in nature (Karabatsos, 1985b; Shope, 2003), or in several laboratory infections probably due to aerosol transmission (Apoi and Rio Bravo virus) (Karabatsos, 1985a; Sulkin et al., 1962). Most common manifestation of human disease from these viruses is febrile illness and rarely encephalitis (Calisher and Gould, 2003; Shope, 2003).
History and Evolution of Human Dengue Emergence
5
II. DENGUE VIRUSES A. Classification of dengue viruses DENV is considered a species within the genus Flavivirus (family Flaviviridae) and includes four distinct but antigenically related serotypes (DENV-1, -2, -3, and -4) in DEN antigenic complex (Calisher et al., 1989). Of the flaviviruses, DENV are among the most restricted with regard to their natural vertebrate host range, which is believed to include only primates. Currently, all four DENV serotypes can be found in nearly all urban and peri-urban environments throughout the tropics and neotropics where the principal vector, Aedes aegypti, is abundant. This distribution puts at risk of infection nearly a third of the global human population. Initially, DENV of all serotypes were classified genetically into clusters called topotypes using T1 RNase fingerprinting (Repik et al., 1983; Trent et al., 1990). Later, nucleic acid sequencing allowed for the classification of DENV into genetically distinct groups or genotypes within each serotype (Rico-Hesse, 1990). Rico-Hesse defined these ‘genotypes’ as clusters of DENV viruses having nucleotide sequence divergence not greater than 6% within a given genome region (in this case the E/NS1 junction), which was based on the clustering of strains for which associations could be inferred on epidemiological grounds (Rico-Hesse, 1990). Various phylogenetic analyses based on partial E/NS1 or complete E nucleotide sequences indicated that DENV-1 are grouped in five genotypes: (1) genotype I, representing strains from Southeast Asia, China, and East Africa; (2) genotype II, representing strains from Thailand collected in the 1950s and 1960s; (3) genotype III, representing the sylvatic strain collected in Malaysia; (4) genotype IV, representing strains from the West Pacific islands and Australia; and (5) genotype V, representing all strains collected in the Americas, strains from West Africa, and a limited number of strains collected from Asia (Goncalvez et al., 2002; Rico-Hesse, 1990) (Fig. 2). Similar phylogenetic analyses based on E nucleotide sequences indicated that DENV-2 comprise five genotypes: (1) the Asian genotype, consisting of Asian genotype 1 representing strains from Malaysia and Thailand, and Asian genotype 2 representing strains from Vietnam, China, Taiwan, Sri Lanka and the Philippines; (2) the cosmopolitan genotype, representing strains of wide geographic distribution including Australia, East and West Africa, the Pacific and Indian ocean islands, the Indian subcontinent and the Middle East; (3) the American genotype, representing strains from Latin America and older strains collected from the Caribbean, the Indian subcontinent and Pacific Islands in the 1950s and 1960s; (4) the Southeast Asian/American genotype, representing strains from Thailand and Vietnam and strains collected in the Americas over the last 20 years; and (5) the sylvatic genotype, representing strains
6
Nikos Vasilakis and Scott C. Weaver
Hawaii/45
5% nucleotide sequence divergence 98
Thai/0023/81 Thai/0081/82 Thai/K0485/93 Thai/K0109/92 Thai/K0229/90 Thai/K0848/90 Thai/K0379/93 Thai/K0127/94 Thai/0398/89 Thai/K0875/97 Thai/0384/87 Thai/0336/88 Thai/PUO359/80 Thai/K0113/99 Thai/K0088/95 Thai/K0048/97 Thai/K0120/00 Thai/K0107/98 Thai/K0051/99 Chin/99020/99 Chin/060117/06 Chin/06098/06 Myan/Mos30511/01 Myan/Mos30501/01 Myan/4857237/02 Myan/4857228/02 Thai/0005/02 Myan/4354902/01 Myan/3704512/00 Chin/02031/02 Austral/HATI7/83 Hawaii/01Q/01 Hawaii/01P/01 FrPoly/192206/01 Seych/1480/04 Indon/SC01/04 02RBD008 Phil/SA073/02 Phil/01St219/01 Phil/228682/74 Thai/AHF82/82 Carib/9241/81 Carib/1636/77 Phil/8631/84 Phil/1848361/81 Nauru/145A25/74 Thai/2543/63 Reunion/191/04 Reunion/305/04 Arg/297/00 PR/94/94 Peru/DEI0151/91 Col/INS347869/85 Mex/1298/80 Mex/1463/84 Mex/1756/86 Mex/1378/83 Mex/1379/82 Mex/4642/95 Mex/494295 CRico/Cesara1/93 Aruba/4951/85 FrPoly/5103/89 Gren/778156/77 Jam/PRS288690/77 Ven/D5736/95 Ven/D28164/97 Ven/D141602/94 Ven/D125239/94 Ven/D6222/95 Braz/BeH58452697 Per/IQT6152/00 Comoros/D04-329 Thai/0442/80 IC/DakAr1520/85/93 Nig/IBH28328/68 Mal/P72-1244/72
I
97
89
95
IV
Endemic/ epidemic
II
V
96
Sylvatic
FIGURE 2 Phylogenetic relationships of DENV-1. The phylogeny was inferred based on the E nucleotide sequence of 79 DENV-1, using Bayesian analysis (one million reiterations) and all horizontal branches are scaled according to the number of substitutions per site. The scale represents a genetic distance of 0.05 or 5% nucleotide sequence divergence. Bootstrap values are shown for key nodes. Strains are abbreviated as follows: Country abbreviation/strain/year.
collected from humans, forest mosquitoes, or sentinel monkeys in West Africa and Southeast Asia (Lewis et al., 1993; Rico-Hesse et al., 1997; Twiddy et al., 2002; Vasilakis, Tesh, and Weaver, 2008; Wang et al., 2000) (Fig. 3) Important phenotypic differences among the endemic DENV-2 genotypes are discussed below.
History and Evolution of Human Dengue Emergence
5% nucleotide sequence divergence
100
100
100
93
100
Braz/40274/90 Boliv/124B/98 Braz/49255/95 Peru/6663/01 Peru/6658/01 Col/360236/92 Col/360281/92 Ven/Mara3/90 Ven/19966/96 Ven/102954/91 PRico/PR1991A/91 Mex/Oax468/00 Nicar/541/99 Ven/15957/96 Ven/LARD1811/96 Cuba/CUB32/95 Cuba/CUB115/95 Cuba/CUB137/95 Jam/N1409/83 Jam/JAM1983/83 VietN/CTD44/98 Taiw/1897/87 Phil/19406aTw/94 Phil/2088/83 PapuaNG/NGC/44 Mal/M56309/86 Mal/MS8455/87 Thai/THnH-28/93 Thai/THnH-52/93 Thai/THnH-7/93 Myan/0410aTw/04 Thai/ThK0062/00 Thai/ThK0123/00 Mvan/0207aTw/02 Thai/ThK0196/98 Thai/ThK0001/95 Thai/Th0032/88 VietN/0408aTw/04 VietN/0307aTw/03 Thai/Th0194/95 Thai/ThK0010/01 Thai/THnH-p11/93 Thai/Th0015/84 Thai/Puo218/80 Thai/Th0044/77 Thai/Th0066/74 Thai/16681/64 Thai/TH36/56 Som/10/84 BFaso/1349/82 BFaso/190/83 Indo/1051/76 Taiw/TW32/02 Phil/00U18/00 Phil/NCH35/00 Seych/44554/77 Seych/44552/77 SriL/1583/85 SriL/1592/85 SriL/271206/90 Mal/P7-863/69 SriL/206714/89 Sri L/271235/90 India/GWL228/01 Sri L/NIID23/04 Chin/Zhejiang-01/04 Mal/P8-377/69 PRico/PR158/60 PRico/1328/77 Trin/780477/78 Mex/131/92 Mex/328298/95 Mex/132/92 Ven/Ven2/87 Peru/IQT2133/96 Peru/IQT2913/96 Peru/IQT1950/95 India/P9122/57 Mex/200787/83 Trin/1751/53
IC/DakAr578/80 BF/DakAr2039/80 IC/DakArA510.80 BF/DakArA2022/80 IC/DakArA1247/80 Guin/PM33974/81 Sen/DakAr141070/99 Sen/DakAr141069/99 Sen/DakArD75505/91 Sen/DakArHD10674/70 Sen/DakAr20761/74 Niq/IBH11234/66 Niq/IBH11664/66 Niq/IBH11208/66 Mal/P8-1407/75
7
SE Asian/ American
Asian I
Asian genotype Asian II Endemic epidemic
Cosmopolitan
American genotype
Sylvatic
FIGURE 3 Phylogenetic relationships of DENV-2. The phylogeny was inferred based on the E nucleotide sequences in the GenBank library, using Bayesian analysis (one million reiterations) and all horizontal branches are scaled according to the number of substitutions per site. Bayesian probability values are shown for key nodes. Strains are abbreviated as follows: Country abbreviation/strain/year.
Initially DENV-3 were grouped into five genotypes based on T1 RNase fingerprinting analysis (Trent et al., 1990), but subsequent analyses based on prM/E nucleotide (Lanciotti et al., 1994) and later complete genome sequences (Chao et al., 2005) indicate clustering into four genotypes: (1) genotype I, representing strains from Indonesia, Malaysia, the Philippines and recent isolates from the South Pacific islands; (2) genotype II, representing strains from Thailand, Vietnam and Bangladesh; (3) genotype III, representing strains from Sri Lanka, India, Africa and Samoa; however,
8
Nikos Vasilakis and Scott C. Weaver
the complete genome phylogenetic analysis includes the 1962 strain from Thailand within this genotype (Chao et al., 2005); and (4) genotype IV, representing strains from Puerto Rico, Latin and central America and a 1965 Tahiti strain. Sylvatic strains of DENV-3 have not been isolated, but are believed to exist in Malaysia, based on the seroconversion of sentinel monkeys (Rudnick, 1984) (Fig. 4). Lastly, DENV-4 comprise four genotypes based on the E gene (AbuBakar et al., 2002; Foster et al., 2003; Lanciotti et al., 1997) or complete genome sequences (Klungthong et al., 2004): (1) genotype I, representing strains from Thailand, the Philippines, Sri Lanka, and Japan (strains were imported into Japan from Southeast Asia); (2) genotype II, representing strains from Indonesia, Malaysia, Tahiti, the Caribbean and the Americas. Subsequent analysis with additional strains revealed putative evidence of intra-serotypic recombination among DENV-4 from independent lineages (most likely Indonesia 1976 and Malaysia 1969), which may have contributed to the emergence of a distinct genotype, representing all Malaysian strains (AbuBakar et al., 2002). Genotype II has become well established in the Caribbean since its introduction in the area in the early 1980s from Southeast Asia (Bennett et al., 2003; Foster et al., 2003); (3) genotype III, representing recently sampled Thai strains that are distinct from other Thai strains (Klungthong et al., 2004); and (4) genotype IV, representing the sylvatic strains of DENV-4 (Fig. 5).
III. DENGUE EPIDEMIOLOGY A. History of dengue virus as a human pathogen The geographic site of origin of DENV has been a subject of intense speculation; some argue for an African origin due to the same origin of the principal vector, Ae. aegypti (Christophers, 1960; Edwards, 1932). However the ecological and serological work of Smith and Rudnick (Rudnick and Lim, 1986; Smith, 1956), the relative insusceptibility to infection to DENV infection of the ancestral Ae. aegypti formosus from Africa (Diallo et al., 2005) and phylogenetic analyses (Wang et al., 2000) point towards an Asian origin. Regardless of its geographic origin, DENV probably evolved as an arboreal mosquito virus before adapting to lower primates in sylvan environments (Gubler, 1997). The DENV most likely moved out of the forest and into the peridomestic environment at a time congruent with the clearing of the forests and development of human settlements. The earliest known clinical descriptions of a DEN-like illness are found in the Chinese literature during the Chin Dynasty [Common Era (CE) 265–420], Tang Dynasty (CE 610) and Northern Sung Dynasty
9
History and Evolution of Human Dengue Emergence
Tahiti/1327/65 PRico/63 PRico/1339/77
89
84
5% nucleotide sequence divergence
IV
Thai/5987/62 Thai/CH3489/73 Thai/D30033/74 Thai/D30273/80 Thai/D30285M/77 Thai/D30649/80 Thai/D30137/84 Thai/D86007/86 Thai/D30177/81 Thai/D30402/85 Thai/D30220/85 Myan/0508a/05 Bang/Apu/01 Bang/058/00 Bang/0108a/01 Bang/Jacob/01 Thai/D30134/93 Thai/MK315/87 Thai/D30029/90 Thai/D30393/91 Thai/D30396/88 Thai/D30213/88 Thai/LN7029/94 Thai/LN7933/94 Mal/LN6083/95 Thai/D3039694/94 Thai/D30111/02 Thai/D30654/01 Indo/98901640/98 Thai/KPS40657/98 Thai/0211a/02 Thai/D30240/92 Viet/0507a/05 Viet/9809a/98 Thai/D30595/99 Thai/D30650/97 Thai/D30903/98 Phil/87/56 Phil/J1682/83 Phil/168AP2/83 Fiji/29472/92 Tahiti/2167/89 Indon/228761/73 Indon/Sleman/78 Indon/1280/78 Mal/1300/74 Taiw/813KH9408a/94 Phil/0508a/05 Phil/9808a/98 Indo/9108a/91 Indo/9909a/99 ETimor/0153/05 ETimor/0167/05 Indo/0508TW/05 Indo/9804a/98 Indo/TB16/04 Indo/TB55i/04 Moz/1559/85 Samoa/1696/88 SirL/1326/81 India/1416/84 SriL/2783/91 SriL/260698/89 Mex/6584/96 Mex/6883/97 Ven/6668/01 Ven/7984/01 Mex/0AX/00 Nic/24/94 Pan/94 Peru/2812/00 Cuba/21/02 Cuba/580/01 Peru/FSL1212/04 Peru/OBT4024/05 Peru/OBT412/00 OBT1467 Mart/1243/99 Cuba116 Mart/2336/01 Parag/PJ6/06 Parag/YA2/03 Parag/AS10/03 Braz/SG2/02 Braz/ST14/04
II
I
III
FIGURE 4 Phylogenetic relationships of DENV-3. The phylogeny was inferred based on the E nucleotide sequence in the GenBank library, using Bayesian analysis (a million reiterations) and all horizontal branches are scaled according to the number of substitutions per site. Bayesian probability values are shown for key nodes. Strains are abbreviated as follows: Country abbreviation/strain/year.
10
Nikos Vasilakis and Scott C. Weaver
5% nucleotide sequence divergence
93
100
100
100 Mal/P75514/75 Mal/P731120/73 Mal/P752415/73
Indo/1036/76 Indo/1132/77 ElsaI/1411/82 NewCal/5489/81 PRico/M32/82 PRico/M42/86 PRico/6387 PRico/9/87 Bahamas/09160/98 Ven/113/95 Montser/11970/94 Trin/08820/99 PRico/85PR/94 Barb/12102/93 CRica/108CR/96 Surin/9411427/94 Martin/112MQ/95 PRico/41/92 PRico/1650/86 PRico/96/90 PRico/88/94 Tahiti/114094/85 PRico/M33/85 EISal/6494/94 EISal/110/93 Braz/1385/82 Col/371813/96 Ecuad/109/94 Hond/1991/91 Mex/1492/84 PRico/M5/82 PRico/M7/82 PRico/M16/82 PRico/M20/82 Domin/M44/81 Domin/814669/81 Mex/111/95 Mex/5962/96 Mex/6637/96 Jam/830886/83 Jam/8110828/81 Mex/1551/85 Mex/1554/85 Tahiti/S44754/79 Surin/824188/82 Trin/4233/82 Chin/7842/78 Chin/7856/78 Indo/30153/73 Mal/123264/01 Mal/123314/01 Indo/SW36i/84 Indo/SW38i/84 Jap/0221HUJA/02 Thai/41571/98 SriL/SL17/78 Thai/40034/94 Thai/40557/91 Jap/9910HuJA/99 Thai/40100/95 Thai/40229/96 Thai/40358/92 Thai/40600/94 Thai/40438/02 Thai/40501/02 Thai/40761/00 Thai/40521/99 Thai/41448/98 Thai/40109/96 Thai/40485/95 Thai/40261/92 Thai/40792/93 Thai/40348/91 Thai/40420/93 Thai/40233/86 Thai/40096/82 Thai/40417/84 Thai/40194/83 Thai/24024/84 Thai/85 Thai/40182/85 Thai/40104/86 Thai/40116/81 Thai/7801/78 Thai/40087/77 Thai/40113/76 Thai/40092/77 Thai/40696/80 Mal/P71006/69 Jap/331HUJA/96 SriL/44750/78 Jap/461NIID/61 Jap/473NIID/73 Phil/H241/56 Phil/12123/84 Phil/16589/64 Thai/C2443/63 Thai/40017/97 ThD40164 Thai/40476/97 Thai/41270/98
II
Endemic/ epidemic
I
III
Sylvatic
FIGURE 5 Phylogenetic relationships of DENV-4. The phylogeny was inferred based on the E nucleotide sequence in the GenBank library, using Bayesian analysis (one million reiterations) and all horizontal branches are scaled according to the number of substitutions per site. Bayesian probability values are shown for key nodes. Strains are abbreviated as follows: Country abbreviation/strain/year.
History and Evolution of Human Dengue Emergence
11
(CE 992) (Gubler, 1997). These reports described a disease called ‘water poison,’ due to its association with water-associated flying insects, and whose clinical description included fever, rash, arthralgia, myalgia and hemorrhagic manifestations. The next reports of a similar illness appear almost seven centuries later, describing an acute illness with prolonged convalescence in the French West Indies and Panama during 1635 and 1699 respectively (Gubler, 1997). A century later (1779–1788), the first reports of a possible DEN pandemic in Batavia (present day Jakarta) (Bylon, 1780; Pepper, 1941), Cairo (Christie, 1881; Hirsch, 1883), Philadelphia (Rush, 1789), and Cadiz and Seville, Spain (Christie, 1881) were described. These accounts describe for the first time evidence for a widespread DENV geographic distribution (or at the very least of an illness very similar to DEN), reaching pandemic proportions by 1788. The etymological origins of the term ‘dengue’ are uncertain. Christie provided an early account of the origin of the term based on his personal experiences from the Zanzibar epidemic in 1870 (Christie, 1881). Early in the epidemic some of his younger native patients called it ‘baridiyabis,’ meaning rheumatism, and his Indian patients used the term ‘Homa mguu,’ meaning leg fever; the Hadramaut Arabs used ‘abou-ndefu,’ which was adapted to ‘abou-madefu’ by the black population, meaning father of beards. As the epidemic intensified, the older residents of Zanzibar recognized the disease as identical to that of the 1822 epidemic, and gave it its earlier designation, ki-dinga pepo, which in Swahili means ‘a disease characterized by a sudden cramp-like seizure, caused by an evil spirit’ (Christie, 1881). It is believed that this phrase entered into the Caribbean via the flourishing slave trade out of East Africa sometime in the early 1800s. Early accounts from St. Thomas, refer to the disease as ‘Dandy fever’ and ‘the Dandy’, reflecting the stiffness in motion of people affected by severe the joint and muscle pain associated with the disease (Christie, 1881; Steadman, 1828). A few years later, when the disease arrived in Cuba the designation had changed to ‘dunga,’ which was later transformed into ‘dengue’, meaning fastidiousness and prudery from the Spanish ‘andar en dengue’ (Christie, 1881). Although the disease was known with several other names (Table I), reflecting cultural or geographic etymologies, the term ‘dengue’ had been universally adopted. Although Benjamin Rush provided the first well-known, detailed clinical descriptions of the illness now believed to represent DENV infection, as well as applied the term ‘‘break-bone fever’’ to describe an epidemic in Philadelphia (Rush, 1789), credit should also attributed to David Bylon, the medical officer who first observed and described a 1779 epidemic taking place in Jakarta, Indonesia (Bylon, 1780). He observed the sudden onset and spread of the disease that swept through the region, and to which he also fell victim. He called it ‘knokkel-koorts’ or knuckle fever, and emphasized the severity of the pains, the presence of rash, and
12
Nikos Vasilakis and Scott C. Weaver
TABLE I Terms used to describe dengue in various parts of the world
a b
Terma
Location (period)
Coup de barre Scarlatina rheumatica Bilious remitting fever Knokkel-koorts (knuckle fever) Abu rocab (knee trouble) La Piadosa (the merciful) Ephemeral fever Three days’ fever Dandy fever Bouquet Dunga Fievre des dates (fever of the dates) Polka feverb Bonon (sighs) Mal de genoux (knee fever) Trancazo (stroke) Baridiyabis (rheumatism) Homa mguu (leg fever) Abou-ndefu Ki-dinga pepo Fievre rouge, giraffe
French West Indies (1635) Philadelphia (1779) Philadelphia (1779) Batavia (1779) Cairo (1780) Cadiz and Sevilla (1784–1785) Rangoon (1824) Calcutta (1824) St. Thomas (1820s) St. Thomas (1827) Cuba (1829) Jeddah (1847–1856) Brazil (1846–1849) Hawaii (1847–1856) Benghazi, Tripoli (1856) East Africa (1870) East Africa (1870) East Africa (1870) East Africa (1870) East Africa (1880) Syria (1870–1873)
Information for this table was obtained from Christie, 1881; Hirsch, 1883; Leichtenstern, 1896; Rush, 1789; Steadman, 1828. Christie believes that the term does not describe dengue (Christie, 1881).
tendency to relapse. Bylon cryptically concludes his report that ‘. . . a well known disease which, however, in the memory of man here in Batavia has never reached an epidemic, and which has, therefore, seemed wondrous to the inhabitants’ (Bylon, 1780; Pepper, 1941), suggests that the disease (most likely endemic DENV) had been known to Indonesians for quite some time. Around the same time, an epidemic of a similar disease was occurring in Cairo, as well as Alexandria, Egypt. The onset of the epidemic in the local population was described by the chronicler Gaberti, as ‘abu rokab’ or knee-trouble, and was characterized by sudden onset of self-limited fever, followed by prolonged defervescence accompanied by pain of the joints, knees and extremities (Hirsch, 1883). Although, the clinical descriptions of the 1699 Panama, as well as the 1789 Indonesia, Batavia and Cairo outbreaks were compatible with DEN fever (DF), it is possible that some these outbreaks were caused by Chikungunya virus, which causes clinical
History and Evolution of Human Dengue Emergence
13
illness nearly indistinguishable from DEN (Carey, 1971; McSherry, 1982). Interestingly, Theiler cast similar doubts, but only for the 1779 Cairo epidemic (Theiler and Downs, 1973). A series of DEN or DEN-like epidemics cris-crossed the globe, from Africa to India to Oceania to the Americas, from 1823 to 1916 (Table II) (Brown, 1977; Christie, 1881; Hirsch, 1883; More, 1904). The historical record suggests the occurrence of at least five pandemics during this time period, lasting 3–7 years, and probably caused by the same DENV serotype and transported among geographic regions by the slave trade and commerce (Brown, 1977; Christie, 1881; Gubler, 1997). Furthermore, Leichtenstern recognized DEN as a disease of seaports and costal regions that could also spread inland along rivers like the Ganges and the Indus in India, or the Mississippi in the United States (Leichtenstern, 1896). Of course there is no way in knowing what DENV strain or serotype was involved, since all these epidemics took place before serotype identification was possible.
1. Modern DENV diagnostics and elucidation of transmission The first step in serodiagnostics took place in Fort McKinley in the Philippines, where Ashburn and Craig, experimenting with human volunteers, concluded that the etiologic agent of DEN was filterable (i.e., a virus) (Ashburn and Craig, 1907). At about the same time, Graham in Beirut, and Cleland in Australia were investigating the role of Ae. aegypti in the transmission of DENV using human volunteers (Cleland et al., 1916; Graham, 1903). Ae. aegypti transmission was confirmed in 1926 by the extensive and well-controlled experiments of Siler, Hall and Hitchens (Siler et al., 1926), followed by the incrimination of Ae. (Stegomyia) albopictus in 1931 (Simmons et al., 1931). By the end of the second decade of the twentieth century, DEN behavior in Southeast Asia, the Indian subcontinent and the Philippines had changed from the sudden onset of urban epidemics to endemicity, a trend attributed to the gradual invasion of Ae. aegypti (Table III) (Daniels, 1908; Edwards, 1932; Smith, 1956; Stanton, 1920; White, 1934). On the other hand, in the Caribbean DEN remained intermittently active. Nevertheless during the epidemic of 1922, which began in Galveston, Texas and spread throughout the Gulf and southern Atlantic states, as well as the Caribbean, close to 2 million people were infected. Of lesser impact was the epidemic in South Africa and Egypt, where close to 100,000 infections may have occurred (Edington, 1927; Kamal, 1928), followed by the great epidemic of 1927–1929 in Greece where at least one million people were infected (Cardamatis, 1929; Copanaris, 1928). By the mid 1940s DEN was eliminated altogether from the Mediterranean basin, mainly due to the elimination of Ae. aegypti.
TABLE II
Epidemics of dengue from 1824–1916a
Year(s)
Geographic location
Comments
1823–1828
Zanzibar, Suez, and Pondicherry to Calcutta; Burma, and Ganges Valley; Caribbean Islands and Savannah; Lesser Antilles, Cuba, North Colombia, Mexico, Southeast USA
1835–1851
Arabian coast; Senegambia, Cairo and Rio de Janeiro; India and Hawaii; New Orleans; Gulf Coast and eastern seaboard USA; Reunion, Mauritius and Tahiti Zanzibar and Dar es Salaam; Port Said, Arabian Peninsula, and gradually throughout India; Burma, Singapore, Indonesia, Shanghai, Taiwan, Mauritius; Southern USA Gibraltar; Cyprus; Greek Islands, Turkey (Aegean and Black Sea ports) and Syria Australia, Indo-China, and China China, Indo-China, India, Singapore, Australia, Galveston, Panama, Cuba, and Colombia Panama, Chile, Argentina, Australia and India
Pandemic was preceded by an epidemic in Peru affecting nearly 50,000 people. First observation from the W. Indies epidemic of 1827 that patients of African origin had a lower incidence and severity of the disease Epidemic returned to Lima (1851), and appeared in Spain (1865) and Port Said (1868)
b
1870–1873b
1887–1889 1894–1897 1901–1907 1912–1916 a b
Pandemic was followed by silence until 1880, when it brought dengue for the first time in the Mediterranean, appearing in ports in Greece, the Levant and the Red Sea
First reports of higher incidence of disease among Chinese patients
Information for this table was compiled from Brown, 1977; Christie, 1881; Hirsch, 1883; Khan, 1913; Leichtenstern, 1896; Maxwell, 1839; More, 1904; Skae, 1902; Smith, 1956; Steadman, 1828. Carey suggested that the dengue epidemics in Zanzibar (1823 and 1870), as well as India (1824, 1871, and 1902), may be attributed to chikungunya infections due to severe arthralgia (Carey, 1971).
TABLE III Epidemics of dengue worldwide from 1922–2007 Year(s)
Geographic location
Comments
1922–1929
Galveston to southern Atlantic states; Caribbean Islands; South Africa to Egypt; Greece
1941–1945
East Africa; Caribbean Islands; Australia to Hawaii; Papua New Guinea to Japan
1956–1959
The Philippines and Thailand
1963–1969
Jamaica, Puerto Rico, northern Colombia; Nigeria; India to throughout southeast Asia
1971–1989
Oceania, Myanmar and Malaysia to India; Thailand to Indonesia to China; Caribbean to Central America to northern South
Epidemic started in Galveston, Texas and spread throughout the gulf, southern Atlantic states and most of Caribbean Islands. First documented cases in Africa (Durban, 1927). Great epidemic of Athens, Greece (1927–1929); a million were infected; last DENV epidemic of record in Europe Dengue epidemics among military personnel in East Africa and the Caribbean. Pandemic throughout the entire area of the Pacific theater of operations. First isolation of DENV-1 (1943) and DENV-2 (1943–1944). First detection of homotypic immunity following infection and development of HI test First documented cases of DHF. First isolation of DENV-3 and DENV-4. Paucity of DENV epidemics in the Americas, Africa and Oceania Reappearance of dengue in the Americas. First isolation of DENV-1 in Africa and evidence of sylvatic DENV-2 activity in humans (Ibadan, 1966). Increased disease severity in DENV epidemics in SE Asia. Formulation of ADE theory Reintroduction of DENV-1 and DENV-4 in Oceania; Reappearance in China after a 30-year absence; reintroduction of DENV-1 in the Caribbean; sylvatic DENV-2 isolation from humans (1970), monkeys (1981) and sylvatic Aedes spp. mosquitoes (1974) in West Africa; introduction of SE Asian (continued)
TABLE III (continued) Year(s)
1990–2007
Geographic location
Comments
America; West Africa; Cuba; East Africa SE Asia, India, Bangladesh, Singapore, Australia, Indonesia, East Africa, Senegal, Brazil, Argentina, Nicaragua, French Guinea, Peru, Oceania
DENV-2 genotypes in the Americas (1981) associated with increased disease severity Global distribution of all DENV serotypes complete; intense activity of epidemics with global peak of morbidity and mortality rates
History and Evolution of Human Dengue Emergence
17
2. Effect of World War II on DEN epidemiology The onset of World War II brought immense ecologic, demographic, and epidemiologic changes leading to a new relationship between DENV and humans. Ecologically, the destruction of existing water distribution systems led to domestic water storage practices, and along the abandonment of war materials led to an abundance of sites ideal for the development of Ae. aegypti larvae. Furthermore, transport of troops and supplies over long distances resulted in the importation of mosquitoes into new geographic regions. These ecological changes not only greatly enhanced the densities of Ae. aegypti but also expanded their geographic distribution. Demographically, the war brought of large numbers of troops and refugees susceptible to DENV infection, contributing to the dispersal of viruses as well as increasing the pool of susceptible people for epidemic DENV transmission. In fact, between 1941 and 1945, a series of DEN epidemics were raging among military personnel in East Africa and the Caribbean, as well as a pandemic that encompassed in the entire pacific theater of operations, from Australia to Hawaii and from Guinea to Japan (Brown, 1977; Gubler, 1997; Hota, 1952; Sabin, 1952). The events of World War II heightened awareness of DEN, resulting in the establishment of scientific commissions to study the disease and its etiologic agent. Japanese scientists isolated DENV-1 (Mochizuki strain) in Nagasaki in 1943, as well as other DENV-1 strains (Sota and Kin-A) from affected patients elsewhere in Japan (Hota, 1952). Later, Sabin isolated both DENV-1 (Hawaii strain) and DENV-2 (New Guinea C strain) from U.S. soldiers in 1944 (Sabin, 1952). Sabin’s group also identified the presence of homotypic immunity following infection, and developed hemagglutination inhibition test for serodiagnosis (Sabin, 1952; Sabin and Schlesinger, 1945). The cessation of World War II in the Pacific theatre led to uncontrolled urbanization, where millions of people moved into cities with inadequate housing, water distribution systems, as well as sewer and waste management. In such environments, Ae. aegypti reached high densities and the movement of people dispersed the DENV serotypes among regions. Overall, these ecologic and demographic changes created ideal conditions for the emergence of DHF in Southeast Asia. Although occurrences of severe and fatal hemorrhagic disease associated with DENV infection were reported as early as the 1780 Philadelphia and 1927–1929 Greece epidemics (Copanaris, 1928; Rush, 1789), they were probably rare and did not pose a severe public health problem. The first well documented cases of DHF were associated with epidemics in Thailand and in the Philippines during in the 1950s, and initially were thought to be a new disease (Hammon et al., 1960a,b). In fact, the viruses isolated from patients with hemorrhagic disease during the 1956 Philippine epidemic were members of the serotypes 3 (H87 strain) and 4 (H241 strain) (Hammon et al., 1960a,b).
18
Nikos Vasilakis and Scott C. Weaver
While a number of epidemics occurred in Southeast Asia in the aftermath of World War II, no epidemics were reported in the Americas, Africa and Oceania for the next 20 years. A major factor in this quiescence, at least in the Americas, was the initiation of an Ae. aegypti eradication program under the auspices of the Pan American Health Organization (PAHO); in Africa, the absence of epidemics may be attributed to poor surveillance. The PAHO program, undertaken to prevent urban epidemics of yellow fever, led to the eradication of Ae. aegypti in all American countries except Suriname, Guyana, French Guiana, Venezuela, some Caribbean Islands, and the United States (Gubler, 1997). Discontinuation of the program during the early 1970s allowed for the gradual reinfestation of the region by Ae. aegypti, a process that continued well into the 1990s. The 1960s saw a dramatic increase in DENV activity in many tropical locations. Dengue reappeared in the Americas in 1963 in Puerto Rico and Jamaica with the introduction of a DENV-3 of Asian origin that caused several epidemics in the Caribbean, northern Colombia and possibly Venezuela (Ehrenkranz et al., 1971; Morales et al., 1973; Neff et al., 1967; Russell et al., 1966; Spence et al., 1969; Ventura and Ehrenkranz, 1976). In Africa, the newly established surveillance program (1964) by the Rockefeller Foundation at the University of Ibadan, Nigeria, documented the endemic transmission of DENV-1 and DENV-2 in humans (Anonymous, 1969; Carey et al., 1971). In Oceania, two small outbreaks of DENV-3 occurred in 1964 and 1969 after an absence of 20 years (Laigret et al., 1967; Saugrain et al., 1970). However, in India (Balaya et al., 1969; Myers et al., 1965, 1968; Ramakrishnan et al., 1964) and in Southeast Asia there were a series of epidemics in Vietnam (Halstead et al., 1965), the Philippines (Basaca-Sevilla and Halstead, 1966), Singapore (Chan et al., 1965; Lim et al., 1961), Malaysia (Rudnick et al., 1965) and Thailand (Halstead et al., 1967), associated with increased incidence of disease severity. By the end of the decade, all four DENV serotypes were circulating throughout Southeast Asia and the Indian subcontinent. Subsequently, through prospective field studies in Thailand, an association was inferred between secondary infections and severity of DEN disease (Halstead et al., 1967; Russell et al., 1967), which eventually led to the antibody-dependent enhancement theory (ADE) of DEN pathogenesis (Halstead et al., 1973c). In the 1970s, there were several DENV epidemics in Oceania (Barnes and Rosen, 1974; Gubler et al., 1978; Loison et al., 1973; Maguire et al., 1974; Moreau et al., 1973), that allowed for the first time the evaluation of the epidemic potential of DENV, based on viremia, disease severity and dynamics of transmission as it moved through human populations (Gubler et al., 1978). Some of the epidemics that occurred in the area in the late 1970s were mainly due to the reintroduction of DENV-1 and DENV-4 (Gubler, 1997). Similarly, several epidemics of increased severity
History and Evolution of Human Dengue Emergence
19
took place throughout Southeast Asia in a wave-like fashion starting in Myanmar and Malaysia (George et al., 1974; Lim et al., 1974; Thaung et al., 1975; Wallace et al., 1980) then moving into India (Mathew et al., 1977), Thailand and Indonesia (Kho et al., 1981; Okuno et al., 1980) and finally into China in 1978 for the first time after an absence of 30 years (Fan et al., 1989). Reintroduction of DENV-1 into the Caribbean in 1977 led to epidemics in Central and northern South America (Gubler, 1997; Morens et al., 1986; Uzcategui et al., 2001). While only limited DEN outbreaks were reported in Africa in the 1970s (Fagbami and Fabiyi, 1976), serological surveillance indicated the endemicity of DEN in West Africa (Fagbami, 1977, 1978; Saluzzo et al., 1986a).
3. Discovery of sylvatic DENV In 1970, DENV-2 (strain DakAr HD10674) was recovered from a young girl in Bandia, Senegal (Robin et al., 1980), which subsequent phylogenetic analysis classified as an ecologically and genetically distinct sylvatic genotype (Rico-Hesse, 1990; Wang et al., 2000). Sylvatic DENV are now understood to be both ecologically and evolutionary distinct DENV lineages whose enzootic transmission cycle occurs in the sylvan environs of southeast Asia and west Africa, presumably between non-human primates and arboreal canopy–dwelling Aedes mosquitoes. Most DENV strains isolated in Africa are sylvatic, as determined genetically. The sylvatic DENV and their transmission are described in greater detail below.
4. The rise of DEN hyperendemism and DHF An important characteristic of DEN epidemics in the Americas during the 1960s and 1970s was the circulation of a single serotype at any given time within a region (hypoendemicity). This trend changed with the introduction of a Southeast Asian strain of DENV-2 into Cuba, probably from Vietnam in 1981 (Kouri et al., 1983; Rico-Hesse, 1990), followed by an increase in the severity DEN during both Cuban and Venezuelan epidemics (Kouri et al., 1989; Uzcategui et al., 2001). Important observations from the Cuban epidemic of 1981 were the putative role of host genetics (Bravo et al., 1987; Kouri et al., 1987), gender and age (Guzman et al., 1984, 2002) in influencing the severity of disease. Several other epidemics of DENV-1, -2, and -4 took place during the 1980s throughout Central America (Kouri et al., 1991; Lorono Pino et al., 1993) and the Caribbean islands (Pinheiro, 1989), albeit of reduced disease severity. In essence, the introduction of new genotypes and the gradual increase in circulation of multiple serotypes, as well as the increased severity of disease in the Americas, mirrored the events that occurred in Southeast Asia in the 1950s and 1960s. In Africa, even in the absence of effective surveillance, the 1980s were characterized by an increase in apparent clinical disease and increased circulation of all serotypes, which curiously was not
20
Nikos Vasilakis and Scott C. Weaver
associated with any increase in disease severity, except on rare occasions (Gubler et al., 1986). In 1983, the first evidence of autochthonous DENV-4 transmission in Senegal was detected (Saluzzo et al., 1986b), and a year later DENV-3 transmission was identified in Mozambique (Gubler et al., 1986). Furthermore, several epidemics were reported in Kenya ( Johnson et al., 1982), Burkina Faso (Gonzalez et al., 1985), Somalia (Botros et al., 1989; Saleh et al., 1985), and Sudan (Hyams et al., 1986). In Southeast Asia, two major epidemics, in Malaysia (Fang et al., 1984) and Thailand (Ungchusak and Kunasol, 1988), were the worst experienced at that point in their history in terms of morbidity and mortality. Several other countries in the latter region recorded also increased incidence in disease severity at this time (Hayes et al., 1988; King et al., 2000; Rathavuth et al., 1997). By the 1990s, the global distribution of all DENV serotypes had been completed mainly due to expanding urban populations, increased vector density due to unsustained control programs, and the dramatic increase in commercial air travel facilitating the rapid movement of viremic humans. Since the end of World War II, these factors have converged into a potent mix for the rapid and dramatic re-emergence of DENV associated with increasing disease severity throughout the tropics. By the middle of the decade several epidemics were documented globally (Cobra et al., 1995; da Cunha et al., 1997; Padbidri et al., 1995; Rathavuth et al., 1997; Reynes et al., 1994; Rodier et al., 1996; Sharp et al., 1995; Strobel et al., 1998; Traore-Lamizana et al., 1994), but the end of the decade was characterized with intense activity of epidemics whose morbidity and mortality rates peaked globally in 1998 (Aziz et al., 2002; Bouree et al., 2001; Corwin et al., 2001; Cunha et al., 1999; Dove, 1998; Ha et al., 2000; Harris et al., 2000; Hussin et al., 2005; Thomas et al., 2003). A longitudinal study examining the spatiotemporal dynamics of DENV infections with increased severity in Thailand during a 14-year period (1983–1997) demonstrated a 3-year periodicity of epidemics with successive predominance of different serotypes (Cummings et al., 2004). A wavelike pattern of radially moving infections from the metropolitan epicenter (Bangkok), underscored the complex vector- and host-pathogen, as well as environmental and ecological interactions that governs DEN epidemics. The annual incidence of DF and DHF, as well as intensity of epidemics, has increased dramatically around the world with the dawn of the new century. In the Americas, several major epidemics have been documented. Although DENV-3 was introduced in Brazil in 1998 (Rocco et al., 2001), the first autochthonous DENV-3 cases were not detected until 2000, leading to explosive epidemics throughout the country (Cordeiro et al., 2007; Nogueira et al., 2001). Within a span of 8 years (1994–2002) the incidence of DEN increased from 37 to 454 per 100,000 inhabitants (Siqueira et al., 2005). Dengue epidemics continued in 2006 and 2007, with over 600,000 documented human infections (PAHO, 2007), probably
History and Evolution of Human Dengue Emergence
21
leading to spillover in Paraguay to cause the first major epidemic there with over 25,000 confirmed cases (PAHO, 2007). Dengue activity also increased in Central America and Mexico, peaking in 2007 with over 100,000 confirmed cases (Diaz et al., 2006; PAHO, 2007). By 2007, 24 countries in the region had reported over 22,000 confirmed cases of DHF (PAHO, 2007). In Southeast Asia DF is so widespread that eight of the region’s 11 countries reported disease to the WHO every year up to 2003. By 2006, all countries in the region except the Democratic Peoples’ Republic of Korea reported DEN cases (WHO, 2007). In 2005, an outbreak, with a high case fatality rate of 3.55% was first reported in Timor-Leste (Kalayanarooj et al., 2007). Nepal reported DF cases for the first time in 2004 (Pandey et al., 2004). In China, an outbreak of DENV-1 was initiated by a viremic traveler from Thailand, and its spread throughout the region was attributed to high concentrations of Ae. albopictus mosquitoes. These mosquito populations resulted from the creation of extensive larval development sites by logging caused by typhoon Rananim (Xu et al., 2007). In Singapore, after a 2 year decline, annual incidence rates of DF skyrocketed from 25/100,000 in 2000 to 340/100,000 in 2005 (Ooi et al., 2006). In Indonesia, where DEN is hyperendemic, annual incidence rates more than doubled from 159/100,000 in 2000 to 344/100,000 in 2004 (Kusriastuti and Sutomo, 2005). In 2006 Indonesia alone reported 57% of all DF cases and over 70% of fatalities due to DHF occurring in the region (WHO, 2006). As described above, DENV have probably had a close relationship with humans for the last 1,700 years, which only during the past few decades has intensified due to expanded commerce, large population movements, changing ecologic conditions and unstainable vector control programs. At present, all DENV serotypes have reached global hyperendemicity and will likely continue to cause epidemics of various intensities and pathogenic severity in rolling cycles 3–5 years apart. Today, it is estimated that about 3 billion people are at risk for DENV infection in large urban and periurban areas located throughout the tropics. By current estimates, approximately 100 million DENV infections occur annually, leading to 500,000 cases of DHF and 20,000 deaths. Thus DENV have become the most important arboviral pathogens of humans.
B. DENV transmission cycles Although most human infections today are caused by DENV strains that rely only on humans as reservoir and amplification hosts, and principally on Ae. aegypti and/or Ae. albopictus as vectors, the ancestral forms of DENV are believed to be viruses that circulate in forest habitats, presumably among nonhuman primates, transmitted by arboreal mosquitoes. These DENV sylvatic cycles have been demonstrated in Asia, where serologic evidence as well as virus isolation suggests transmission of
22
Nikos Vasilakis and Scott C. Weaver
sylvatic strains of DENV-1, -2, and -4 among Macaca and Presbytis monkeys vectored by Ae. niveus mosquitoes (Peiris et al., 1993; Rudnick and Lim, 1986). In West Africa, only sylvatic DENV-2 has been shown to circulate regularly between Erythrocebus patas monkeys and various sylvatic Aedes sp., including Ae. taylori, Ae. furcifer, Ae. vitattus, and Ae. luteocephalus, in a sylvatic focus near Kedougou, Senegal (Diallo et al., 2003, 2005; Rodhain, 1991; Saluzzo et al., 1986b). The transmission cycle that today results in most human DENV infections includes Ae. Aegypti as the principal vector. This mosquito originated in Africa, where the ancestral form, Ae. aegypti formosus, uses treeholes as larval development habitats (Tabachnick and Powell, 1979). The subspecies that transmits DENV in peridomestic habitats, Ae. aegypti aegypti, adapted in Africa to use artificial water containers as its larval habitat and was later transported to nearly all tropical and subtropical locations. The derived form, Ae. aegypti aegypti, now lives in close contact with people in urban settings by relying on artificial water containers for its larval habitats, resulting in endophilicity that increases contact with people. The reliance on blood (instead of plant carbohydrates) for its energetic needs, as well as its endophilicity result in a high frequency of multiple host contacts during a single gonotrophic cycle (Harrington et al., 2001). These behavioral and ecological traits, probably more than its innate susceptibility to DENV, contribute to Ae. aegypti’s success as an endemic and epidemic vector.
1. Sylvatic DENV cycles Sylvatic DENV are both genetically and ecologically distinct from their urban, endemic/epidemic counterparts. Their transmission cycle most likely involves non-human primates as reservoir hosts and several arboreal canopy-dwelling Aedes spp. mosquitoes. Although little attention has been paid to these sylvatic cycles in recent years, the seminal work of Smith and Rudnick in the 1950s and 1960s in Asia remains illuminating. Gordon Smith, working in Penang, Malaya, demonstrated the presence of DENV antibodies in tree-dwelling animals, such as wild monkeys, slow lorises, civets and squirrels. In contrast, very few ground-dwelling animals were seropositive, thus suggesting for the first time a canopy-dwelling DENV vector (Smith, 1956). However, in a later study, he confirmed serologically that only monkeys were DENV-positive (Smith, 1958). Subsequently, working in Malaysian forests of various ecologic types (primary dipterocarp, freshwater peat swamp and mangrove swamp) away from normal human activity, and where Ae. aegypti were completely absent, Rudnick demonstrated the presence of widespread DENV-neutralizing antibodies in wild monkeys (Macaca nemestrina, M. fascicularis, Presbytis cristata, and P. melaphos)(Rudnick, 1965). Follow up studies, using 27 sentinel monkeys (M. fascicularis and P. obscura) in the forest
History and Evolution of Human Dengue Emergence
23
canopy of isolated primary rain forest, led to DENV-1 (P72–1244), DENV-2 (P8–1407, P72–1273, and P72–1274 strains), and DENV-4 (P75– 481 and P75–514 strains) isolation, but no virus isolation was reported from 19 sentinel monkeys exposed on the ground. Although DENV-3 were not isolated, seroconversions in sentinel monkeys suggested their existence (Rudnick and Lim, 1986). An endemic strain of DENV-2 was also isolated from Ae. albopictus (P8–377; see Fig. 3), a vector that was found only at ground level, and DENV ––– 4 (P75–215 strain) was isolated from Aedes (Finlaya) niveus s. l. The latter is abundant in the forest canopy, but will descend to the ground if primates are present, and in Malaysia consists of six species (Ae. pseudoniveus, Ae. subniveus, Ae. vanus, Ae. albolateralis, Ae. niveoides and Ae. novoniveus); all are primatophilic (Rudnick, 1986). Furthermore, in a serum survey of 300 forest-dwelling Orang Asli aborigines, the vast majority had neutralizing antibodies due to DENV, although no clinical DEN was reported among this group (Rudnick, 1986). Similar observations have been reported from the Philippines, where high rates of DENV neutralizing antibodies were present among isolated aborigines living in a region devoid of Ae. aegypti (Rudnick et al., 1967). Although in this area there was no evidence of sylvatic DENV transmission, these data suggest sylvatic transmission there as well. Collectively, the studies of Rudnick and colleagues suggested enzootic, sylvatic DENV cycles in the canopy of the forests of Malaysia with all 4 DENV serotypes transmitted by members of the Ae. niveus spp., among Macaca and Presbytis spp. monkeys. (Fig. 6). In theory this enzootic cycle
Ae. furcifer Ae. luteocephalus Ae. aegypti formosus
Ae. furcifer
“Zone of emergence”
Ae. aegypti aegypti Ae. albopictus
??
FIGURE 6 The transmission cycles of DENV, showing the sylvatic origins and the ‘‘zone of emergence’’ where these cycles contact human populations in rural areas of West Africa (DENV-2) and Asia (probably all 4 DENV serotypes).
24
Nikos Vasilakis and Scott C. Weaver
could occur in all primary forests of tropical Asia where the zoonotic reservoir hosts and vectors exist (Yuwono et al., 1984). Sylvatic DENV transmission cycles were also suspected to exist in West Africa, where DENV-2 antibodies had been detected in nonhuman primates inhabiting both gallery and lowland forests in Nigeria (Fagbami et al., 1977). However, isolation of DENV-2 in 1974 from forest Ae. luteocephalus mosquitoes in eastern Senegal, collected far from inhabited areas, provided for the first credible evidence that sylvatic DENV cycles occur in West Africa (Robin et al., 1980). Furthermore, evidence that non-human primates serve as amplification hosts was provided from a retrospective serologic study of non-human primates and humans inhabiting the same Senegal region. This study indicated the presence of successive epizootics among non-human primates in 1974 and 1981, in the absence of epidemic transmission among humans residing in outlying forest villages (Saluzzo et al., 1986a). During the latter epizootic, DENV-2 was isolated from Erythrocebus patas monkeys (Cornet et al., 1984), whereas in the previous rainy season over 100 strains of DENV-2 were isolated from primatophilic Ae. taylori, Ae. furcifer, Ae. opok, Ae. luteocephalus and Ae. africanus in Guinea, Coˆte d’Ivoire, and Burkina Faso (Cordellier et al., 1983; Hervy et al., 1984; Roche et al., 1983; Rodhain, 1991). As in Senegal, there was no indication of a spillover epidemic (enzootic virus transmission into a small, localized groups of people, confined to a village or a small area due to favorable ecological conditions, such as increased vector densities) in the human populations of these countries. Moreover, the 1982 DENV-2 epidemic in Burkina Faso is suspected to have originated through the introduction of an endemic DENV2 strain from the Seychelles Islands (Rico-Hesse, 1990). Similarly, the last recorded DENV amplification cycle of 1999–2000 in Kedougou, Senegal led to isolation of several DENV-2 strains from mosquito pools, with no recorded human clinical cases in the region (Diallo et al., 2003). This suggested that either the sylvatic virus is confined to the forest habitat or, if human–mosquito–human transmission occurs, it remains at low levels and results in mild illness. Surprisingly, six sylvatic DENV-2 strains were isolated within four villages (Ngari, Silling, Bandafassi, and Kenioto) from collections of the arboreal Ae. furcifer mosquito, which suggests that it could act as a bridge vector for sylvatic DENV dissemination into human habitats (Diallo et al., 2003). Domestic Ae. aegypti, the principle vector of epidemic DENV worldwide, is scarce or possible absent from this area, whereas the sylvatic Ae. aegypti formosus is abundant. Further research suggested that the zoophilic Ae. aegypti formosus plays little or no role in sylvatic DENV transmission because it is relatively refractory to infection (Diallo et al., 2005). As the historical record indicates, sylvatic amplification cycles of DENV-2 in West Africa appear with oscillating frequency (1974,
History and Evolution of Human Dengue Emergence
25
1980–1982, 1989–1990, 1999–2000), with silent intervals (lack of virus isolates from mosquitoes) of about 8 years in length. Collectively, the data suggest that in West Africa, only sylvatic DENV-2 circulates regularly, between Erythrocebus patas monkeys and various sylvatic Aedes sp., including Ae. (Diceromyia) taylori, Ae. (Diceromyia) furcifer, and Ae. (Stegomyia) luteocephalus (the latter two as the principal enzootic vectors), in a sylvatic focus in Kedougou, Eastern Senegal (Fig. 6). Sylvatic DENV transmission cycles have not been documented in the Americas. An ecological study conducted in Panama in 1954, where sera from 105 wild-caught non-human primates were examined for the presence of DENV-1 and DENV-2-specific antibodies, revealed no evidence of enzootic circulation (Rosen, 1958b). Evidence of DENV-2 neutralizing antibodies was obtained in Ayoreo Indians living in an isolated forested region (Rincon del Tigre) of Bolivia, where Ae. aegypti are not present, suggests the presence of a sylvatic transmission cycle (Roberts et al., 1984). Ae. (Gymnometopa) mediovittatus, a forest mosquito that is also adapted to peridomestic habitats and shares larval sites with Ae. aegypti, could in theory support such cycles (Gubler et al., 1985b). Several species of New World non-human primates, including Cebus capucinus, Ateles geoffroyi, Ateles fusciceps, Alouatta palliata, Marikina geoffroyi, Saimiri orstedii and Aotus trivirgatus, are susceptible to DENV-1 and DENV-2 infection. They develop viremia in the absence of clinical illness, as well as neutralizing antibodies. However, the viremia profiles are believed to be insufficient to initiate oral mosquito infection (Rosen, 1958a), and there is no evidence that nonhuman primates are exposed to or acquire these viruses in forests of the Americas. Further studies of the potential for the establishment of a sylvatic DENV cycles in the Americas are needed. Yellow fever virus, which was also introduced into the Americas on sailing ships from Africa, subsequently became established in a sylvatic primate cycle throughout many areas of South America. In Africa, Yellow fever virus shares many characteristics of its enzootic cycle with DENV, including overlaps in vector species and primate hosts. It also uses Ae. aegypti as its epidemic mosquito vector but the human–mosquito–human cycles does not appear to be sustained for decades or centuries like the DENV. Considering these many similarities, it is certainly plausible that, with increased levels of endemic DENV transmission in the Americas, opportunities for the establishment of an enzootic, sylvatic cycle are abundant. In rural areas of Africa, and Asia (also known as the ‘zone of emergence’) where enzootic vector(s) often reach high densities, DENV-2 can transfer between non-human primates and humans (Fig. 6). In Asia, the studies of Rudnick suggested that zoonotic A. (Finlaya) niveus vectors descend to the ground to feed on humans where Ae. albopictus are abundant, thus allowing the transfer of virus from the forest into human
26
Nikos Vasilakis and Scott C. Weaver
habitats. In fact, in a companion study where the incidence of DENV infection was measured in rural areas adjacent to forests characterized by low, immobile human densities and devoid of Ae. aegypti, it was established that (i) the highest rates of DENV infection were among the rural populations living adjacent to the forest, (ii) mild fevers of short duration, presumably due to DENV infection, occasionally occurred, and (iii) Ae. albopictus was the principal vector on the ground (Rudnick, 1986). This scenario parallels that in rural areas adjacent to forests of West Africa, where the principal bridge vector between forest and village is Ae. furcifer. Similarly, in West Africa (albeit at lower incidence rates), DENV-2 circulates among rural populations in the absence detected clinical illness, probably with the presentation of mild signs and symptoms (Monlun et al., 1992; Saluzzo et al., 1986a). Other species of Aedes mosquitoes, such as Ae. (Stegomyia) polynensiensis, Ae. mediovittatus, and Ae. (Stegomyia) scutellaris have been reported to be responsible for DENV transmission in rural areas of the Pacific, based on epidemiological observations (Mackerras, 1946; Rosen et al., 1954) or their ability to transmit experimentally (Gubler et al., 1985; Rosen et al., 1985). Ae. (Protomacleaya) triseriatus could also be considered a potential DENV vector based on experimental transmission studies (Freier and Grimstad, 1983), but its distribution is temperate and does not overlap with DENV-endemic locations. In the latter experimental studies, the tested mosquitoes exhibited a higher susceptibility to DENV oral infection than did Ae. aegypti. This observation has been also confirmed with Ae. albopictus (Jumali et al., 1979; Moncayo et al., 2004; Rosen et al., 1985).
2. Endemic/epidemic DENV cycles The most important DENV transmission cycle in terms of public health importance is that occurring in urban and periurban environments throughout the tropics (Fig. 6). In this cycle, DENV are transmitted among humans, which are both reservoir and amplification hosts, by the peridomestic Ae. aegypti mosquito. Other Aedes species, such as Ae. albopictus, and Ae. polynesiensis can serve as secondary vectors. This cycle, which has become endemic in many parts of the tropics and is responsible for periodic epidemics of various intensities, hereafter will be referred to as ‘endemic.’ The acquisition of Ae. aegypti as principal vector in urban settings resulted in the independence of the endemic cycle from the enzootic cycle for DENV maintenance. Although Ae. aegypti has an African origin (Christophers, 1960; Edwards, 1932), the establishment of extensive trade routes in the 1700s–1800s, the movement of people and water storage practices resulted in the widespread infestation of the tropics worldwide. As Leichtenstern initially and subsequently by others in the tropics recognized, DEN was a disease of ports and towns in coastal regions where the disease would travel inland along rivers (Barraud, 1928; Leichtenstern, 1896; More, 1904; Skae, 1902). Although none of these
History and Evolution of Human Dengue Emergence
27
early workers made an association between the disease and the mosquito vector, it was Stanton in his 1915–1916 mosquito survey of all the major seaports of Southeast Asia and Indonesia, who confirmed the presence of Stegomyia fasciata (as Ae. aegypti was known at the time) and the replacement of the native Ae. albopictus (Stanton, 1919; Theobald, 1901). Although Ae. albopictus is often more susceptible and thus a better laboratory vector of DENV (Gubler and Rosen, 1976; Jumali et al., 1979; Moncayo et al., 2004; Rosen et al., 1985) and under certain circumstances a better vector in nature (Hotta, 1952; Metselaar et al., 1980; Qiu et al., 1981), the adaptation of Ae. aegypti to domestic habitats and its feeding behavior allows it to surpass in epidemiological importance all other species of Aedes mosquitoes. Ae. aegypti have become highly domesticated and lay their eggs in artificial water containers commonly found in domestic habitats, such as flower pots, rainwater collection buckets, large water storage cisternae, and discarded tires, producing large number of adult mosquitoes in close proximity to humans. The adults are almost exclusively anthropophilic, prefer to feed during two peaks (early morning and afternoon) and afterwards prefer to rest on indoor walls, where they remain unobtrusive. However, being nervous feeders they interrupt their feeding process at the slightest human movement, only to return moments later to the same or a different host. Thus, in the process of obtaining a single blood meal (even if only probing) they can transmit DENV to multiple hosts during a very short time span and within a single gonotrophic cycle (Platt et al., 1997; Putnam and Scott, 1995a,b). In essence, this feeding behavior overcompensates for Ae. aegypti’s limited oral susceptibility, which varies greatly in both Ae. albopictus and Ae. aegypti based on their geographic origin (Failloux et al., 2002; Gubler and Rosen, 1976; Gubler et al., 1979; Vazeille et al., 2001, 2003). The limited susceptibility of Ae. aegypti suggests that only DENV associated with high human viremia levels would be transmitted efficiently, thus selecting for viruses that generate high viremia titers. Because viremia is correlated with the severity of human disease (Guilarde et al., 2008; Libraty et al., 2002; Vaughn et al., 1997, 2000; Wang et al., 2003), mosquito infection may select for virulence of endemic DENV. Although the mechanism of maintenance of DENV during the dry season or during interepidemic periods has not been clearly elucidated, evidence suggests DENV maintenance by vertical (transovarial) transmission. For a long time it was believed that transovarial transmission (TOT) of arboviruses by mosquitoes was not possible based on negative experimental evidence (Siler et al., 1926; Simmons et al., 1931). This view was revised when experimental evidence demonstrated TOT of vesicular stomatitis virus (VSV) by phlebotomus flies (Tesh et al., 1972), and TOT of LaCrosse virus by Ae. triseriatus in nature (Watts et al., 1973). The first evidence of DENV transovarial transmission (TOT) in nature was
28
Nikos Vasilakis and Scott C. Weaver
demonstrated by the isolation of DENV-2 (presumably sylvatic strains) from a pool of male Ae. taylori in Coˆte d’Ivoire in 1980 (Roche et al., 1983) and a year later in an Ae. furcifer mosquito pool in Senegal (Cornet et al., 1984). At about the same time, DENV-2 (presumably an endemic strain) was also isolated from 3 out of 123 Ae. aegypti larvae pools collected in natural breeding containers located throughout Rangoon, Burma, as well as from 2 of 76 pools of male Ae. aegypti larvae that were reared to adults. The calculated minimum field infection rates were 1:2067 and 1:3865 respectively (Khin and Than, 1983). Low filial infection rates for Ae. aegypti seen in nature have also been observed in experimental studies (Rosen et al., 1983). Further evidence of TOT, was also obtained in Trinidad were DENV-4 was isolated in a pool of adult Ae. aegypti collected as eggs (Hull et al., 1984), and in India where TOT of DENV-3 was demonstrated in nature and experimentally ( Joshi et al., 1996). More recently, field-caught male Ae. aegypti mosquitoes from four diverse locations of Southern Thailand tested positive for both DENV-2 and DENV-3 infection (Thavara et al., 2006). Nonetheless, no evidence of TOT was reported in earlier extensive mosquito larvae surveys in Bangkok, Thailand (Watts et al., 1985). Dengue virus TOT has been also demonstrated in other Aedes mosquito species that play a role in the transmission of DENV in nature, such as Ae. albopictus (Mitchell and Miller, 1990; Rosen, 1988; Rosen et al., 1985), Ae. mediovittatus (Freier and Rosen, 1988), and several members of the Aedes (Stegomyia) scutellaris group, which are important in the transmission of DENV in the Indonesian archipelago and Polynesia (Freier and Rosen, 1987). However, TOT rates in Ae. albopictus exhibited extensive variability depending on the strain of virus as well as the geographic origin of the mosquitoes used (Rosen et al., 1985), which may explain the lack of positive pools in a study conducted in Malaysia (Lee et al., 1997). Interestingly, at least for Ae. albopictus, it has been demonstrated that TOT can be sustained for several generations for DENV-1 (Shroyer, 1990) and DENV-3 ( Joshi et al., 2002) in the absence of any horizontal transmission from vertebrate blood meals, a notion initially suggested by Rosen 15 years earlier (Rosen, 1987). At face value this phenomenon suggests that mosquitoes can act as DENV reservoirs. Collectively, the data suggest that TOT may play a significant role in the maintenance of DENV in nature when unfavorable conditions limit horizontal transmission. Such conditions may include periods when low populations of susceptible non-immune vertebrate hosts cannot sustain horizontal transmission (interepidemic period), or when climatic conditions are unfavorable for mosquito activity (dry season), during which even very low transovarial transmission rates could preserve the virus. Most importantly, vertical transmission of DENV implies that priority and emphasis should be
History and Evolution of Human Dengue Emergence
29
placed on sustainable programs for the control and elimination of immature mosquito stages, as well as their artificial larval habitats.
IV. EVOLUTION A. DENV evolutionary relationships—origins and emergence Gubler’s (Gubler, 1997) hypothesis that endemic DENV evolved from sylvatic progenitors has been the subject of intense discussion and speculation. As mentioned above, sylvatic DENV cycles occur in the forests of West Africa and Southeast Asia involving only the DENV-2 serotype in the former region, whereas in the latter all four serotypes may be represented. Virus maintenance in both regions involves sylvatic Aedes spp. mosquito vectors, and presumably non-human primates serving as reservoir hosts. These sylvatic cycles are representative of ancestral cycles from which the endemic strains (all four serotypes) are thought to have arisen independently several hundreds to thousands of years ago. In comparison, current tropical urban endemic cycles occur in evolutionarily independent and ecologically distinct environments through transmission of DENV between anthropophilic Aedes spp. mosquito vectors and humans serving as reservoir hosts. The first support for this hypothesis came from phylogenetic studies by Rico-Hesse, which demonstrated that sylvatic, West African DENV-2 strains are genetically distinct from endemic isolates (Rico-Hesse, 1990). Gubler’s hypothesis was tested by Wang et al. (Wang et al., 2000) by assembling phylogenetic trees derived from complete E protein gene sequences of sylvatic DENV-1, -2, and -4 strains of Southeast Asian origin (Rudnick and Lim, 1986), as well as DENV-2 sylvatic strains from West Africa (Cornet et al., 1984; Saluzzo et al., 1986b). Sylvatic DENV-3 are believed to circulate in Southeast Asia based on the seroconversion of sentinel non-human primates (Rudnick, 1986), although no virus isolation has been reported to date. The analyses indicated that the Southeast Asian endemic serotypes evolved independently from progenitor sylvatic DENV of each serotype, in a series of divergence events occurring after the establishment of sufficiently large urban populations in the AsiaOceania region capable to support a human transmission cycle (Figs. 2, 3, 5) (Gubler, 1997; Kuno, 1995). However, a more recent and robust analysis using complete DENV genomes does not place the Malaysian canopy DENV-1 isolates in a basal position within this serotype (Fig. 7) as was seen in the E gene tree (Fig. 3). If the P72-1244 DENV-1 strain actually represents an endemic strain that circulated in the Malaysian forest
30
Nikos Vasilakis and Scott C. Weaver
Sylvatic?
100
Mal/P72-1244/72 Thai/488/94 Thai/97/94 Sing/S275/90 Thai/102/91 99 Thai/49/2001 Thai/336/91 Djibouti/1998 Thai/323/91 Chin/Guangzhou/80 Thai/81/82 Thai/08/1981 Jap/Mochizuki/43 86 Indo/A88/1988 IvCoast/Abidjan/98 FrGuiana/89 Braz/90 Braz/111/97 100 Braz/233/97 Braz/MR/01 Par/259/00 Arg/297/00 Arg/293/00 Thai/673/80 Thai/442/80 Thai/1687/98 Thai/1283/98 Thai/104/93 Thai/55/93 Thai/07/87 Thai/10/87 Martin/99 SriL/00
Endemic/ epidemic
DENV-1
DENV-3 Sylvatic?
IC/A510/80 IC/A578/80 IC/A1247/80 BFaso/2039/80 BFaso/A2022/80 Guin/PM33974/81 Sen/75505/91 Sen/319/99 100 Sen/320/99 Nig/11208/66 Nig/11664/66 100 Nig/IBH11234/66 Sen/20761/74 Sen/A10674/70 Mal/P8-1407/70 Peru/1950/95 Peru/2913/96 Peru/1797/95 100 Ven/1319/92 Ven/2/87 PR/1328/77 BFaso/1349/82 Austral/TSV01/93 Chin/FJ11/99 Thai/74 100 Thai/38/74 Thai/NH73/93 Thai/NH36/93 Thai/NH55/93 Thai/17/98 Thai/78/01 Thai/C0167/96 Thai/263/95 Thai/K0010/94 100 Thai/55/99 Thai/26/88 Thai/433/85 Thai/NHp14/93 Thai/168/79 Thai/498/84 Thai/16681/64 NewGuin/C/44 China/44/87 China/43/89 Braz/64022/98 Ven/Mara4/90 Martin/98 Jam/N1409/83 China/04/85 Thai/284/90 DomRep/1981 Thai/734/00 Thai/348/91 100 Thai/485/01 Thai/87/77 100 Thai/476/97 Thai/17997 Mal/P75-215/75
Sylvatic
100
DENV-2
Endemic/ epidemic
10% nucleotide sequence divergence
Endemic/ epidemic Sylvatic
DENV-4
FIGURE 7 Evolutionary relationships of the 4 DENV serotypes. Phylogenetic tree derived from complete genome nucleotide sequences in the GenBank library. Dashed branches represent predicted ancestral, sylvatic lineages and solid branches represent the emergence of the endemic lineages. Bayesian probability values are shown for key nodes. The most recent common ancestor (MRCA) estimates are from previous publications (Holmes and Twiddy, 2003; Wang et al., 2000). Strains are abbreviated as follows: country abbreviation/strain/year.
during the studies of Rudnick et al., than the assumption of emergence of endemic DENV-1 from a sylvatic ancestor implies that, like DENV-3, the DENV-1 sylvatic cycle has not yet been sampled.
History and Evolution of Human Dengue Emergence
31
The hyperendemic DENV cycles seen today probably required further urbanization of the twentieth century combined with the distribution of the highly efficient vector, Ae. aegypti, throughout the tropics. Assuming that DENV evolves at a constant rate, an assumption that may not be completely valid (see rates of evolution section below), it was estimated that the endemic DENV-2 genotypes diverged from the sylvatic genotypes no more than 1000 500 years ago, DENV-4 no more than 600 300 years ago, and DENV-1 no more than 200 100 years ago (Fig. 7) (Wang et al., 2000). Within DENV-2, the African and Malaysian sylvatic lineages diverged from each other about 800 400 years ago, presumably when Asian viruses were introduced into Africa. Slightly more recent dates have been estimated for most recent common ancestors using maximum likelihood methods (Holmes and Twiddy, 2003). As the above estimated times of endemic DENV emergence indicate a series of recent events, it is estimated that the ancestor for all DENV most likely occurred much earlier and probably at a time when sylvatic DENV utilized only non-human primates as reservoirs (Wang et al., 2000). The geographic origin of DENV has been the subject of debate for some time. It has been suggested that DENV originated in Africa based on the circulation of many mosquito-borne flaviviruses (Fig. 1) and the origin of the most important vector (Ae. aegypti) for interhuman transmission (Gaunt et al., 2001). However, as indicated previously (see transmission cycles) Ae. aegypti has only recently (300–400 years ago) been adopted as a vector for human transmission, long after the evolution of the 4 DENV serotypes. Ecological and phylogenetic evidence argues for an Asian origin of DENV: (1) greater diversity of sylvatic serotypes (possibly all four) in Southeast Asia, whereas in Africa only circulation of sylvatic DENV-2 has been demonstrated (Cordellier et al., 1983; Roche et al., 1983; Rudnick, 1986); (2) phylogenetic analysis demonstrating the deep phylogenetic position of the Asian sylvatic strains (Twiddy et al., 2002; Wang et al., 2000). Nonetheless, conclusive determination of the geographic origin of DENV will require increased sampling of sylvatic strains from both Asia and Africa; currently only seven sylvatic strains of DENV-1, -2, and -4 from Southeast Asia are known to exist (see transmission cycles).
1. Venues for DENV emergence from sylvatic cycles
The studies of Wang et al. (Wang et al., 2000) suggested that emergence of the four endemic DENV serotypes was facilitated by vector switching, from enzootic arboreal Aedes mosquito species to peridomestic Ae. albopictus and other Aedes (i.e., Ae. polynesiensis) mosquitoes and human reservoir hosts, in a convergent process that occurred independently and
32
Nikos Vasilakis and Scott C. Weaver
repeatedly (Wang et al., 2000). The emergence of distinct DENV lineages (serotypes) was most likely facilitated by the allopatric and perhaps ecological partitioning of ancestral sylvatic DENV strains in different non-human primate populations. However, the studies of Rudnick in Malaysia demonstrated that sylvatic serotypes probably utilize similar non-human primate hosts (see transmission cycles), which suggests that if the DENV serotypes evolved allopatrically and later expanded into overlapping regions, that occurred after their divergence had reached adequate levels of strain variation to allow maintenance through only limited cross-reactive immune protection. Antibody-dependent immune enhancement (ADE), although it has not been demonstrated to occur in nonhuman primates, could select for the maintenance of sufficiently antigenically distinct serotypes such that each benefits in a cooperative manner from immunity generated due to the others (Ferguson et al., 1999). Alternatively, emergence of distinct DENV lineages may have occurred sympatrically, which would have produced antigenically similar lineages with complete immunological cross-protection. However, this scenario is not favorable for the emergence of distinct lineages, since strong cross-reactive immune protection leads to direct competition among lineages and may result in competitive exclusion of different strains that occupy the same ecological niche if host resources are limiting (Ferguson et al., 1999). However, if antigenically distinct DENV lineages evolved sympatrically, then one could argue that sustained virus transmission was maintained through ADE. Unlike other viruses that are subject to strong host immune pressures, such as influenza (Bush et al., 1999) or the human immunodeficiency virus (HIV) (Zanotto et al., 1999), the selective pressures acting upon DENV appear to be rather weak and located at least partly within putative T or B cell epitopes (Twiddy et al., 2002a,b), reflecting the strong purifying selection characteristic of many arboviruses ( Jenkins et al., 2002; Weaver et al., 1999). Furthermore the historical record indicates that ADE in humans is a relatively recent phenomenon that arose due to the contact of antigenically distinct DENV viruses coincident with hyperendemicity during the twentieth century. Overall the major selection pressure acting on DENV genomes is purifying (negative selection), manifested by the low ratio of nonsynonymous (dN) to synonymous (dS) substitutions per site (dN/dS << 1), with sporadic occurrences of positive selection (Holmes, 2003; Twiddy et al., 2002a,b). These selective pressures vary among serotypes, genotypes and viral proteins, and where positive selection has been identified, amino acid sites reside within the E glycoprotein and are implicated in virulence and transmissibility (Sanchez and Ruiz, 1996; Twiddy et al., 2002a,b), or in the nonstructural NS2B, NS3, and NS5 genes (Twiddy et al., 2002a,b). DENV phylogenetic analyses have also revealed a complex pattern of evolution within multiple lineages that is partly fueled by the extensive
History and Evolution of Human Dengue Emergence
33
travel movement of humans and mosquitoes (especially in the days of sailing ships). The advent of commercial air travel, facilitating the rapid movement of viremic humans, led to introduction of DENV into new locations resulting in the displacement or extinction of local lineages (Carrington et al., 2005; Diaz et al., 2006; Myat Thu et al., 2005; RicoHesse et al., 1997; Salda et al., 2005; Zhang et al., 2005). Several reasons have been proposed for these population shifts and lineage replacements, such as: (1) adaptive mutations in nonstructural proteins (Bennett et al., 2003), which has been observed in epidemics of other viruses (Knowles et al., 2001); (2) adaptive mutations in cytotoxic T cell (CTL) epitopes (Appanna et al., 2007; Hughes, 2001; Zivna et al., 2002); and (3) fitness for transmission which may differ among DENV genotypes, as is evidenced by the greater susceptibility of Ae. aegypti for the invading Asian genotypes than for the American genotype it has displaced in some locations (Armstrong and Rico-Hesse, 2003). Differences in replication in human DCs may also give the Asian genotype a competitive advantage (Cologna et al., 2005). Extinction and emergence of new virus lineages can also be attributed to natural selection, or to drift associated with genetic bottlenecks in the viral population size (Klungthong et al., 2004; Sittisombut et al., 1997; Wittke et al., 2002). For arboviruses, bottlenecks may occur during seasonal and spatial fluctuations in vector population sizes and densities during inter-epidemic years or following increases in herd immunity, which could lead to genetic drift in DENV (Zhang et al., 2005). Although virus population shifts and lineage replacements are temporally associated with changes of incidence and severity of DEN disease, and could be attributed to a number of factors, these events are thought to be mainly stochastic in nature (Diaz et al., 2006; Myat Thu et al., 2005).
2. Recombination Lastly, recombination may be an alternative force shaping DENV evolution. Genetic recombination between RNA viruses serves a dual, yet antithetical purpose; it allows exploration of novel combinations between genetic regions from different parental genomes, or facilitates the ‘rescue’ of viable genomes from mutational debilitation due to Muller’s ratchet and genetic load (Domingo and Holland, 1997; Lai, 1992a). Recombination has been described, albeit very rarely, in nonsegmented RNA viruses, including alphaviruses (Hahn et al., 1988), picornaviruses (Cooper et al., 1974) and other members of the Flaviviridae family (Becher et al., 2001; Worobey and Holmes, 2001). Evidence of recombination has been reported within all four DENV serotypes (Aaskov et al., 2007; AbuBakar et al., 2002; Craig et al., 2003; Domingo et al., 2006; Holmes et al., 1999; Tolou et al., 2001; Uzcategui et al., 2001; Worobey et al., 1999), but not between the serotypes. The putative recombination events described
34
Nikos Vasilakis and Scott C. Weaver
within DENV serotypes were detected only by utilizing computationally demanding phylogenetic analyses (split decomposition and/or maximum likelihood methods), and caution should be used when inferring conclusions about these putative recombination events. To date the most convincing evidence of DENV recombination was described in a recent report of multiple DENV-1 isolates from a single patient in New Caledonia (Aaskov et al., 2007). In this report putative DENV-1 recombinants were detected with both phylogenetic analyses and genetic confirmation of identical crossover breakpoints (Aaskov et al., 2007). However, because viable clonal recombinant viruses are rarely observed in nature, the implications of the recombinant genomes observed in this report are uncertain. For natural recombination leading to the transmission of a recombinant strain to be conclusively confirmed, the following prerequisites should be met: (1) the recombinant crossover should be demonstrated in a single PCR amplicon following cloning to ensure it occurs in a single DNA molecule; (2) the recombination should be demonstrated repeatedly in clonal populations of viable virus (e.g., a plaque harvest or limited endpoint dilution; and (3) the recombinant should maintain adequate sequence conservation during post-recombination evolution (Tolou et al., 2001). This caution of interpreting evidence of recombination based on phylogenetic analyses does not exclude the possibility that recombination in DENV can occur. If DENV recombination occurs, it most likely involves a copy-choice mechanism where the polymerase ‘switches’ between viral genomes during the replicative process (Cooper et al., 1974; Jarvis and Kirkegaard, 1992; Lai, 1992b). Taking into account the feeding behavior of Ae. aegypti, as well as the detection of simultaneous, multiple DENV infections in the vertebrate host (Araujo et al., 2006; Gubler et al., 1985a) (albeit with strains of different serotypes), or in the mosquito vector (Craig et al., 2003; Lorono-Pino et al., 1999), ample opportunities for recombination are present. The possibility of DENV recombination may have significant epidemiological and clinical implications in endemic regions if licensure of the attenuated multivalent DENV candidate vaccines results in widespread human use. Overall, phylogenetic analyses indicated that the Southeast Asian endemic DENV serotypes evolved independently from progenitor sylvatic DENV in a series of convergence events after the establishment of sufficiently large urban populations in the Asia-Oceania region capable of supporting a human transmission cycle. On the other hand, the divergence of sylvatic Southeast Asian and African DENV-2 lineages is a more recent event. The Asiatic origin of DENV sylvatic progenitors is further supported by serological surveys of ecologically diverse rural habitats in Southeast Asia, which may also suggest that the emergence of the endemic lineages was facilitated by vector switching (anthropophilic
History and Evolution of Human Dengue Emergence
35
mosquitoes). The explosive emergence of new DENV lineages and hyperendemic DEN can be attributed to the explosive growth of human travel after the end of World War II, as well as the colonization of most tropical regions by the highly efficient vector, Ae. aegypti.
B. Rates of DENV evolution The unique requirement of arthropod-borne RNA viruses (arboviruses) for replication in divergent hosts may impose additional selective constraints compared to single-host virus cycles. Because arboviruses utilize both vertebrate and invertebrate hosts, they have adapted to replicate efficiently in either host, where optimal replication in one host involves a fitness tradeoff for the alternate host (Cooper and Scott, 2001; Greene et al., 2005; Weaver et al., 1992, 1999). DENV sequence comparisons suggest the presence of strong purifying selection in nature, supporting strong selective constraints on arbovirus evolution (Holmes, 2003; Klungthong et al., 2004). However, some experimental evidence does not support the notion that alternating host replication produces fitness trade-offs in arboviruses (Novella et al., 1999). Reconstruction of a molecular time-scale evolution of DENV by estimation of the nucleotide substitution rates between viruses sampled at different times and locations can provide valuable insights into the history of DENV (Rambaut, 2000). As described above, the prevalence of all DENV serotypes worldwide increased dramatically after the end of World War II, a rise accompanied by increased genetic diversity within each serotype, leading to the emergence of strains with increased pathogenicity. Therefore, utilizing tools that allow us to gain insights into the epidemiological potential, as well as the processes that drive DENV evolution could help us develop effective control countermeasures. Initial evaluations on the rate of DENV evolution were based on pairwise comparisons of phylogenetically divergent, yet related E genes (sister strains), where the divergence (number of nonsynonymous substitutions per nonsynonymous site) occurring between their sampling time was plotted against the difference between their time of isolation (Zanotto et al., 1996). The rate of evolution for DENV-1 was estimated at 4.4 104, for DENV-4 at 8.5 104, for DENV-3 at 2.9 10–4–6.8 104, and for DENV-2 at 1.6 104–7.5 105 substitutions/site/year. Using these rates, they estimated the divergence from sylvatic ancestors (or the age of the most recent common ancestor [MRCA]) at 1500–2000 years ago. A few years later, the rate of evolution of DENV-4 was estimated at 8.3 104 subs/site/year by employing linear regression analysis (Lanciotti et al., 1997), a rate that is similar to the estimated rate described in the previous study. Wang et al. employed both methods described in the two previous studies to estimate a rate of 5 104 subs/site/year for DENV-2
36
Nikos Vasilakis and Scott C. Weaver
(Wang et al., 2000). They estimated the earliest endemic serotype to diverge was DENV-2 at approximately 1000 500 years ago, followed by DENV-4 and DENV-1 at 600 300 and 200 100 years ago, respectively. Using similar methods, Goncalvez et al. estimated a rate of 5.8 104 subs/site/year for DENV-1, and a very recent divergence (approximately 100 years ago). However, the methods of rate estimation employed in these studies were not statistically rigorous because they either utilized limited data sets for pairwise comparisons, had biases in sampling times or used an untested assumption of a molecular clock or a constant rate of evolution (Twiddy et al., 2003). These limitations were addressed by the development of a maximum likelihood (ML) method that takes into account the times of sampling of a large set of E gene sequences to generate appropriate confidence intervals around the generated rates of evolution and times of divergence and allows the use of complex models of nucleotide substitution (Twiddy et al., 2003). They utilized the following criteria to assemble their dataset: (1) known date of virus sampling, (2) use of single sequence from any patient, (3) exclusion of putative recombinants, and (4) exclusion of sequences that shared >99% nucleotide identity to any other sequence in the set. Estimated rates of endemic DENV-1, -2, -3, and -4 were 4.5 104, 6.1 104, 9.0 104, and 6.02 104 subs/site/year, respectively. These rates were very similar to those estimated for DENV in a comprehensive study of several RNA viruses (Jenkins et al., 2002). Their analyses also revealed that DENV evolution conforms to a molecular clock, although some serotype and genotype-specific rate differences were observed. Interestingly, the estimated times of DENV divergence were significantly different from previous analyses (Wang et al., 2000; Zanotto et al., 1996), placing the emergence of human epidemic DENV transmission at more recent times (approximately 300 years ago) (Twiddy et al., 2003). A more recent analysis of DENV-4 with an extensive dataset of temporal sampling and employing similar methods as described in the previous study estimated a rate of 1.1 103 subs/site/year (Klungthong et al., 2004). It is not known whether this considerably higher rate of DENV-4 evolution compared to the rate generated from previous analyses (Lanciotti et al., 1997; Twiddy et al., 2003), reflects a high DENV-4 turnover in Thailand, or is a consequence of the extensive sample size. However, if one considers the strong purifying selection exerted on DENV and frequent lineage extinction due to the natural occurrence of deleterious mutations, then it is possible that all estimated rates of DENV evolution are overestimates because these sequences have not been removed from the data set (Klungthong et al., 2004; Pybus et al., 2007). Until recently, the study of Twiddy et al. was the only on that attempted to evaluate the evolutionary rates of sylvatic DENV within a large dataset of endemic DENV (Twiddy et al., 2003). Although their data
History and Evolution of Human Dengue Emergence
37
suggested that inclusion of sylvatic DENV in the analyses did not alter the overall rate estimates, these inferences may have been affected by the small sample size of sylvatic DENV. Although, a very limited number of sylvatic DENV-1 (n ¼ 1), DENV-2 (n ¼ 4) and DENV-4 (n ¼ 2) samples was available, the existence of over 300 putative sylvatic DENV2 samples, collected mainly from amplifications in West Africa, offers a unique opportunity for elucidating the nature of evolutionary processes that characterize sylvatic DENV. Our recent analysis of 14 complete coding regions of sylvatic DENV-2 virus sampled over a 33-year period in West Africa and Southeast Asia demonstrated that both the rate of evolutionary change and nature of natural selection are similar among endemic and sylvatic DENV lineages, although the latter has a uniquely high frequency of positive selection in the NS4B protein gene (Vasilakis et al., 2007a). These analyses are the first to imply that the dynamics of mutation, replication and selection are similar for DENV-2 across its host range and suggest of rapid sylvatic DENV turnover (rapid generation of viral diversity) due to their high nucleotide substitution rates. Surprisingly, the large virus population sizes of endemic DENV-2 associated with hyperendemicity do not appear to have major impacts on evolutionary rates.
C. Evolution of virulence 1. DEN animal models A major limitation in DENV research is the lack of suitable animal or in vitro models that recapitulate human disease and viremia. Consequently, identifying the mechanisms of pathogenicity, as well as characterizing viral determinants of virulence (which for DENV infections implies progression into DHF) has been difficult. Various non-human primates support DENV replication without developing clinical signs of illness (Halstead et al., 1973a; Rosen, 1958a), although the duration and magnitude of virus replication often correlates with patterns of replication in humans. For this reason, primates have become the standard for evaluation of live attenuated DENV vaccine candidates (Angsubhakorn et al., 1988; Blaney et al., 2005; Edelman et al., 1994; Hanley et al., 2004; Markoff et al., 2002; Men et al., 1996; Robert Putnak et al., 2005). Murine models (Boonpucknavig et al., 1981; Chaturvedi et al., 1991; Cole and Wisseman, 1969; Hotta et al., 1981) have also been developed, but have proven ineffective due to the absence of natural DENV strain replication, illness, and the resulting requirement for murine-adapted DENV strains for use in these animals (Cole and Wisseman, 1969; Sabin, 1952; Shresta et al., 2006). The development of the SCID-xenograft model (intraperitoneally grafted HepG2 or Huh-7 cells) has provided a model where DENV replication is similar to that which accompanies human infection (An et al., 1999) and proved useful for virulence testing of DENV vaccine
38
A
Nikos Vasilakis and Scott C. Weaver
In vivo (SCID-xenograft model)
B
Ex vivo (monocyte-derived DCs [mo-DCs]) Obtain blood from consenting healthy volunteers
Separate PBMCs and isolate CD14+ monocytes by MACS
Transplant 1⫻107 Huh-7 cells i.p. ~4–6 wks
IL-4 + GM-CSF selection of CD14+ monocytes 6–7 days 200
Counts
Development of peritoneal tumors
160
Isotype
120
DC-SIGN
80
CD83
40 0 0 10
1 103 102 10 Mouse IgG2a FITC
Confirm immature status of mo-DCs by FACS
104
Infect immature mo-DCs with DENV (MOI= 2) Inject 1⫻104 virus directly into tumor 2 days 7 days Collect serum, isolate and characterize virus in tumor, liver, brain
Collect supernatant and quantify virus output
FIGURE 8 Surrogate human models of DENV used to estimate the infection phenotypes of sylvatic versus endemic DENV-2 strains. (A) SCID-xenograft model. (B) monocyte derived dendritic cell (moDC) model.
candidates (Fig. 8A) (Blaney et al., 2002; Whitehead et al., 2003). Similarly, a humanized mouse model based on the grafting of human CD34þ cells in non-obese diabetic/severely compromised immunodeficient mice (NOD/SCID), has also been proposed as a model for studying the pathogenicity of DENV infection (Bente et al., 2005). A number of vertebrate and invertebrate cells are permissive for DENV replication but their relevance to association with human disease and virulence is questionable. One limitation for the establishment of in vitro models of virulence lies in our lack of understanding of the target cells for infection in humans. For many years, mononuclear phagocytes (macrophages and/or monocytes) were presumed to be the target cells (Halstead, 1989; Halstead and O’Rourke, 1977a,b; Halstead et al., 1977, 1980; Morens, 1994). However, subsequent in vitro studies employing sensitive methods of viral RNA and antigen detection repeatedly demonstrated that human monocytes and/or macrophages are either minimally
History and Evolution of Human Dengue Emergence
39
infected with DENV (Blackley et al., 2007; Bosch et al., 2002; Diamond et al., 2000; Goncalvez et al., 2007; O’Sullivan and Killen, 1994; Wu et al., 2000), or readily infected in the presence of DENV-specific antibodies (Brandt et al., 1982; Diamond et al., 2000; Goncalvez et al., 2007; King et al., 2002; Kliks et al., 1989; Kou et al., 2008). Several other non-myeloid, human cell lines such as endothelial (Anderson et al., 1992; Avirutnan et al., 1998; Bonner and O’Sullivan, 1998; Huang et al., 2000), epithelial (Bosch et al., 2002), fibroblasts (Diamond et al., 2000; Kurane et al., 1992), and hepatocytes (Hilgard and Stockert, 2000; Marianneau et al., 1996, 1997) have been shown to be readily infected with DENV. More recently human skin dendritic cells (DCs) and monocyte-derived dendritic cells (moDCs) have been demonstrated to be permissive for DENV infection and replication in the absence of DENV-specific antibodies (Tassaneetrithep et al., 2003; Wu et al., 2000). This work suggests that skin DCs (i.e., Langerhans cells) are the initial target of DENV infection following mosquito transmission, and could be used in assessing important biological differences among viral genotypes and their contribution to pathogenesis (Fig. 8B) (Cologna and Rico-Hesse, 2003; Cologna et al., 2005; Vasilakis et al., 2007b). A limitation of any model that is often overlooked in the context of DENV evolution of virulence is the passage history of the sample virus. It is well documented that virus cultivation in vivo (i.e., mosquitoes or rodents)(Morimoto et al., 1998; Schlesinger et al., 1996) or in vitro results in altered viral phenotypes (Bernard et al., 2000; Byrnes and Griffin, 2000; Lee et al., 2004, 2006). Therefore use of primary viral isolates or viruses rescued from infectious clones to eliminate passages is essential when attempting to elucidate the mechanisms or evolution of DENV virulence. Unfortunately, a major limitation of the availability of primary isolates is the requirement of patient samples with high viral titers from the acute phase of the disease, which occurs before the onset of severe symptoms.
2. DENV virulence One of the proposed mechanisms of DENV virulence-antibody-dependent enhancement (ADE) is based on epidemiological and experimental (both in vitro and in vivo) observations where secondary infection with a heterotypic DENV is often associated with increased risk in developing DHF due to existence of non-neutralizing heterologous antibodies (Halstead, 1979; Halstead et al., 1976; Kliks et al., 1989). Other proposed mechanisms examined the role of host immune factors, such as cytokines (Bosch et al., 2002; Cardier et al., 2006; Chakravarti and Kumaria, 2006; Chen et al., 2006; Hober et al., 1993; Kurane et al., 1991, 1993; Libraty et al., 2001), coagulation abnormalities (Wills et al., 2002), genetic predisposition (Bravo et al., 1987), gender and age (Guzman et al., 1984, 2002; Halstead
40
Nikos Vasilakis and Scott C. Weaver
et al., 2001), and nutrition (Kalayanarooj and Nimmannitya, 2005; Thisyakorn and Nimmannitya, 1993). Epidemiological reports and genetic studies have also indicated that viral factors (i.e., specific genotypes and/or serotypes, viral structures, etc) may be important indicators of DENV virulence. Although no clear correlation has been established between specific genotypes or serotypes and virulence, epidemiological and phylogenetic evidence suggests that certain Asian DENV-2 genotypes are more virulent than American genotypes or those circulating in the South Pacific (Fig. 3) (Harris et al., 2000; Leitmeyer et al., 1999; Murgue et al., 2000; Rico-Hesse et al., 1997; Rosen, 1977; Watts et al., 1999). Similarly DENV-3 genotype III, which belongs to a genetically distinct clade (Fig. 4), has been responsible for geographically distant epidemics of DHF in the Indian subcontinent where it first originated, East Africa (in the 1980s), and Latin America (in the 1990s) (Messer et al., 2003). In the context of DENV-2, subsequent studies have indicated that Asian genotypes replicate to higher viral outputs than American genotypes in monocyte-derived DCs (moDCs) (Cologna and Rico-Hesse, 2003; Cologna et al., 2005; Vasilakis et al., 2007b) or macrophages (Pryor et al., 2001). Complete genome analysis and comparisons between Asian and American genotypes revealed numerous differences at the nucleotide and amino acid levels within the open reading frame, as well as predicted RNA structural differences in the 30 noncoding regions (NCR) (Leitmeyer et al., 1999)(Vasilakis et al., 2008c). Of particular interest is a particular amino acid difference at position 390 of the E protein, which has been suggested to be a critically determinant of virulence. Residue 390 is located within the putative glycosaminoglycan binding motif (386L-411M) responsible for the binding of DENV to the host cell via a non-Fc receptor (Chen et al., 1996). This residue shows extensive polymorphism among DENV-2 genotypes: for ancestral sylvatic and Asian genotypes residue 390 is an Asparagine (N), for the majority of cosmopolitan genotypes is a serine (S)[a few retain the ancestral N], whereas for American genotypes is an aspartic acid (D) [N390D] (Twiddy et al., 2002)(Vasilakis et al., 2008c). Phylogenetic analyses have shown that residue N390 is under positive selective pressure (Twiddy et al., 2002), whereas D390 has been shown to reduce virus production by both human monocyte-derived macrophages (Pryor et al., 2001) and moDCs (Cologna and Rico-Hesse, 2003), as well as to alter virulence for mice (Sanchez and Ruiz, 1996). The putative attenuating property of D390 is further enhanced in chimeras where the Asian genotype 30 -NCR is replaced with the American genotype 30 -NCR (Cologna and Rico-Hesse, 2003), suggesting a synergistic function. In summary, these observations suggest that the amino acid residue at E-390 may play an important role in determining key aspects of DENV phenotype, although further investigation is needed.
History and Evolution of Human Dengue Emergence
3. Virulence changes associated with DEN emergence from sylvatic cycles
41
In a recent study, the in vivo and ex vivo replication profiles of representative American genotypes were shown to be similar to those of the ancestral sylvatic DENV-2 genotypes (Vasilakis et al., 2007b), which are also considered of low virulence potential (Monlun et al., 1992; Robin et al., 1980; Saluzzo et al., 1986b; Vasilakis et al., 2008b). This observation suggests that American genotypes have retained their ancestral, lowvirulence phenotype. This finding implies that DENV-2 virulence increased recently, possibly the result of selection by Ae. aegypti, which requires higher viremia titers than ancestral vectors for efficient transmission (Gubler, 1997). The correlation between virulence and viremia, which facilitates efficient transmission, as well as the greater infectivity of the Asian strains for Ae. aegypti, predicts that American genotypes will exhibit poor transmission potential and be at a competitive disadvantage compared to other DENV-2 genotypes (Armstrong and Rico-Hesse, 2001; Cologna et al., 2005). During the 1995 DENV-2 epidemic in the Peruvian Amazon region, caused by an American genotype strain, 60.5% of students aged 7–20 years experienced secondary infection. By extrapolation, a minimum of 887 cases of DHF would have been expected, yet no DHF/ DSS cases were reported despite of thousands of DF cases (Watts et al., 1999). This study provided strong epidemiological evidence of the limited pathogenicity of the American genotype, even during secondary infection. Because the Asian genotypes had not yet reached this region of Peru, their possible competitive advantage for viremia and mosquito infection could not be evaluated. Other correlates of virulence include viral load (as reflected in viremia) and viral proteins (i.e., soluble nonstructural protein 1 [sNS1]) in the bloodstream of the infected. Early studies by Halstead (Halstead, 1988; Rosen, 1986) hypothesized an association between virulence (disease severity) and virus load. Several subsequent studies in southeast Asia (Avirutnan et al., 2006; Endy et al., 2004; Libraty et al., 2002; Vaughn et al., 2000; Wang et al., 2003, 2006) and Oceania (Murgue et al., 2000) have provided support for this hypothesis. However, contradictory evidence from other DENV epidemics in Asia (Chen et al., 2005; Sudiro et al., 2001; Yeh et al., 2006) is also found in the literature. Other viral proteins have also been shown to exhibit an indirect role in DENV virulence. Evidence suggests that NS2A, NS4A, NS4B, and NS5 are involved in the inhibition of signal transduction pathways from interferon (IFN) receptors, thus facilitating evasion from the host’s immune responses and allowing viral spread and replication (Appanna et al., 2007; Ho et al., 2005; Jones et al., 2005; Munoz-Jordan et al., 2003, 2005). Collectively, the contradictory lines of evidence indicate that elucidation of the causative pathways, genotypes and genes responsible for
42
Nikos Vasilakis and Scott C. Weaver
DENV virulence in humans will be quite complex. Similarly complex have become the virulence and disease models that include primary human cell cultures (moDCs), mosquitoes, xenografted severe combined immunodeficient or IFN knock-out mice, and viral infectious clones.
V. POTENTIAL FOR SYLVATIC DENV REEMERGENCE A. Epidemics and human contact Smith (Smith, 1956) and Rudnick (Rudnick, 1986), based on their studies of DENV ecology and seroprevalence in the Malay peninsula, hypothesized that the rural population, especially those living in close proximity to forested and rubber plantation areas, were exposed periodically to sylvatic DENV. As described above in the ‘transmission cycles’ section, all of the sylvatic DENV isolates were obtained from sentinel monkeys or Ae. niveus species mosquito pools, collected within sylvan environs (Rudnick, 1986). To date, the paucity of ecological and non-human primate epidemiological data in Southeast Asia do not allow the formulation of meaningful inferences about the vector–host interactions of sylvatic DENV. However, in West Africa, several sylvatic DENV amplification cycles have been documented since the early 1980s (Cornet, 1993; Diallo et al., 2003; Saluzzo et al., 1986b), but the degree of human contact during these events remains largely unknown. There has been no evidence that sylvatic amplification cycles are involved in major outbreaks of human DEN, which involve the genetically and ecologically distinct endemic strains. The available data suggest that the sylvatic strains are either confined to forest habitats and/or produce relatively mild illness (DEN fever). Most of our current understanding of human illness after infection with sylvatic DENV comes from the case histories of DENV infections in two Senegalese and three expatriate Caucasian patients, which led to the isolation and genetic characterization of DENV-2. The first case was described in 1970, in a 6 year old Senegalese girl living in the prefecture of Bandia; although this patient was parasitemic with Plasmodium falciparum, a blood sample yielded the DakAr HD10674 strain of DENV-2 (Robin et al., 1980). The second case involved a Caucasian scientist whose infection occurred while investigating the 1983 DENV amplification cycle in southeastern Senegal. The clinical illness was characterized as severe, with persistent rash and arthralgia lasting for approximately a year. The third case, also in 1983, involved a Caucasian male who, upon return from the southwestern Senegalese province of Casamance, developed a febrile illness characterized by classical DF signs and symptoms including a maculopapular rash on the fifth day after onset (Saluzzo et al., 1986b). The next two cases occurred during the 1990 epizootic outbreak in the
History and Evolution of Human Dengue Emergence
43
southeastern Senegalese province of Kedougou, and led to the isolation of DENV from humans concomitant with isolation from sylvatic Aedes mosquito species, suggesting the presence of a sylvatic cycle (Monlun et al., 1992). The fourth case involved a 31 year-old Caucasian man returning to Dakar from military maneuvers; two days after his return he developed a flu-like illness, characterized by sudden onset of classic DF signs and symptoms, including high fever, frontal headache, arthralgia, myalgia, emesis, and generalized asthenia. His blood indicators were within the normal range and he exhibited no signs of hepatomegaly or splenomegaly, neurological syndrome or rash. Serological analyses performed 77 days after the onset of symptoms indicated past exposure to WNV, ZIKV and WESSV viruses (Monlun et al., 1992). The last case involved a local 15 year-old Senegalese boy presenting near Kedougou with a malaria-like disease. He developed mild arthralgia that lasted nearly a month without any other symptoms. Recent phylogenetic evidence suggests that human contact with sylvatic DENV also took place in Ibadan, Nigeria during 4 years of DEN activity in the 1960s (Carey et al., 1971; Vasilakis et al., 2008b). From 1964– 1968, 14 of the 32 strains of DENV isolated from febrile patients were classified as DENV-2, of which 10 strains were isolated in 1966. Phylogenetic analysis that included the complete genomic sequences of 3 of the 1966 DENV-2 isolates (the only ones known to exist in reference collections) indicated that they were genetically distinct from endemic DENV-2 isolates, and fell within the sylvatic DENV-2 clade (Vasilakis et al., 2008b). Although there are no written records on the above patients’ locations of residence or places of exposure, all resided within the Ibadan city limits. Furthermore, the clinical presentation of DENV infection due to these Nigerian sylvatic DENV-2 strains was indistinguishable from classic DF due to endemic DENV (D. E. Carey, personal communication). These findings extend the temporal and spatial span in which sylvatic DENV are known to circulate in West Africa and provide further evidence that the West African DENV-2 strains can cause typical DF. Furthermore, a retrospective serological survey provided evidence that non-human primates are involved in the transmission cycle of sylvatic DENV in Nigeria (Fagbami et al., 1977). In that study, 38% of sera collected from non-human primates within the rainforest were positive for DENV antibodies, which was similar to levels observed (43%) for humans living in communities within the forest. The prevalence of DENV antibodies in non-human primates living within the forest gallery in the Nupeko forest, a rainforest preserve located along the upper middle Niger river, were the highest at 74% (Fagbami et al., 1977). However, the availability of limited epidemiological information and the paucity of serosurvey data prevent an accurate assessment of the overall human exposure to sylvatic DENV-2 in Nigeria, Senegal and other parts of West Africa.
44
Nikos Vasilakis and Scott C. Weaver
Even though sylvatic transmission cycles occupy ecologically distinct environments, the studies of Smith and Rudnick in Southeast Asia and the small number of documented human infections from Senegal and Nigeria suggest that sylvatic DENV comes into regular contact with humans, but with little or no secondary transmission within the human population (spillover epidemics). The mechanisms for the apparent inability of sylvatic DENV strains to regularly establish transmission among humans are not known, but perhaps could be attributed to their limited genetic diversity, lower pathogenicity potential, limited frequency or intensity of transmission. Another possible explanation for the confinement of sylvatic DENV strains to the forest is that they generally do not contact the peridomestic vectors, Ae. aegypti and Ae. albopictus, which are not abundant in regions where sylvatic DENV circulate. However, recent studies have shown that the gallery forest-dwelling mosquito Ae. furcifer is highly susceptible to sylvatic DENV infection (Diallo et al., 2005), and disperses from the forest into villages in eastern Senegal (Diallo et al., 2003). This suggests that this species may act as a bridge vector for exchange between forest and peridomestic habitats. Furthermore, the ability of Ae. aegypti and Ae. albopictus to transmit sylvatic DENV (Diallo et al., 2005), as well as the lack of evidence that any adaptation of sylvatic DENV is needed to replicate efficiently in humans (Vasilakis et al., 2007b), suggests that the transfer between forest and human habitats could occur regularly.
B. The influence of natural immunity or vaccination on potential sylvatic DENV emergence As described in previous sections (see DENV evolutionary relationships—origin and emergence) the four DENV serotypes have evolved from a common ancestral progenitor resulting in viruses sharing several common antigenic sites. However our current understanding of antibody-mediated protection (i.e., virus neutralization) against DENV infection, albeit epidemiologically well-founded, lacks an inherent understanding of human antibody responses during primary or heterotypic DENV infections. This gap is mainly attributed to our lack of knowledge of the precise identity of cells that support DENV replication, as well as their contribution to disease during the early events of DENV infection. Subsequently very little is known about any biologically relevant receptor(s) responsible for attachment and entry of DENV into human cells. Among the molecules that have been suggested as primary receptors for DENV are dendritic cell-specific ICAM-grabbing non-integrin (DCSIGN) (Navarro-Sanchez et al., 2003; Tassaneetrithep et al., 2003), glucoseregulating protein 78 (GRP78/BiP) ( Jindadamrongwech et al., 2004), and CD14-associated molecules (Chen et al., 1999). Glycosaminoglycans, such
History and Evolution of Human Dengue Emergence
45
as heparin, and its structural analogues have also been proposed as attractive candidates for DENV attachment to target cells due to their widespread distribution on cells, involvement in ligand recognition as well as signaling processes (Chen et al., 1997; Germi et al., 2002; Hung et al., 1999, 2004; Lin et al., 2002; Martinez-Barragan and del Angel, 2001; Pattnaik et al., 2007). There is ample epidemiological evidence indicating long lasting (lifelong) homotypic protective immunity (Papaevangelou and Halstead, 1977). Studies by Sabin on military recruits during World War II demonstrated failure to develop viremia in challenges with homotypic DENV strains (Sabin, 1952). Similarly during the 1981 Cuban DENV-2 epidemic, older Cubans were protected against infection because of their prior exposure to DENV-2 during the early 1940s (Guzman et al., 1990). However, a different story emerges with heterotypic infections. Sabin demonstrated in human volunteers that heterotypic infection within two months of primary infection offers complete protection from development of clinical illness (Sabin, 1952). Subsequently, protection declined gradually for the following months, as challenges with heterotypic DENV produced malaise and slight fever, and cross challenges 9 months post primary exposure led to the development of classic DF (Sabin, 1952). Although detailed studies in non-human primates are limited, similar responses are elicited in these animals. A study by Halstead with Macaca mulatta monkeys demonstrated that all naı¨ve primates developed viremia following infection with DENV-1 to -4, had developed neutralizing antibodies and thus were protected against challenge by homotypic virus (Halstead et al., 1973a). Upon heterotypic infection there was evidence of crossprotection within two weeks from primary exposure, however a number of non-human primates developed viremia characterized by significantly delayed onset, shortened duration and depressed peak viremia titers (Halstead et al., 1973b). Our knowledge of the neutralization potential of sylvatic strains by prior immunity to endemic strains is limited and has not been examined by organized research. Even though sylvatic transmission cycles occupy ecologically distinct environments, the studies of Smith (Smith, 1956) and Rudnick (Rudnick, 1986; Rudnick and Lim, 1986) in Southeast Asia and the small number of documented human infections from Senegal (Robin et al., 1980; Saluzzo et al., 1986b) and Nigeria (Carey et al., 1971; Vasilakis et al., 2008b) suggest that sylvatic DENV comes into regular contact with humans. Recent reports have also shown that the forest mosquito Ae. furcifer is highly susceptible to sylvatic DENV infection (Diallo et al., 2005), and has been shown a pattern of movement into villages in eastern Senegal (Diallo et al., 2003), which suggests that may act as a bridge vector for exchange of DENV between sylvatic and peridomestic habitats. Furthermore, since little or no change in vector infectivity accompanied
46
Nikos Vasilakis and Scott C. Weaver
endemic emergence from sylvatic progenitors (Diallo et al., 2005), the ability of sylvatic DENV-2 to replicate efficiently in humans (Vasilakis et al., 2007b), as well as the rapid sylvatic DENV turnover due to their high nucleotide substitution rates (Vasilakis et al., 2007a), suggest that reemergence of sylvatic DENV into the endemic cycle could occur at any time, which on the short term could affect public health by fueling human epidemics in areas with minimal or non-existent public health infrastructure. On a longer timescale, emergence of sylvatic DENV into the human transmission cycle could limit the potential for eradicating this transmission cycle with effective vaccines now under development. In a recent study the likelihood of current strains of sylvatic DENV to re-emerge in the face of immunity to endemic strains was assessed by evaluating the neutralization capacity of sera from DENV vaccinees and convalescent patients after primary infection with DENV-2 and -3 serotypes (Vasilakis et al., 2008a). One limitation of this study was the scarce availability of primary convalescent sera, which prohibited evaluation of their neutralization capacity with a larger collection of DENV. Nonetheless, the data indicated robust homotypic cross-immunity between human sera and sylvatic DENV strains, but limited heterotypic neutralization. A possible explanation for this observation is that the sera of DENV vaccinated subjects were collected 42 days post vaccination; this timeframe is too short for the development of long lasting homotypic humoral immunity, and is in line with previous observations where DENV infection to any serotype produces a short-lived heterotypic response, lasting up to 12 weeks (Sabin, 1952). On the other hand, the strong homotypic virus neutralization demonstrated by the sera of convalescent patients after primary infection, could explain the periodicity of sylvatic amplification cycles and absence of epidemics in the human population. This implies that sylvatic amplification cycles are limited by the effects of herd immunity within the nonhuman reservoir hosts and thus reduce the amplification of sylvatic DENV in the enzootic cycle, which ultimately reduces the chance of incidental infections in humans by bridge vectors (i.e., Ae. furcifer). Consequently, sylvatic amplification cycles occur when the population of non-immune non-human primate reservoir hosts rises to sufficient levels to support a transmission cycle. Therefore, should a licensed vaccine lead to the eradication of the endemic transmission cycle in the future, re-emergence of sylvatic strains into the endemic cycle would be limited by homotypic immunity mediated by virus neutralizing antibodies. However, as the experience of previous vector eradication efforts in the Americas can attest, abatement of vector control programs could send any eradication efforts into obsolescence. Similarly, the recent experience of polio eradication in West Africa suggests that abatement of blanket vaccination efforts can lead to the re-introduction of virus into areas where the virus was eradicated. Therefore, reduction and ultimately
History and Evolution of Human Dengue Emergence
47
eradication of DEN from human populations solely due to vaccination campaigns may be at best short-lived and unrealistic in the long term, if sylvatic DENV can readily re-emerge from sylvatic cycles not amenable to intervention. However, success could hinge on universal vaccination coverage of the susceptible population at risk and on the virtual elimination of the endemic vector mosquitoes as a result of sustainable vector control programs. Active employment of these methods should prevent the re-emergence of sylvatic DENV into the human transmission cycle and thus eliminate or dramatically reduce a major public health problem. Nonetheless, cessation of these public health measures for a sufficiently long time will lead to the rise of susceptible human populations and establishment of populations of endemic vector mosquitoes, which can serve as a platform for the re-emergence of disease-producing sylvatic DENV into the human population.
C. Selection pressures Several years ago Gubler questioned whether some DENV genotypes have a greater epidemic potential than others, thus rendering them good candidates for spawning epidemics when introduced in new geographic locations (Gubler et al., 1981). In other words, is there a selective basis for the differences in geographical distribution that is observed among the DENV genotypes within each serotype? Phylogenetic (Messer et al., 2003; Rico-Hesse et al., 1997) and epidemiological evidence (Messer et al., 2002; Uzcategui et al., 2003) suggests that specific genotypes are responsible for the onset of DEN epidemics characterized by severe pathogenicity. We now know that the introduction of Southeast Asian genotypes in the Americas coincided with the origin and spread in the New World of DHF (Rico-Hesse, 1990), whereas the American DENV genotypes are not associated with severe disease due to their low virulence potential (Leitmeyer et al., 1999). The selective forces that influence the occurrence of outbreaks and disease severity remain poorly understood. Several of detailed phylogenetic analyses based on diverse strains of endemic DENV genotypes have revealed evidence of limited, localized adaptive evolution. However, the selective targets appear vary among serotypes, genotypes and viral proteins, and very little is known about the evolutionary pressures that characterize sylvatic DENV evolution. Twiddy et al. utilized a ML method that measures rates of synonymous and nonsynonymous substitution codon-by-codon (Yang et al., 2000) on the E gene (Twiddy et al., 2002). A ratio of dN/dS < 1, indicates purifying (negative) selection, whereas a ratio of dN/dS > 1, indicates positive selection. A limitation of these studies that focus on codons effected convergently is that unique mutations, involving codons selected only once during the course of evolution, cannot be identified. The analysis revealed that the E gene of DENV-2 is
48
Nikos Vasilakis and Scott C. Weaver
subject to strong purifying selective constraints (expressed as the extremely low ratio of dN/dS), with evidence of weak positive selection in the Cosmopolitan (1 site) and lineage 2 (17 sites) of the Asian genotypes. The level of observed positive selection in the Cosmopolitan genotypes was much higher than that in the Asian genotypes (0.190 vs 0.056), which suggests that increased fitness may correlate with their dispersal potential. Of particular interest was the positively selected E390 site within the Cosmopolitan genotypes, which has been previously identified as a key virulence determinant (Sanchez and Ruiz, 1996) and maps within the distal face of domain III, a region associated with viral attachment to host cells. The same group also performed expanded analyses to measure the selection pressures within all DENV serotypes on larger datasets of E gene sequences (for DENV-2 only) (Twiddy et al., 2002). Due to the small sample size of DENV-1 sequences, no positively selected sites were detected; however, weak positive selection was detected in both DENV-3 (2 sites) and DENV-4 (5 sites), and in the Cosmopolitan (2 sites) and lineage 2 of the Asian (17 sites) DENV-2 genotypes. The majority of the selected sites (E-169 of DENV-3; E-357 and E-429 of DENV-4; and E-52, E85, E-90, E-122, E-131, E-144, E-170, E-330, E-334, E-342, E-378, and E-392 of DENV-2) were located within or near potential B- or T-cell epitopes (Aaskov et al., 1989; Innis et al., 1989; Kutubuddin et al., 1991; Leclerc et al., 1993; Megret et al., 1992; Roehrig et al., 1994), an association that suggests immune evasion as a selective factor (Twiddy et al., 2002). A number of selected sites were also located within functional domains involved in cell tropism (E-380 of DENV-3; E-342, E-378 and E-392 of DENV-2) or fusion (E108 and E-131 of DENV-4; E-52, E-98, E-100, E-105, E112 and E-113 of DENV2), suggesting that cell tropism and/or virus-mediated membrane fusion may also confer selective advantages by increasing fitness. A dataset of 36 DENV-2 complete genomes was also used to reveal the nature of selection pressures acting across the DENV genome. Interestingly, there was no evidence of positive selection in either PrM or capsid, and only localized positive selection was observed in NS2B (2 sites) and NS5 (2 sites). The selected sites (NS2B-57 and NS2B-63) are located within a 40 amino acid segment in NS2B that has been shown to be essential for NS2B/NS3 protease activity (Falgout et al., 1993), suggesting that these sites may play a role in the efficiency of the polyprotein processing. One of the selected sites in NS5 (NS5-135), is located within the conserved S-adenosylmethionine-utilizing methyltransferase (SAM) domain, suggesting a involvement in capping of virus genomes, whereas the other site (NS5-637) is located within the RNA-dependent RNA polymerase (RdRp) domain. Twiddy et al. performed additional analyses that included a limited number of sylvatic DENV-2 sequences (P8-1407, DakAr A578, PM339474 and DakArHD10674) (Twiddy et al., 2002). Surprisingly, there was no
History and Evolution of Human Dengue Emergence
49
detection of any positive selection (dN/dS ¼ 0.031) in the E gene. In a more extensive study that included 14 complete sylvatic DENV-2 sequences, gene-specific dN/dS ratios presented a similar picture of strong selective constraints (dN/dS << 1) (Vasilakis et al., 2007a). However, 16 amino acid sites showed evidence for putative weak positive selection and were unevenly distributed across the DENV genome. Of these, two were located in the E (177, 329), one in NS2A (195), and thirteen in NS4B (12, 19, 21, 24, 26, 49, 96, 113, 116, 123, 197, 242, 246) genes. The functional importance of the observed changes is not known and none has of these amino acid residues has previously been shown to mediate adaptive evolution in DENV (Twiddy et al., 2002). Interestingly, the analysis showed no evidence of positive selection acting upon NS4B in endemic DENV-2, and no positive selection was detected in the sylvatic lineage at amino acid E390, a key virulence determinant that has previously been reported to exhibit positive selection in endemic DENV-2 (Sanchez and Ruiz, 1996; Twiddy et al., 2002; Vasilakis et al., 2007a). Overall, these studies imply that DENV-2 have not undergone extensive adaptive evolution after emergence of the endemic, urban transmission cycles. At face value this suggests that emergence into the endemic transmission cycle was not facilitated by natural selection or host-specific (vertebrate and/or vector) adaptations, but rather by changing ecologic conditions. The major implication of the latter inference is that sylvatic DENV that are currently circulating in the forests of West Africa and Southeast Asia would have no trouble reinitiating an endemic transmission cycle should the proper conditions arise. In fact recent phylogenetic analyses based on complete DENV-2 genomes from human isolates associated with a 1966 outbreak of DF in Nigeria (Carey et al., 1971) suggest that limited spillover outbreaks of sylvatic DENV into urban settings can occur (Vasilakis et al., 2008b). These spillover outbreaks are defined as the enzootic transmission into a small, localized group of people often confined to a village, due to favorable ecological conditions such as increased vector densities. Enhanced passive or active surveillance of human populations in enzootic regions of West Africa and Asia are needed to evaluate the frequency of these spillover outbreaks and their public health impacts. Quantification of human viremias following infection with sylvatic DENV strains is also needed to further evaluate the facility with which these strains could initiate endemic transmission.
D. Adaptation for urban transmission As described in previous sections, four independent evolutionary events that occurred repeatedly in Asia probably resulted in the emergence of the four endemic DENV serotypes from sylvatic progenitors. Emergence was also facilitated by vector switching, from arboreal canopy-dwelling Aedes mosquito species to peridomestic Aedes mosquitoes, as well as
50
Nikos Vasilakis and Scott C. Weaver
vertebrate host changes, from non-human primates to humans (Wang et al., 2000). The gradual rise of urban populations large enough to sustain transmission, as well as the colonization of most tropical regions by the peridomestic mosquito vectors (Ae. aegypti and Ae. albopictus), provided a unique opportunity for adaptation and emergence into the major urban centers of the tropics worldwide. Today, urban DENV are both ecologically and evolutionarily independent from the ancestral sylvatic cycles. Since Ae. aegypti mosquitoes had not colonized the tropics until the establishment of the commercial trade routes in seventeenth Century, Ae. albopictus was most likely the original human mosquito vector in the establishment of sustained human transmission cycles. To test the hypothesis that sylvatic DENV strains required adaptation to these vectors to establish endemic cycles, Moncayo et al. compared the oral infectivity and dissemination profiles of endemic (Asian genotypes 1349 and NGC) and sylvatic DENV-2 strains from West Africa (PM33974 and DakAr 2022) and Asia (P8–1407) in Ae. aegypti and Ae. albopictus from diverse geographic locations in southeast Asia and the Americas (Moncayo et al., 2004). Ae. albopictus, regardless of its geographic origin, was more susceptible to endemic DENV-2 strains than Ae. aegypti (94–69% respectively), suggesting a higher degree of adaptation. This supports the idea of Ae. albopictus mosquitoes being the original peridomestic vector due to their longer historical contact with DENV in Asia. When the infection and dissemination rates from each group were pooled, endemic DENV-2 demonstrated higher rates of infection, but not dissemination from the midgut, than sylvatic DENV-2 strains in most populations of both mosquito species (Moncayo et al., 2004). However, American genotype DENV-2 strains, which are now known to exhibit reduced infectivity for Ae. aegypti compared to Asian genotypes (Armstrong and Rico-Hesse, 2001, 2003), were not included in this study. More recent studies suggest that the infection profiles the sylvatic and American DENV-2 genotypes for Ae. aegypti are similar, and that adaptation for enhanced transmission by this vector occurred after the emergence of the Asian genotype (K. Hanley, N. Vasilakis, unpublished). Diallo et al. addressed potential differences in DENV-2 genotypes with experimental infections of sylvatic and various peridomestic populations of Senegalese mosquitoes (Kedougou and Koung Koung), using both sylvatic (PM33974 and DakAr 2022) and endemic (1349 and NGC) DENV-2 strains (Diallo et al., 2005). Surprisingly, the sylvatic mosquito vectors Ae. furcifer and Ae. luteocephalus were highly susceptible to both sylvatic and endemic DENV-2 strains. In contrast, sylvatic Aedes (Stegomyia) vittatus and both sylvatic and peridomestic populations of Ae. aegypti were relatively refractory to all DENV-2 strains tested. This study involved Ae. aegypti populations collected from distinct ecological environments: mosquitoes from Kedougou are zoophilic and colonize
History and Evolution of Human Dengue Emergence
51
tree holes and other sylvatic breeding sites, whereas those from Koung Koung are anthropophilic, are usually to be found within human dwellings, and use artificial water containers as larval development sites. Surprisingly, these two Senegalese Ae. aegypti populations were not significantly different in their susceptibility to DENV-2 (Diallo et al., 2005), suggesting limitations on the emergence potential of enzootic strains. Likewise, a lack of correlation was observed between infection and dissemination rates; Ae. furcifer and Ae. luteocephalus exhibited high infection rates but low dissemination rates, whereas Ae. aegypti and Ae. vittatus exhibited low infection but high dissemination rates, which suggests a tradeoff between infection and dissemination rates that influences the transmission potential of these vectors. Collectively the data described above contrast with the results for the peridomestic DENV-2 vectors by Moncayo et al., where infection and dissemination rates were high for both sylvatic and epidemic strains (Moncayo et al., 2004). The data also indicate that adaptation of the Asian genotype DENV-2 to urban vectors did not result in a loss of infectivity for some African sylvatic vectors. Thus little or no change in vector infectivity accompanied the emergence of endemic DENV-2 from sylvatic progenitors, implying that peridomestic vector susceptibility, at least in some locations, is not an impediment to further sylvatic DENV-2 emergence. Whether emergence of endemic DENV was also facilitated by adaptation of sylvatic DENV to utilize humans as reservoir and amplification hosts for peridomestic transmission was only recently evaluated in two surrogate human model hosts, mice engrafted with human hepatoma (Huh-7) cells (Fig. 8A)(Table IV) and moDCs (Fig. 8B) (Table V) (Vasilakis et al., 2007b). In this study, six strains of low passage history that represented all major DENV-2 genotypes, including both African and Asian sylvatic strains, and Asian, African and American endemic strains, were included. Both Asian and American endemic strains were included for evaluation because they are reported to differ in their human virulence (Cologna et al., 2005; Rico-Hesse et al., 1997). The in vivo data demonstrated significant overall differences among mean replication titers of the DENV-2 strains examined, and statistical analyses revealed significant differences among individual pairs of strains. Surprisingly, this complex pattern of differences did not reveal a consistent or overall difference between sylvatic and endemic strains or between Asian endemic and American endemic strains. For example, the titer of Asian endemic strain 16681 was significantly higher than any of the other three endemic strains examined, but not significantly different from sylvatic strain P8–1407 (Table IV). Furthermore, the sylvatic strains replicated to lower titers than the endemic Asian strains, but did not differ consistently from the endemic American strains (Vasilakis et al., 2007b)(Table IV). Similarly, the
TABLE IV Replication profile of endemic and sylvatic DENV-2 in vivo
b c d e f
Statistical groupf
Virusa
Host
Location/yearb
Epidemiological typec
16681 1349 1328 IQT-1950 P8-1407
Human Human Human Human Sentinel Monkey Ae. africanus
Thailand/1964 Burkina Faso/1982 Puerto Rico/1977 Peru/1995 Malaysia/1970
Asian Endemic Asian Endemic American Endemic American Endemic Sylvatic
7 6 6 6 7
5.9 0.2 4.9 0.3 2.9 0.3 4.0 0.3 5.4 0.2
A B, C D C, E A, B
Republic of Guinea/1981
Sylvatic
6
3.6 0.2
D, E
PM33974 a
Mean peak virus titerd,e (log10 ffu/ml S.E.)
No. of mice
Groups of SCID-Huh-7 mice were inoculated into the tumor with 4.0–4.2 log10 ffu of the indicated virus. Serum was collected on day 7 and virus titer was determined on C6/36 cells. Indicates the country and year that virus strain was isolated. ‘‘Endemic’’ denotes human or Ae. aegypti isolates or strains that are associated with peridomestic transmission. Virus titer in serum was determined by focus forming immunoassay (FFA) on C6/36 cells. The limit of detection of the assay is 0.5 ffu/ml. Mean peak titers were assigned to statistical groups using one-way ANOVA, df ¼ 5; F ¼ 25.6; P < 0.0001 and the Tukey–Kramer post hoc test (P < 0.05). Groups with the same letter designation are not significantly different; all means not sharing a letter are significantly different.
TABLE V
a b c d e f
Replication profile of endemic and sylvatic DENV-2 ex vivo
Virusa
Host
Location/yearb
Epidemiological typec
Mean peak virus titerd,e (log10 ffu/ ml S.E.)
16681 1349 1328 IQT-1950 P8-1407 PM33974 DakAr A510 DakAr A1247 DakAr 2022 DakAr 2039
Human Human Human Human Sentinel Monkey Ae. africanus Ae. Taylori Ae. Taylori Ae. africanus Ae. luteocephalus
Thailand/1964 Burkina Faso/1982 Puerto Rico/1977 Peru/1995 Malaysia/1970 Republic of Guinea/1981 Coˆte d’Ivoire/1980 Coˆte d’Ivoire/1980 Burkina Faso/1980 Burkina Faso/1980
Asian Endemic Asian Endemic American Endemic American Endemic Sylvatic Sylvatic Sylvatic Sylvatic Sylvatic Sylvatic
5.4 0.1 5.0 0.3 3.1 0.6 4.3 0.2 3.2 0.0 0.5 0.0 4.4 0.2 0.5 0.0 2.6 0.1 4.6 0.2
Statistical groupf
A A B B B C B C B B
moDCs (2.5 105 per sample) from two healthy human volunteers were infected with an MOI ¼ 2 of the indicated virus. Supernatants were collected 48 h p.i. and viral output was determined on C6/36 cells. Indicates the country and year that virus strain was isolated. ‘‘Endemic’’ denotes human or Ae. aegypti isolates or strains that are associated with peridomestic transmission. Virus titer in serum was determined by focus forming immunoassay (FFA) on C6/36 cells. The limit of detection of the assay is 0.5 log10 ffu/ml. Mean peak titers were assigned to statistical groups using the Tukey–Kramer post hoc test (P < 0.05). Groups with the same letter designation are not significantly different; all means not sharing a letter are significantly different. Unpaired t-test endemic versus sylvatic using mean values from four endemic and six sylvatic: df ¼ 8; t ¼ 1.35; P ¼ 0.21.
54
Nikos Vasilakis and Scott C. Weaver
ex vivo data detected significant inter-strain variation in mean replication titers among the various DENV-2 strains, but no overall difference between sylvatic and endemic strains. The ex vivo moDCs model (Fig. 8B) also provided an opportunity to determine whether host genetics (i.e., different racial backgrounds) may influence the outcome of DENV infection. Although moDCs were obtained from a small number of human donors representing different ethnicities (Caucasian, West African, and Southeast Asian), no differences were observed in their ability to support DENV-2 replication (Table V) (Vasilakis et al., 2007b). A potential limitation of this study was the small sample size (n ¼ 2) of strains representing the major DENV-2 genotypes. It is possible that there is a detectable difference between sylvatic and endemic strains, but that overall differences were masked by the limited sample size. Future studies with extended sample size are needed. These studies should also extend to the other DENV serotypes, although very few sylvatic DENV1 and -4 strains are available and sylvatic DENV-3 has not been isolated. Overall, these findings do not support the hypothesis that emergence of endemic DENV was also facilitated by adaptation of sylvatic DENV to utilize humans as reservoir and amplification hosts for peridomestic transmission. This implies that the potential for re-emergence of sylvatic DENV strains from Africa or Asia into the endemic cycle is high. Emergence and adaptation are significant issues for arbovirology and pose enormous public health implications, especially considering the potential for eradicating the human transmission cycle with effective vaccines now under development.
E. Conclusions and future work Although details remain unclear, especially for DENV-3 and DENV-4, most evidence supports the hypothesis that all four DENV serotypes emerged into the human population independently from sylvatic progenitors that circulate in forests of Asia among nonhuman primates via arboreal mosquito vectors. The emergence of endemic DENV is a relatively recent event that probably occurred within the past 2 thousand years after human populations reached densities and sizes amenable to sustained transmission. The dramatic increase in DENV incidence and its geographic spread probably depended primarily on the colonization of most tropical locations by the highly anthropophilic and efficient vector, Ae. aegypti, as well as on the acceleration of urbanization following World War II. The evolution of enhanced pathogenicity and infectivity for Ae. aegypti by the Asian genotype of DENV-2 probably played an important role in the recent increase in severe DENV in the Americas. Studies examining the fitness of the ancestral, sylvatic DENV-2 compared to currently circulating, endemic/epidemic strains, using
History and Evolution of Human Dengue Emergence
55
surrogates for human infection and experimental infections of vectors, suggest that little or no adaptation was required for the initial emergence of endemic DENV-2 strains. This implies that the sylvatic cycles in Asia and West Africa will remain a source of re-emergence. Although several promising DENV vaccines under development hold the hope for the eradication of endemic DENV strains (because humans are the only reservoir hosts), the sylvatic strains are not amenable to control and will probably remain a source of re-emergence. However, human herd immunity from exposure to endemic DENV strains and immunity generated by some vaccines is capable of sylvatic strain neutralization, indicating that sustained vaccination could prevent re-emergence. The predictions described above are based on very limited studies with imperfect models of human infection, and the testing of limited DENV strains despite extensive evidence that these viruses exhibit a high degree of strain variation in phenotypes. Therefore, additional studies of the fitness of sylvatic DENV, especially serotypes other than DENV-2, are needed. Ultimately, the data needed to evaluate the human infection potential and disease burden will come from epidemiological studies in West Africa and Asia where the sylvatic strains circulate near human populations. Of particular importance will be characterization of the disease caused by the sylvatic strains, quantification of viremia titers to determine the potential for infected people to initiate interhuman transmission if suitable vectors are present, and ecological contacts between the forest cycles and people via studies of human and sylvatic vector movement and contact.
ACKNOWLEDGMENTS We thank Edward C. Holmes for helpful advice with the phylogenetic methods and divergence time estimates. NV was supported by the Centers for Disease Control and Prevention Fellowship Training Program in Vector-Borne Infectious Diseases, T01/CCT622892.
REFERENCES Aaskov, J. G., Geysen, H. M., and Mason, T. J. (1989). Serologically defined linear epitopes in the envelope protein of dengue 2 ( Jamaica strain 1409). Arch. Virol. 105(3–4):209–221. Aaskov, J., Buzacott, K., Field, E., Lowry, K., Berlioz-Arthaud, A., and Holmes, E. C. (2007). Multiple recombinant dengue type 1 viruses in an isolate from a dengue patient. J. Gen. Virol. 88:3334–3340. AbuBakar, S., Wong, P. F., and Chan, Y. F. (2002). Emergence of dengue virus type 4 genotype IIA in Malaysia. J. Gen. Virol. 83(Pt 10):2437–2442. An, J., Kimura-Kuroda, J., Hirabayashi, Y., and Yasui, K. (1999). Development of a novel mouse model for dengue virus infection. Virology 263(1):70–77. Anderson, R., King, A. D., and Innis, B. L. (1992). Correlation of E protein binding with cell susceptibility to dengue 4 virus infection. J. Gen. Virol. 73(Pt 8):2155–2159.
56
Nikos Vasilakis and Scott C. Weaver
Angsubhakorn, S., Yoksan, S., Bhamarapravati, N., Moe, J. B., Marchette, N. J., Pradermwong, A., and Sahaphong, S. (1988). Dengue-4 vaccine: Neurovirulence, viraemia and immune responses in rhesus and cynomolgus monkeys. Trans. R. Soc. Trop. Med. Hyg. 82(5):746–749. Appanna, R., Huat, T. L., See, L. L., Tan, P. L., Vadivelu, J., and Devi, S. (2007). Cross-reactive T-cell responses to the nonstructural regions of dengue viruses among dengue fever and dengue hemorrhagic fever patients in Malaysia. Clin. Vaccine Immunol. 14(8):969–977. Araujo, F. M., Nogueira, R. M., de Araujo, J. M., Ramalho, I. L., Roriz, M. L., de Melo, M. E., and Coelho, I. C. (2006). Concurrent infection with dengue virus type-2 and DENV-3 in a patient from Ceara, Brazil. Mem. Inst. Oswaldo Cruz 101(8):925–928. Armstrong, P. M., and Rico-Hesse, R. (2001). Differential susceptibility of Aedes aegypti to infection by the American and Southeast Asian genotypes of dengue type 2 virus. Vector Borne Zoonotic Dis. 1(2):159–168. Armstrong, P. M., and Rico-Hesse, R. (2003). Efficiency of dengue serotype 2 virus strains to infect and disseminate in Aedes aegypti. Am. J. Trop. Med. Hyg. 68(5):539–544. Ashburn, P. M., and Craig, C. F. (1907). Experimental investigations regarding the etiology of dengue fever. J. Infect. Dis. 4:440–475. Autorino, G. L., Battisti, A., Deubel, V., Ferrari, G., Forletta, R., Giovannini, A., Lelli, R., Murri, S., and Scicluna, M. T. (2002). West Nile virus epidemic in horses, Tuscany region, Italy. Emerg. Infect. Dis. 8(12):1372–1378. Avirutnan, P., Malasit, P., Seliger, B., Bhakdi, S., and Husmann, M. (1998). Dengue virus infection of human endothelial cells leads to chemokine production, complement activation, and apoptosis. J. Immunol. 161(11):6338–6346. Avirutnan, P., Punyadee, N., Noisakran, S., Komoltri, C., Thiemmeca, S., Auethavornanan, K., Jairunsgsri, A., Kanlaya, R., Tangthawornchaikul, N., Puttikhunt, C., Pattanakisakul, S. N., Yenchitsomanus, P. T., et al. (2006). Vascular leakage in severe dengue virus infections: A potential role for the nonstructural viral protein NS1 and complement. J. Infect. Dis. 193:1078–1088. Aziz, M. M., Hasan, K. N., Hasanat, M. A., Siddiqui, M. A., Salimullah, M., Chowdhury, A. K., Ahmed, M., Alam, M. N., and Hassan, M. S. (2002). Predominance of the DEN-3 genotype during the recent dengue outbreak in Bangladesh. Southeast Asian J. Trop. Med. Public Health 33(1):42–48. Baer, G. M., and Woodall, D. F. (1966). Bat salivary gland virus carrier state in a naturally infected Mexican freetail bat. Am. J. Trop. Med. Hyg. 15(5):769–771. Balaya, S., Paul, S. D., D’Lima, L. V., and Pavri, K. M. (1969). Investigations on an outbreak of dengue in Delhi in 1967. Indian J. Med. Res. 57(4):767–774. Barnes, W. J., and Rosen, L. (1974). Fatal hemorrhagic disease and shock associated with primary dengue infection on a Pacific island. Am. J. Trop. Med. Hyg. 23(3):495–506. Barraud, P. J. (1928). The distribution of ‘Stegomyia fasciata’ in India, with remarks on dengue and yellow fever. Indian J. Med. Res. 16:377–398. Basaca-Sevilla, V., and Halstead, S. B. (1966). Recent virological studies on haemorrhagic fever and other arthropod-borne virus infections in the Philippines. J. Trop. Med. Hyg. 69 (9):203–208. Becher, P., Orlich, M., and Thiel, H. J. (2001). RNA recombination between persisting pestivirus and a vaccine strain: Generation of cytopathogenic virus and induction of lethal disease. J. Virol. 75(14):6256–6264. Belov, G. F., Tofaniuk, E. V., Kurzhukov, G. P., and Kuznetsova, V. G. (1995). [The clinicoepidemiological characteristics of Omsk hemorrhagic fever in 1988–1992]. Zh. Mikrobiol. Epidemiol. Immunobiol. 4:88–91. Bennett, S. N., Holmes, E. C., Chirivella, M., Rodriguez, D. M., Beltran, M., Vorndam, V., Gubler, D. J., and McMillan, W. O. (2003). Selection-driven evolution of emergent dengue virus. Mol. Biol. Evol. 20(10):1650–1658.
History and Evolution of Human Dengue Emergence
57
Bente, D. A., Melkus, M. W., Garcia, J. V., and Rico-Hesse, R. (2005). Dengue fever in humanized NOD/SCID mice. J. Virol. 79(21):13797–13799. Bernard, K. A., Klimstra, W. B., and Johnston, R. E. (2000). Mutations in the E2 glycoprotein of Venezuelan equine encephalitis virus confer heparan sulfate interaction, low morbidity, and rapid clearance from blood of mice. Virology 276(1):93–103. Blackley, S., Kou, Z., Chen, H., Quinn, M., Rose, R. C., Schlesinger, J. J., Coppage, M., and Jin, X. (2007). Primary human splenic macrophages, but not T or B cells, are the principal target cells for dengue virus infection in vitro. J. Virol. 81(24):13325–13334. Blaney, J. E., Jr., Johnson, D. H., Manipon, G. G., Firestone, C. Y., Hanson, C. T., Murphy, B. R., and Whitehead, S. S. (2002). Genetic basis of attenuation of dengue virus type 4 small plaque mutants with restricted replication in suckling mice and in SCID mice transplanted with human liver cells. Virology 300(1):125–139. Blaney, J. E., Jr., Matro, J. M., Murphy, B. R., and Whitehead, S. S. (2005). Recombinant, liveattenuated tetravalent dengue virus vaccine formulations induce a balanced, broad, and protective neutralizing antibody response against each of the four serotypes in rhesus monkeys. J. Virol. 79(9):5516–5528. Bonner, S. M., and O’Sullivan, M. A. (1998). Endothelial cell monolayers as a model system to investigate dengue shock syndrome. J. Virol. Methods 71(2):159–167. Boonpucknavig, S., Vuttiviroj, O., and Boonpucknavig, V. (1981). Infection of young adult mice with dengue virus type 2. Trans. R. Soc. Trop. Med. Hyg. 75(5):647–653. Bosch, I., Xhaja, K., Estevez, L., Raines, G., Melichar, H., Warke, R. V., Fournier, M. V., Ennis, F. A., and Rothman, A. L. (2002). Increased production of interleukin-8 in primary human monocytes and in human epithelial and endothelial cell lines after dengue virus challenge. J. Virol. 76(11):5588–5597. Botros, B. A., Watts, D. M., Soliman, A. K., Salib, A. W., Moussa, M. I., Mursal, H., Douglas, C., and Farah, M. (1989). Serological evidence of dengue fever among refugees, Hargeysa, Somalia. J. Med. Virol. 29(2):79–81. Bouree, P., Lancon, A., Anquetil, R., and Menager, C. (2001). [Dengue in New Caledonia. Study of 68 pediatric cases]. Arch. Pediatr. 8(12):1311–1317. Brandt, W. E., McCown, J. M., Gentry, M. K., and Russell, P. K. (1982). Infection enhancement of dengue type 2 virus in the U-937 human monocyte cell line by antibodies to flavivirus cross-reactive determinants. Infect. Immun. 36(3):1036–1041. Bravo, J. R., Guzman, M. G., and Kouri, G. P. (1987). Why dengue haemorrhagic fever in Cuba? Part 1. Individual risk factors for dengue haemorrhagic fever/dengue shock syndrome (DHF/DSS) Trans. R. Soc. Trop. Med. Hyg. 81(5):816–820. Brinker, K. R., and Monath, T. P. (1980). The acute disease. In ‘‘St. Louis Encephalitis’’ (T.P Monath, Ed.), pp. 503–534. American Public Health Association, Washington, DC. Brown, A. W. A. (1977). Yellow fever, dengue and dengue haemorrhagic fever. In ‘‘A World Geography of Human Diseases’’ (G. M Howe, Ed.), pp. 271–317. Academic Press, New York. Bush, R. M., Fitch, W. M., Bender, C. A., Cox, N. J. (1999) Positive selection on the H3 hemagglutinin gene of human influenza virus A. Mol. Biol. Evol. 16(11):1457–1465. Burke, D. S., Nisalak, A., Johnson, D. E., and Scott, R. M. (1988). A prospective study of dengue infections in Bangkok. Am. J. Trop. Med. Hyg. 38(1):172–180. Bylon, D. (1780). Korte Aantekening, wegens eene Algemeene Ziekte, Doorgaans Genaamd de Knokkel-Koorts. In ‘‘Verhandelungen van het bataviaasch Genootschop der Konsten in Wetenschappen’’,Vol. 2, pp. 17–30. Batavia. Johannes Allart (Amsterdam). Byrnes, A. P., and Griffin, D. E. (2000). Large-plaque mutants of Sindbis virus show reduced binding to heparan sulfate, heightened viremia, and slower clearance from the circulation. J. Virol. 74(2):644–651. Calisher, C. H., and Gould, E. A. (2003). Taxonomy of the virus family Flaviviridae. Adv. Virus Res. 59:1–19.
58
Nikos Vasilakis and Scott C. Weaver
Calisher, C. H., Karabatsos, N., Dalrymple, J. M., Shope, R. E., Porterfield, J. S., Westaway, E. G., and Brandt, W. E. (1989). Antigenic relationships between flaviviruses as determined by cross-neutralization tests with polyclonal antisera. J. Gen. Virol. 70(Pt. 1):37–43. Cardamatis, J. P. (1929). La dengue en Grece. Bull. Soc. Path. Exot. Filiales 22:272–292. Cardier, J. E., Balogh, V., Perez-Silva, C., Romano, E., Rivas, B., Bosch, N., and Rothman, A. L. (2006). Relationship of thrombopoietin and interleukin-11 levels to thrombocytopenia associated with dengue disease. Cytokine 34:155–160. Carey, D. E. (1971). Chikungunya and dengue: A case of mistaken identity? J. Hist. Med. Allied Sci. 26(3):243–262. Carey, D. E., Causey, O. R., Reddy, S., and Cooke, A. R. (1971). Dengue viruses from febrile patients in Nigeria, 1964–1968. Lancet 1(7690):105–106. Carrington, C. V., Foster, J. E., Pybus, O. G., Bennett, S. N., and Holmes, E. C. (2005). Invasion and maintenance of dengue virus type 2 and type 4 in the Americas. J. Virol. 79 (23):14680–14687. Chakravarti, A., and Kumaria, R. (2006). Circulating levels of tumor necrosis factor-alpha & interferon-gamma in patients with dengue & dengue hemorrhagic fever during an outbreak. Indian J. Med. Res. 123:25–30. Chan, Y. C., Kanapathipillai, K., and Chew, K. S. (1965). Isolation of two strains of dengue virus type 3 in Singapore. Singapore Med. J. 6(3):127–132. Chao, D. Y., King, C. C., Wang, W. K., Chen, W. J., Wu, H. L., and Chang, G. J. (2005). Strategically examining the full-genome of dengue virus type 3 in clinical isolates reveals its mutation spectra. Virol. J. 2:72. Charrel, R. N., Attoui, H., Butenko, A. M., Clegg, J. C., Deubel, V., Frolova, T. V., Gould, E. A., Gritsun, T. S., Heinz, F. X., Labuda, M., Lashkevich, V. A., Loktev, V., et al. (2004). Tickborne virus diseases of human interest in Europe. Clin. Microbiol. Infect. 10(12):1040–1055. Chaturvedi, U. C., Dhawan, R., Khanna, M., and Mathur, A. (1991). Breakdown of the blood– brain barrier during dengue virus infection of mice. J. Gen. Virol. 72(Pt. 4):859–866. Chen, Y., Maguire, T., and Marks, R. M. (1996). Demonstration of binding of dengue virus envelope protein to target cells. J. Virol.. 70(12):8765–8772. Chen, Y., Maguire, T., Hileman, R. E., Fromm, J. R., Esko, J. D., Linhardt, R. J., and Marks, R. M. (1997). Dengue virus infectivity depends on envelope protein binding to target cell heparan sulfate. Nat. Med. 3(8):866–871. Chen, Y. C., Wang, S. Y., and King, C. C. (1999). Bacterial lipopolysaccharide inhibits dengue virus infection of primary human monocytes/macrophages by blockade of virus entry via a CD14-dependent mechanism. J. Virol. 73(4):2650–2657. Chen, R. F., Liu, J. W., Yeh, W. T., Wang, L., Chang, J. C., Yu, H. R., Cheng, J. T., and Yang, K. D. (2005). Altered T helper 1 reaction but not increase of virus load in patients with dengue hemorrhagic fever. FEMS Immunol. Med. Microbiol. 44(1):43–50. Chen, L. C., Lei, H. Y., Liu, C. C., Shiesh, S. C., Chen, S. H., Liu, H. S., Lin, Y. S., Wang, S. T., and Shyu, H. W. (2006). Correlation of serum levels of macrophage migration inhibitory factor with disease severity and clinical outcome in dengue patients. Am. J. Trop. Med. Hyg. 74:142–147. Christie, J. (1881). On epidemics of dengue fever: Their diffusion and etiology. Glasgow Med. J. 16:161–176. Christophers, S. R. (1960). Systematic. In ‘‘Aedes aegypti, The Yellow Fever Mosquito: Its life History, Bionomics and Structure’’, pp. 21–53. Cambridge University Press, London. Cleland, J. E., Bradley, B., and McDonald, W. (1916). On the transmission of Australian dengue by the mosquito Stegomyia fasciata. Med. J. Aust. 2:179–200. Clifford, C. M., Yunker, C. E., Thomas, L. A., Easton, E. R., and Corwin, D. (1971). Isolation of a Group B arbovirus from Ixodes uriae collected on Three Arch Rocks National Wildlife Refuge, Oregon. Am. J. Trop. Med. Hyg. 20(3):461–468.
History and Evolution of Human Dengue Emergence
59
Cobra, C., Rigau-Perez, J. G., Kuno, G., and Vorndam, V. (1995). Symptoms of dengue fever in relation to host immunologic response and virus serotype, Puerto Rico, 1990–1991. Am. J. Epidemiol. 142(11):1204–1211. Cole, G. A., and Wisseman, C. L., Jr. (1969). Pathogenesis of type 1 dengue virus infection in suckling, weanling and adult mice. Part 1. The relation of virus replication to interferon and antibody formation. Am. J. Epidemiol. 89(6):669–680. Cologna, R., and Rico-Hesse, R. (2003). American genotype structures decrease dengue virus output from human monocytes and dendritic cells. J. Virol. 77(7):3929–3938. Cologna, R., Armstrong, P. M., and Rico-Hesse, R. (2005). Selection for virulent dengue viruses occurs in humans and mosquitoes. J. Virol. 79(2):853–859. Constantine, D. G., and Woodall, D. F. (1964). Latent infection of Rio Bravo virus in salivary glands of bats. Public Health Rep. 79:1033–1039. Cooper, L. A., and Scott, T. W. (2001). Differential evolution of eastern equine encephalitis virus populations in response to host cell type. Genetics/Society 157(4):1403–1412. Cooper, 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. Copanaris, P. (1928). L’epidemie de dengue en Grece au cours de l’ete 1928. Office International d’Hygiene Publique Bulletin. 20:1590–1601. Cordeiro, M. T., Schatzmayr, H. G., Nogueira, R. M., Oliveira, V. F., Melo, W. T., and Carvalho, E. F. (2007). Dengue and dengue hemorrhagic fever in the State of Pernambuco, 1995–2006. Rev. Soc. Bras. Med. Trop. 40(6):605–611. Cordellier, R., Bouchite, B., Roche, J. C., Monteny, N., Diaco, B., and Akoliba, P. (1983). Circulation selvatique du virus Dengue 2, en 1980, dans les savanes sub-soudaniennes de Cote d’Ivoire. Cah. ORSTOM. ser Ent. Med. et Parasitol. 21:165–179. Cornet, M. (1993). Dengue in Africa. In ‘‘Monograph on Dengue/Dengue Haemorrhagic Fever’’, (P. Thongcharoen, Ed.), pp. 39–47. WHO Regional Publication, South-East Asia. World Health Organization, Regional Office for South-East Asia, New Delhi, India. Cornet, M., Saluzzo, J. F., Hervy, J. P., Digoutte, J. P., Germain, M., Chauvancy, M. F., Eyraud, M., Ferrara, L., Heme, G., and Legros, F. (1984). Dengue 2 au Senegal oriental: Une poussee epizootique en milieu selvatique; isolements du virus a partir des moustiques et d’un singe et considerations epidemiologiques. Cah. ORSTOM. ser Ent. Med. et Parasitol. 22:313–323. Corwin, A. L., Larasati, R. P., Bangs, M. J., Wuryadi, S., Arjoso, S., Sukri, N., Listyaningsih, E., Hartati, S., Namursa, R., Anwar, Z., Chandra, S., Loho, B., et al. (2001). Epidemic dengue transmission in southern Sumatra, Indonesia. Trans. R. Soc. Trop. Med. Hyg. 95 (3):257–265. Craig, S., Thu, H. M., Lowry, K., Wang, X. F., Holmes, E. C., and Aaskov, J. (2003). Diverse dengue type 2 virus populations contain recombinant and both parental viruses in a single mosquito host. J. Virol. 77(7):4463–4467. Cummings, D. A., Irizarry, R. A., Huang, N. E., Endy, T. P., Nisalak, A., Ungchusak, K., and Burke, D. S. (2004). Travelling waves in the occurrence of dengue haemorrhagic fever in Thailand. Nature 427(6972):344–347. Cunha, R. V., Schatzmayr, H. G., Miagostovich, M. P., Barbosa, A. M., Paiva, F. G., Miranda, R. M., Ramos, C. C., Coelho, J. C., dos Santos, F. B., and Nogueira, R. M. (1999). Dengue epidemic in the State of Rio Grande do Norte, Brazil, in 1997. Trans. R. Soc. Trop. Med. Hyg. 93(3):247–249. da Cunha, R. V., Maspero, R. C., Miagostovich, M. P., de Araujo, E. S., Luz Dda, C., Nogueira, R. M., and Schatzmayr, H. G. (1997). Dengue infection in Paracambi, State of Rio de Janeiro, 1990–1995. Rev. Soc. Bras. Med. Trop. 30(5):379–383. Daniels, C. W. (1908). Notes on the mosquitoes on the river and coast districts on the eastern side of the Peninsula. Stud. Inst. Med. Res. Malaya 3:262–266.
60
Nikos Vasilakis and Scott C. Weaver
Diallo, M., Ba, Y., Sall, A. A., Diop, O. M., Ndione, J. A., Mondo, M., Girault, L., and Mathiot, C. (2003). Amplification of the sylvatic cycle of dengue virus type 2, Senegal, 1999–2000: Entomologic findings and epidemiologic considerations. Emerg. Infect. Dis. 9(3):362–367. Diallo, M., Sall, A. A., Moncayo, A. C., Ba, Y., Fernandez, Z., Ortiz, D., Coffey, L. L., Mathiot, C., Tesh, R. B., and Weaver, S. C. (2005). Potential role of sylvatic and domestic african mosquito species in dengue emergence. Am. J. Trop. Med. Hyg. 73(2):445–449. Diamond, M. S., Edgil, D., Roberts, T. G., Lu, B., and Harris, E. (2000). Infection of human cells by dengue virus is modulated by different cell types and viral strains. J. Virol. 74 (17):7814–7823. Diaz, F. J., Black, W. C. T., Farfan-Ale, J. A., Lorono-Pino, M. A., Olson, K. E., and Beaty, B. J. (2006). Dengue virus circulation and evolution in Mexico: A phylogenetic perspective. Arch. Med. Res. 37(6):760–773. Domingo, E., and Holland, J. J. (1997). RNA virus mutations and fitness for survival. Annu. Rev. Microbiol. 51:151–178. Domingo, C., Palacios, G., Jabado, O., Reyes, N., Niedrig, M., Gascon, J., Cabrerizo, M., Lipkin, W. I., and Tenorio, A. (2006). Use of a short fragment of the C-terminal E gene for detection and characterization of two new lineages of dengue virus 1 in India. J. Clin. Microbiol. 44(4):1519–1529. Dove, A. (1998). Dengue fever on the increase. Nat. Med. 4(5):543. Durban (1927). ‘‘Dengue", as seen in the recent epidemic in Durban. J. Med. Assoc. S. Afr. 1:446–448. Edelman, R., Tacket, C. O., Wasserman, S. S., Vaughn, D. W., Eckels, K. H., Dubois, D. R., Summers, P. L., and Hoke, C. H. (1994). A live attenuated dengue-1 vaccine candidate (45AZ5) passaged in primary dog kidney cell culture is attenuated and immunogenic for humans. J. Infect. Dis. 170(6):1448–1455. Edington, A. D. (1927). ‘‘Dengue", as seen in the recent epidemic in Durban. J. Med. Assoc. S. Afr. 1:446–448. Edwards, F. W. (1932). Diptera.family culicidae. ‘‘Genera Insectorum’’, P. Wystman, ). Vol. 194: Desmet-Verteneuil, Brussels, Belgium. Ehrenkranz, N. J., Ventura, A. K., Cuadrado, R. R., Pond, W. L., and Porter, J. E. (1971). Pandemic dengue in Caribbean countries and the southern United States—past, present and potential problems. N. Engl. J. Med. 285(26):1460–1469. Endy, T. P., Nisalak, A., Chunsuttitwat, S., Vaughn, D. W., Green, S., Ennis, F. A., Rothman, A. L., and Libraty, D. H. (2004). Relationship of preexisting dengue virus (DV) neutralizing antibody levels to viremia and severity of disease in a prospective cohort study of DV infection in Thailand. J. Infect. Dis. 189(6):990–1000. Fagbami, A. (1977). Epidemiological investigations on arbovirus infections at Igbo-Ora, Nigeria. Trop. Geogr. Med. 29(2):187–191. Fagbami, A. (1978). Human arthropod-borne virus infections in Nigeria. Serological and virological investigations and Shaki, Oyo State. J. Hyg. Epidemiol. Microbiol. Immunol. 22 (2):184–189. Fagbami, A. H., and Fabiyi, A. (1976). Epidemiology of dengue infections in Nigeria: Virus isolations and clinical observations, 1972–1975. J. Trop. Med. Hyg. 79(10):226–228. Fagbami, A. H., Monath, T. P., and Fabiyi, A. (1977). Dengue virus infections in Nigeria: A survey for antibodies in monkeys and humans. Trans. R. Soc. Trop. Med. Hyg. 71(1):60–65. Failloux, A. B., Vazeille, M., and Rodhain, F. (2002). Geographic genetic variation in populations of the dengue virus vector Aedes aegypti. J. Mol. Evol. 55(6):653–663. Falgout, B., Miller, R. H., and Lai, C. J. (1993). Deletion analysis of dengue virus type 4 nonstructural protein NS2B: Identification of a domain required for NS2B-NS3 protease activity. J. Virol. 67(4):2034–2042. Fan, W. F., Yu, S. R., and Cosgriff, T. M. (1989). The reemergence of dengue in China. Rev. Infect. Dis. 11(Suppl. 4):S847–S853.
History and Evolution of Human Dengue Emergence
61
Fang, R., Lo, E., and Lim, T. W. (1984). The 1982 dengue epidemic in Malaysia: Epidemiological, serological and virological aspects. Southeast Asian J. Trop. Med. Public Health 15 (1):51–58. Ferguson, N., Anderson, R., and Gupta, S. (1999). The effect of antibody-dependent enhancement on the transmission dynamics and persistence of multiple-strain pathogens. Proc. Natl. Acad. Sci. USA 96(2):790–794. Foster, J. E., Bennett, S. N., Vaughan, H., Vorndam, V., McMillan, W. O., and Carrington, C. V. (2003). Molecular evolution and phylogeny of dengue type 4 virus in the Caribbean. Virology 306(1):126–134. Freier, J. E., and Grimstad, P. R. (1983). Transmission of dengue virus by orally infected Aedes triseriatus. Am. J. Trop. Med. Hyg. 32(6):1429–1434. Freier, J. E., and Rosen, L. (1987). Vertical transmission of dengue viruses by mosquitoes of the Aedes scutellaris group. Am. J. Trop. Med. Hyg. 37(3):640–647. Freier, J. E., and Rosen, L. (1988). Vertical transmission of dengue viruses by Aedes mediovittatus. Am. J. Trop. Med. Hyg. 39(2):218–222. George, R., Kassim, M. S., and Wah, L. T. (1974). Mosquito-borne haemorrhagic fever. Med. J. Malaysia 29(1):11–16. Germi, R., Crance, J. M., Garin, D., Guimet, J., Lortat-Jacob, H., Ruigrok, R. W., Zarski, J. P., and Drouet, E. (2002). Heparan sulfate-mediated binding of infectious dengue virus type 2 and yellow fever virus. Virology 292(1):162–168. Gonzalez, J. P., Du Saussay, C., Gautun, J. C., McCormick, J. B., and Mouchet, J. (1985). [Dengue in Burkina Faso (ex-Upper Volta): Seasonal epidemics in the urban area of Ouagadougou]. Bull. Soc. Pathol. Exot. Filiales 78(1):7–14. Goncalvez, A. P., Escalante, A. A., Pujol, F. H., Ludert, J. E., Tovar, D., Salas, R. A., and Liprandi, F. (2002). Diversity and evolution of the envelope gene of dengue virus type 1. Virology 303(1):110–119. Goncalvez, A. P., Engle, R. E., St Claire, M., Purcell, R. H., and Lai, C. J. (2007). Monoclonal antibody-mediated enhancement of dengue virus infection in vitro and in vivo and strategies for prevention. Proc. Natl. Acad. Sci. USA 104(22):9422–9427. Graham, H. (1903). The dengue: A study of its pathology and mode of propagation. J. Trop. Med. Lond. 6:209–214. Greene, I. P., Wang, E., Deardorff, E. R., Milleron, R., Domingo, E., and Weaver, S. C. (2005). Effect of alternating passage on adaptation of sindbis virus to vertebrate and invertebrate cells. J. Virol. 79(22):14253–14260. Gritsun, T. S., Nuttall, P. A., and Gould, E. A. (2003). Tick-borne flaviviruses. In ‘‘The Flaviviruses: Detection, diagnosis and vaccine development’’, (T. J. Chambers and T. P. Monath, Eds.), pp. 318–372. Elsevier, San Diego. Gubler, D. J. (1997). Dengue and dengue hemorrhagic fever: Its history and resurgence as a global public health problem. In ‘‘Dengue and Dengue Hemorrhagic Fever’’, (D. J. Gubler and G. Kuno, Eds.), pp. 1–22. CABI, New York. Gubler, D. J., and Rosen, L. (1976). Variation among geographic strains of Aedes albopictus in susceptibility to infection with dengue viruses. Am. J. Trop. Med. Hyg. 25(2):318–325. Gubler, D. J., Reed, D., Rosen, L., and Hitchcock, J. R., Jr. (1978). Epidemiologic, clinical, and virologic observations on dengue in the Kingdom of Tonga. Am. J. Trop. Med. Hyg. 27 (3):581–589. Gubler, D. J., Nalim, S., Tan, R., Saipan, H., and Sulianti Saroso, J. (1979). Variation in susceptibility to oral infection with dengue viruses among geographic strains of Aedes aegypti. Am. J. Trop. Med. Hyg. 28(6):1045–1052. Gubler, D. J., Suharyono, W., Lubis, I., Eram, S., and Gunarso, S. (1981). Epidemic dengue 3 in central Java, associated with low viremia in man. Am. J. Trop. Med. Hyg. 30(5):1094–1099. Gubler, D. J., Kuno, G., Sather, G. E., and Waterman, S. H. (1985a). A case of natural concurrent human infection with two dengue viruses. Am. J. Trop. Med. Hyg. 34(1):170–173.
62
Nikos Vasilakis and Scott C. Weaver
Gubler, D. J., Novak, R. J., Vergne, E., Colon, N. A., Velez, M., and Fowler, J. (1985b). Aedes (Gymnometopa) mediovittatus (Diptera: Culicidae), a potential maintenance vector of dengue viruses in Puerto Rico. J. Med. Entomol. 22(5):469–475. Gubler, D. J., Sather, G. E., Kuno, G., and Cabral, J. R. (1986). Dengue 3 virus transmission in Africa. Am. J. Trop. Med. Hyg. 35(6):1280–1284. Guilarde, A. O., Turchi, M. D., Jr., Siqueira, J. B., Feres, V. C., Rocha, B., Levi, J. E., Souza, V. A., Boas, L. S., Pannuti, C. S., and Martelli, C. M. (2008). Dengue and dengue hemorrhagic fever among adults: Clinical outcomes related to viremia, serotypes, and antibody response. J. Infect. Dis. 197(6):817–824. Guzman, M. G., Kouri, G., Morier, L., Soler, M., and Fernandez, A. (1984). A study of fatal hemorrhagic dengue cases in Cuba, 1981. Bull. Pan. Am. Health Organ. 18(3):213–220. Guzman, M. G., Kouri, G. P., Bravo, J., Soler, M., Vazquez, S., and Morier, L. (1990). Dengue hemorrhagic fever in Cuba, 1981: A retrospective seroepidemiologic study. Am. J. Trop. Med. Hyg. 42(2):179–184. Guzman, M. G., Kouri, G., Bravo, J., Valdes, L., Vazquez, S., and Halstead, S. B. (2002). Effect of age on outcome of secondary dengue 2 infections. Int. J. Infect. Dis. 6(2):118–124. Ha, D. Q., Tien, N. T., Huong, V. T., Loan, H. T., and Thang, C. M. (2000). Dengue epidemic in southern Vietnam, 1998. Emerg. Infect. Dis. 6(4):422–425. Hahn, C. S., Lustig, S., Strauss, E. G., and Strauss, J. H. (1988). Western equine encephalitis virus is a recombinant virus. Proc. Natl. Acad. Sci. USA 85(16):5997–6001. Halstead, S. B. (1979). In vivo enhancement of dengue virus infection in rhesus monkeys by passively transferred antibody. J. Infect. Dis. 140(4):527–533. Halstead, S. B. (1988). Pathogenesis of dengue: Challenges to molecular biology. Science 239 (4839):476–481. Halstead, S. B. (1989). Antibody, macrophages, dengue virus infection, shock, and hemorrhage: A pathogenetic cascade. Rev. Infect. Dis. 11(Suppl. 4):S830–S839. Halstead, S. B. (1997). Epidemiology of dengue and dengue hemorrhagic fever. In ‘‘Dengue and Dengue Hemorrhagic Fever’’, (D. J. Gubler and G. Kuno, Eds.), pp. 23–44. CABI, Oxon. Halstead, S. B., and O’Rourke, E. J. (1977a). Antibody-enhanced dengue virus infection in primate leukocytes. Nature 265(5596):739–741. Halstead, S. B., and O’Rourke, E. J. (1977b). Dengue viruses and mononuclear phagocytes. I. Infection enhancement by non-neutralizing antibody. J. Exp. Med. 146(1):201–217. Halstead, S. B., Voulgaropoulos, E. M., Tien, N. H., and Udomsakdi, S. (1965). Dengue hemorrhagic fever in South Vietnam: Report of the 1963 outbreak. Am. J. Trop. Med. Hyg. 14(5):819–830. Halstead, S. B., Nimmannitya, S., Yamarat, C., and Russell, P. K. (1967). Hemorrhagic fever in Thailand; recent knowledge regarding etiology. Jpn. J. Med. Sci. Biol. 20:96–103. Halstead, S. B., Shotwell, H., and Casals, J. (1973a). Studies on the pathogenesis of dengue infection in monkeys. I. Clinical laboratory responses to primary infection. J. Infect. Dis. 128(1):7–14. Halstead, S. B., Shotwell, H., and Casals, J. (1973b). Studies on the pathogenesis of dengue infection in monkeys. II. Clinical laboratory responses to heterologous infection. J. Infect. Dis. 128(1):15–22. Halstead, S. B., Chow, J. S., and Marchette, N. J. (1973c). Immunological enhancement of dengue virus replication. Nat. New Biol. 243(122):24–26. Halstead, S. B., Marchette, N. J., Sung Chow, J. S., and Lolekha, S. (1976). Dengue virus replication enhancement in peripheral blood leukocytes from immune human beings. Proc. Soc. Exp. Biol. Med. 151(1):136–139. Halstead, S. B., O’Rourke, E. J., and Allison, A. C. (1977). Dengue viruses and mononuclear phagocytes. II. Identity of blood and tissue leukocytes supporting in vitro infection. J. Exp. Med. 146(1):218–229.
History and Evolution of Human Dengue Emergence
63
Halstead, S. B., Porterfield, J. S., and O’Rourke, E. J. (1980). Enhancement of dengue virus infection in monocytes by flavivirus antisera. Am. J. Trop. Med. Hyg. 29(4):638–642. Halstead, S. B., Streit, T. G., Lafontant, J. G., Putvatana, R., Russell, K., Sun, W., KanesaThasan, N., Hayes, C. G., and Watts, D. M. (2001). Haiti: Absence of dengue hemorrhagic fever despite hyperendemic dengue virus transmission. Am. J. Trop. Med. Hyg. 65(3):180–183. Hammon, W. M., Rudnick, A., and Sather, G. (1960a). Viruses associated with epidemic hemorrhagic fevers of the Philippines and Thailand. Science 131:1102–1103. Hammon, W. M., Rudnick, A., Sather, G., Rogers, K. D., and Morse, L. J. (1960b). New hemorrhagic fevers of childern in the Philippines and Thailand. Trans. Assoc. Am. Phys. 73:140–155. Hanley, K. A., Manlucu, L. R., Manipon, G. G., Hanson, C. T., Whitehead, S. S., Murphy, B. R., and Blaney, J. E., Jr. (2004). Introduction of mutations into the nonstructural genes or 30 untranslated region of an attenuated dengue virus type 4 vaccine candidate further decreases replication in rhesus monkeys while retaining protective immunity. Vaccine 22(25–26):3440–3448. Harrington, L. C., Edman, J. D., and Scott, T. W. (2001). Why do female Aedes aegypti (Diptera: Culicidae) feed preferentially and frequently on human blood? J. Med. Entomol. 38 (3):411–422. Harris, E., Videa, E., Perez, L., Sandoval, E., Tellez, Y., Perez, M. L., Cuadra, R., Rocha, J., Idiaquez, W., Alonso, R. E., Delgado, M. A., Campo, L. A., et al. (2000). Clinical, epidemiologic, and virologic features of dengue in the 1998 epidemic in Nicaragua. Am. J. Trop. Med. Hyg. 63(1–2):5–11. Hayes, C. G., Manaloto, C. R., Gonzales, A., and Ranoa, C. P. (1988). Dengue infections in the Philippines: Clinical and virological findings in 517 hospitalized patients. Am. J. Trop. Med. Hyg. 39(1):110–116. Heinz, F. X., MS, C., Purcell, R. H., Gould, E. A., Howard, C. R., Houghton, M., Moormann, R. J. M., Rice, C. M., and Thiel, H. J. (2000). Family flaviviridae. In ‘‘Virus Taxonomy: 7th International Committee for the Taxonomy of Viruses’’, (M. H. V. Regenmortel, C. M. Fauquet, D. H. L. Bishop, E. Carstens, M. K. Estes, S. Lemon, J. Maniloff, M. A. Mayo, D. McGeogh, C. R. Pringle, and R. B. Wickner, Eds.). Academic Press, San Diego. Hervy, J. P., Legros, F., Roche, J. C., Monteny, N., and Diaco, B. (1984). Circulation du Dengue 2 dans plusieurs milieux boises des savanes soudaniennes de la region de Bobo-Dioulasso (Burkina Faso). Considerations entomologiques et epidemiologiques. Cah. ORSTOM. ser Ent. Med. et Parasitol. 22:135–143. Hilgard, P., and Stockert, R. (2000). Heparan sulfate proteoglycans initiate dengue virus infection of hepatocytes. Hepatology 32(5):1069–1077. Hirsch, A. (1883). Dengue, a comparatively new desease: Its symptoms. (Trans. C. Creighton, Ed.). In ‘‘Handbook of Geographical and Historical Pathology’’, Vol. I, pp. 55–81. III vols. Syndenham Society, London. Ho, L. J., Hung, L. F., Weng, C. Y., Wu, W. L., Chou, P., Lin, Y. L., Chang, D. M., Tai, T. Y., and Lai, J. H. (2005). Dengue virus type 2 antagonizes IFN-alpha but not IFN-gamma antiviral effect via down-regulating Tyk2-STAT signaling in the human dendritic cell. J. Immunol. 174(12):8163–8172. Hober, D., Poli, L., Roblin, B., Gestas, P., Chungue, E., Granic, G., Imbert, P., Pecarere, J. L., Vergez-Pascal, R., Wattre, P., and Maniez-Montreuil, M. (1993). Serum levels of tumor necrosis factor-alpha (TNF-alpha), interleukin-6 (IL-6), and interleukin-1 beta (IL-1 beta) in dengue-infected patients. Am. J. Trop. Med. Hyg. 48(3):324–331. Holmes, E. C. (2003). Patterns of intra- and interhost nonsynonymous variation reveal strong purifying selection in dengue virus. J. Virol. 77(20):11296–11298. Holmes, E. C., and Twiddy, S. S. (2003). The origin, emergence and evolutionary genetics of dengue virus. Infect. Genet. Evol. 3(1):19–28.
64
Nikos Vasilakis and Scott C. Weaver
Holmes, E. C., Worobey, M., and Rambaut, A. (1999). Phylogenetic evidence for recombination in dengue virus. Mol. Biol. Evol. 16(3):405–409. Hotta, S. (1952). Experimental studies on dengue. I. Isolation, identification and modification of the virus. J. Infect. Dis. 90(1):1–9. Hotta, H., Murakami, I., Miyasaki, K., Takeda, Y., Shirane, H., and Hotta, S. (1981). Localization of dengue virus in nude mice. Microbiol. Immunol. 25(1):89–93. Huang, Y. H., Lei, H. Y., Liu, H. S., Lin, Y. S., Liu, C. C., and Yeh, T. M. (2000). Dengue virus infects human endothelial cells and induces IL-6 and IL-8 production. Am. J. Trop. Med. Hyg. 63(1–2):71–75. Hughes, A. L. (2001). Evolutionary change of predicted cytotoxic T cell epitopes of dengue virus. Infect. Genet. Evol. 1(2):123–130. Hull, B., Tikasingh, E., de Souza, M., and Martinez, R. (1984). Natural transovarial transmission of dengue 4 virus in Aedes aegypti in Trinidad. Am. J. Trop. Med. Hyg. 33(6):1248–1250. Hung, S. L., Lee, P. L., Chen, H. W., Chen, L. K., Kao, C. L., and King, C. C. (1999). Analysis of the steps involved in Dengue virus entry into host cells. Virology 257(1):156–167. Hung, J. J., Hsieh, M. T., Young, M. J., Kao, C. L., King, C. C., and Chang, W. (2004). An external loop region of domain III of dengue virus type 2 envelope protein is involved in serotype-specific binding to mosquito but not mammalian cells. J. Virol. 78(1):378–388. Hussin, N., Jaafar, J., Naing, N. N., Mat, H. A., Muhamad, A. H., and Mamat, M. N. (2005). A review of dengue fever incidence in Kota Bharu, Kelantan, Malaysia during the years 1998–2003. Southeast Asian J. Trop. Med. Public Health 36(5):1179–1186. Hyams, K. C., Oldfield, E. C., Scott, R. M., Bourgeois, A. L., Gardiner, H., Pazzaglia, G., Moussa, M., Saleh, A. S., Dawi, O. E., and Daniell, F. D. (1986). Evaluation of febrile patients in Port Sudan, Sudan: Isolation of dengue virus. Am. J. Trop. Med. Hyg. 35 (4):860–865. Innis, B. L., Thirawuth, V., and Hemachudha, C. (1989). Identification of continuous epitopes of the envelope glycoprotein of dengue type 2 virus. Am. J. Trop. Med. Hyg. 40(6):676–687. Jarvis, T. C., and Kirkegaard, K. (1992). Poliovirus RNA recombination: Mechanistic studies in the absence of selection. EMBO J. 11(8):3135–3145. Jenkins, G. M., Rambaut, A., Pybus, O. G., and Holmes, E. C. (2002). Rates of molecular evolution in RNA viruses: A quantitative phylogenetic analysis. J. Mol. Evol. 54 (2):156–165. Jindadamrongwech, S., Thepparit, C., and Smith, D. R. (2004). Identification of GRP 78 (BiP) as a liver cell expressed receptor element for dengue virus serotype 2. Arch. Virol. 149 (5):915–927. Johnson, B. K., Ocheng, D., Gichogo, A., Okiro, M., Libondo, D., Kinyanjui, P., and Tukei, P. M. (1982). Epidemic dengue fever caused by dengue type 2 virus in Kenya: Preliminary results of human virological and serological studies. East Afr. Med. J. 59 (12):781–784. Jones, M., Davidson, A., Hibbert, L., Gruenwald, P., Schlaak, J., Ball, S., Foster, G. R., and Jacobs, M. (2005). Dengue virus inhibits alpha interferon signaling by reducing STAT2 expression. J. Virol. 79(9):5414–5420. Joshi, V., Singhi, M., and Chaudhary, R. C. (1996). Transovarial transmission of dengue 3 virus by Aedes aegypti. Trans. R. Soc. Trop. Med. Hyg. 90(6):643–644. Joshi, V., Mourya, D. T., and Sharma, R. C. (2002). Persistence of dengue-3 virus through transovarial transmission passage in successive generations of Aedes aegypti mosquitoes. Am. J. Trop. Med. Hyg. 67(2):158–161. Jumali, S, Gubler, D. J., Nalim, S., Eram, S., and Sulianti Saroso, J. (1979). Epidemic dengue hemorrhagic fever in rural Indonesia. III. Entomological studies. Am. J. Trop. Med. Hyg. 28 (4):717–724. Kalayanarooj, S., and Nimmannitya, S. (2005). Is dengue severity related to nutritional status? Southeast Asian J. Trop. Med. Public Health 36(2):378–384.
History and Evolution of Human Dengue Emergence
65
Kalayanarooj, S., Rimal, H. S., Andjaparidze, A., Vatcharasaevee, V., Nisalak, A., Jarman, R. G., Chinnawirotpisan, P., Mammen, M. P., Holmes, E. C., and Gibbons, R. V. (2007). Clinical intervention and molecular characteristics of a dengue hemorrhagic fever outbreak in Timor Leste, 2005. Am. J. Trop. Med. Hyg. 77(3):534–537. Kamal, H. (1928). The 1927 epidemic of Dengue in Egypt. Br. Med. J. 1:1104–1106. Karabatsos, N. (1985a). Apoi virus. In ‘‘International Catalogue of Arboviruses Including Certain Other Viruses of Vertebrates,’’ 4th Ed., p. 174. American Society of Tropical Medicine and Hygiene, San Antonio. Karabatsos, N. (1985b). Rio Bravo virus. In ‘‘International Catalogue of Arboviruses Including Certain Other Viruses of Vertebrates,’’ 3rd Ed p. 860. American Society of Tropical Medicine and Hygiene, San Antonio. Khan, S. (1913). Dengue at Meerut. Indian Medical Gazette, 48:204. Khin, M. M., and Than, K. A. (1983). Transovarial transmission of dengue 2 virus by Aedes aegypti in nature. Am. J. Trop. Med. Hyg. 32(3):590–594. Kho, L. K., Sumarmo, W. H., Jahja, E. C., and Gubler, D. J. (1981). Dengue hemorrhagic fever accompanied by encephalopathy in Jakarta. Southeast Asian J. Trop. Med. Public Health 12 (1):83–86. King, C. A., Anderson, R., and Marshall, J. S. (2002). Dengue virus selectively induces human mast cell chemokine production. J. Virol. 76(16):8408–8419. King, C. C., Wu, Y. C., Chao, D. Y., Lin, T. H., Chow, L., Ht, W., Ku, C. C., Cl, K., Chien, L. J., Chang, H. J., Huang, J. H., Twu, S. J., et al. (2000). Major epidemics of dengue in Taiwan in 1981–2000: Related to intensive virus activities in Asia. Dengue Bull. 24:1–8. Kliks, S. C., Nisalak, A., Brandt, W. E., Wahl, L., and Burke, D. S. (1989). Antibody-dependent enhancement of dengue virus growth in human monocytes as a risk factor for dengue hemorrhagic fever. Am. J. Trop. Med. Hyg. 40(4):444–451. Klungthong, C., Zhang, C., Mammen, M. P., Jr., Ubol, S., and Holmes, E. C. (2004). The molecular epidemiology of dengue virus serotype 4 in Bangkok, Thailand. Virology 329 (1):168–179. Knowles, N. J., Davies, P. R., Henry, T., O’Donnell, V., Pacheco, J. M., and Mason, P. W. (2001). Emergence in Asia of foot-and-mouth disease viruses with altered host range: Characterization of alterations in the 3A protein. J. Virol. 75(3):1551–1556. Kou, Z., Quinn, M., Chen, H., Rodrigo, W. W., Rose, R. C., Schlesinger, J. J., and Jin, X. (2008). Monocytes, but not T or B cells, are the principal target cells for dengue virus (DV) infection among human peripheral blood mononuclear cells. J. Med. Virol. 80(1):134–146. Kouri, G., Mas, P., Guzman, M. G., Soler, M., Goyenechea, A., and Morier, L. (1983). Dengue hemorrhagic fever in Cuba, 1981: Rapid diagnosis of the etiologic agent. Bull. Pan Am. Health Organ. 17(2):126–132. Kouri, G. P., Guzman, M. G., and Bravo, J. R. (1987). Why dengue haemorrhagic fever in Cuba? Part 2. An integral analysis Trans. R. Soc. Trop. Med. Hyg. 81(5):821–823. Kouri, G. P., Guzman, M. G., Bravo, J. R., and Triana, C. (1989). Dengue haemorrhagic fever/ dengue shock syndrome: Lessons from the Cuban epidemic, 1981. Bull. World Health Organ. 67(4):375–380. Kouri, G., Valdez, M., Arguello, L., Guzman, M. G., Valdes, L., Soler, M., and Bravo, J. (1991). [Dengue epidemic in Nicaragua, 1985]. Rev. Inst. Med. Trop. Sao Paulo 33(5):365–371. Kuno, G. (1995). Review of the factors modulating dengue transmission. Epidemiol. Rev. 17 (2):321–335. Kurane, I., Innis, B. L., Nimmannitya, S., Nisalak, A., Meager, A., Janus, J., and Ennis, F. A. (1991). Activation of T lymphocytes in dengue virus infections. High levels of soluble interleukin 2 receptor, soluble CD4, soluble CD8, interleukin 2, and interferon-gamma in sera of children with dengue. J. Clin. Invest. 88(5):1473–1480. Kurane, I., Janus, J., and Ennis, F. A. (1992). Dengue virus infection of human skin fibroblasts in vitro production of IFN-beta, IL-6 and GM-CSF. Arch. Virol. 124(1–2):21–30.
66
Nikos Vasilakis and Scott C. Weaver
Kurane, I., Innis, B. L., Nimmannitya, S., Nisalak, A., Meager, A., and Ennis, F. A. (1993). High levels of interferon alpha in the sera of children with dengue virus infection. Am. J. Trop. Med. Hyg. 48(2):222–229. Kusriastuti, R., and Sutomo, S. (2005). Evolution of dengue prevention and control programme in Indonesia. Dengue Bull. 29:1–7. Kutubuddin, M., Kolaskar, A. S., Galande, S., Gore, M. M., Ghosh, S. N., and Banerjee, K. (1991). Recognition of helper T cell epitopes in envelope (E) glycoprotein of Japanese encephalitis, west Nile and Dengue viruses. Mol. Immunol. 28(1–2):149–154. Lai, M. M. (1992a). Genetic recombination in RNA viruses. Curr. Top. Microbiol. Immunol. 176:21–32. Lai, M. M. (1992b). RNA recombination in animal and plant viruses. Microbiol. Rev. 56 (1):61–79. Laigret, J., Rosen, L., and Scholammer, G. (1967). [On an epidemic of dengue occurring in Tahiti in 1964. Relations to the ‘‘hemorrhagic fevers’’ of Southeast Asia] Bull. Soc. Pathol. Exot. Filiales 60(14):339–353. Lanciotti, R. S., Lewis, J. G., Gubler, D. J., and Trent, D. W. (1994). Molecular evolution and epidemiology of dengue-3 viruses. J. Gen. Virol. 75(Pt. 1):65–75. Lanciotti, R. S., Gubler, D. J., and Trent, D. W. (1997). Molecular evolution and phylogeny of dengue-4 viruses. J. Gen. Virol. 78(Pt. 9):2279–2284. Lanciotti, R. S., Roehrig, J. T., Deubel, V., Smith, J., Parker, M., Steele, K., Crise, B., Volpe, K. E., Crabtree, M. B., Scherret, J. H., Hall, R. A., MacKenzie, J. S., et al. (1999). Origin of the West Nile virus responsible for an outbreak of encephalitis in the northeastern United States. Science 286(5448):2333–2337. Leclerc, C., Deriaud, E., Megret, F., Briand, J. P., Van Regenmortel, M. H., and Deubel, V. (1993). Identification of helper T cell epitopes of dengue virus E-protein. Mol. Immunol. 30 (7):613–625. Lee, H. L., Mustafakamal, I., and Rohani, A. (1997). Does transovarial transmission of dengue virus occur in Malaysian Aedes aegypti and Aedes albopictus? Southeast Asian J. Trop. Med. Public Health 28(1):230–232. Lee, E., Hall, R. A., and Lobigs, M. (2004). Common E protein determinants for attenuation of glycosaminoglycan-binding variants of Japanese encephalitis and West Nile viruses. J. Virol. 78(15):8271–8280. Lee, E., Wright, P. J., Davidson, A., and Lobigs, M. (2006). Virulence attenuation of Dengue virus due to augmented glycosaminoglycan-binding affinity and restriction in extraneural dissemination. J. Gen. Virol. 87(Pt. 10):2791–2801. Leichtenstern, O. (1896). Influenza and dengue. ‘‘Specielle Pathologie und Therapie’’ (H. Nothnagel ed.) Vol. IV:133–226. Alfred Holder Wien. Leitmeyer, K. C., Vaughn, D. W., Watts, D. M., Salas, R., Villalobos, I., de, C., Ramos, C., and Rico-Hesse, R. (1999). Dengue virus structural differences that correlate with pathogenesis. J. Virol. 73(6):4738–4747. Lewis, J. A., Chang, G. J., Lanciotti, R. S., Kinney, R. M., Mayer, L. W., and Trent, D. W. (1993). Phylogenetic relationships of dengue-2 viruses. Virology 197(1):216–224. Libraty, D. H., Pichyangkul, S., Ajariyakhajorn, C., Endy, T. P., and Ennis, F. A. (2001). Human dendritic cells are activated by dengue virus infection: Enhancement by gamma interferon and implications for disease pathogenesis. J. Virol. 75(8):3501–3508. Libraty, D. H., Endy, T. P., Houng, H. S., Green, S., Kalayanarooj, S., Suntayakorn, S., Chansiriwongs, W., Vaughn, D. W., Nisalak, A., Ennis, F. A., and Rothman, A. L. (2002). Differing influences of virus burden and immune activation on disease severity in secondary dengue-3 virus infections. J. Infect. Dis. 185(9):1213–1221. Lim, T. W., Rudnick, A., and Chan, Y. C. (1961). Recent studies of haemorrhagic fever in Singapore. Singapore Med. J. 2(4):158–161.
History and Evolution of Human Dengue Emergence
67
Lim, T. W., Wallace, H. G., Rudnick, A., Cheong, W. H., Knudsen, A. B., and Chew, V. (1974). The 1973 epidemic of dengue haemorrhagic fever in Malaysia: A preliminary report. Southeast Asian J. Trop. Med. Public Health 5(3):453–454. Lin, Y. L., Lei, H. Y., Lin, Y. S., Yeh, T. M., Chen, S. H., and Liu, H. S. (2002). Heparin inhibits dengue-2 virus infection of five human liver cell lines. Antiviral Res. 56(1):93–96. Lloyd, W., Theiler, M., and Ricci, N. I. (1936). Modification of virulence of yellow fever virus by cultivation in tissues in vitro. Trans. R. Soc. Trop. Med. Hyg. 29:481–529. Loison, G., Rosen, L., Papillaud, J., Tomasini, J., Vaujany, J., and Chanalet, G. (1973). La dengue en Nouvelle-Caledonie (1971–1972). Bull. Soc. Pathol. Exot. Filiales 66:511–519. Lorono Pino, M. A., Farfan Ale, J. A., Rosado Paredes, E. P., Kuno, G., and Gubler, D. J. (1993). Epidemic dengue 4 in the Yucatan, Mexico, 1984. Rev. Inst. Med. Trop. Sao Paulo 35 (5):449–455. Lorono-Pino, M. A., Cropp, C. B., Farfan, J. A., Vorndam, A. V., Rodriguez-Angulo, E. M., Rosado-Paredes, E. P., Flores-Flores, L. F., Beaty, B. J., and Gubler, D. J. (1999). Common occurrence of concurrent infections by multiple dengue virus serotypes. Am. J. Trop. Med. Hyg. 61(5):725–730. Lumsden, L. L. (1958). St. Louis encephalitis in 1933; observations on epidemiological features. Public Health Rep. 73(4):340–353. Lvov, D. K., Timopheeva, A. A., Chervonski, V. I., Gromashevski, V. L., Klisenko, G. A., Gostinshchikova, G. V., and Kostyrko, I. N. (1971). Tuleniy virus. A new Group B arbovirus isolated from Ixodes (Ceratixodes) putus Pick.-Camb. 1878 collected on Tuleniy Island, Sea of Okhotsk. Am. J. Trop. Med. Hyg. 20(3):456–460. Mackerras, I. M. (1946). Transmission of dengue fever by Aedes (Stegomyia) scutellaris walk. in New Guinea. Trans. R. Soc. Trop. Med. Hyg. 40:295–312. Maguire, T., Miles, J. A., Macnamara, F. N., Wilkinson, P. J., Austin, F. J., and Mataika, J. U. (1974). Mosquito-borne infections in Fiji. V. The 1971–73 dengue epidemic. J. Hyg. (Lond.) 73(2):263–270. Malkinson, M., and Banet, C. (2002). The role of birds in the ecology of West Nile virus in Europe and Africa. Curr. Top. Microbiol. Immunol. 267:309–322. Marianneau, P., Megret, F., Olivier, R., Morens, D. M., and Deubel, V. (1996). Dengue 1 virus binding to human hepatoma HepG2 and simian Vero cell surfaces differs. J. Gen. Virol. 77 (Pt. 10):2547–2554. Marianneau, P., Cardona, A., Edelman, L., Deubel, V., and Despres, P. (1997). Dengue virus replication in human hepatoma cells activates NF-kappaB which in turn induces apoptotic cell death. J. Virol. 71(4):3244–3249. Markoff, L., Pang, X., Houng, H. S., Falgout, B., Olsen, R., Jones, E., and Polo, S. (2002). Derivation and characterization of a dengue type 1 host range-restricted mutant virus that is attenuated and highly immunogenic in monkeys. J. Virol. 76(7):3318–3328. Martinez-Barragan, J. J., and del Angel, R. M. (2001). Identification of a putative coreceptor on Vero cells that participates in dengue 4 virus infection. J. Virol. 75(17):7818–7827. Mathew, T., Suri, J. C., Suri, N. K., Bhola, S. R., Arora, R. R., Lal, P., Raichaudhari, A. N., Mathur, K. K., and Gupta, J. P. (1977). Investigation on an epidemic of dengue in Jammu, 1974. Indian J. Med. Res. 65(5):613–622. McLean, D. M., and Donohue, W. L. (1959). Powassan virus: Isolation of virus from a fatal case of encephalitis. Can. Med. Assoc. J. 80(9):708–711. McSherry, J. A. (1982). Some medical aspects of the Darien scheme: Was it dengue? Scott. Med. J. 27(2):183–184. Megret, F., Hugnot, J. P., Falconar, A., Gentry, M. K., Morens, D. M., Murray, J. M., Schlesinger, J. J., Wright, P. J., Young, P., Van Regenmortel, M. H., et al. (1992). Use of recombinant fusion proteins and monoclonal antibodies to define linear and discontinuous antigenic sites on the dengue virus envelope glycoprotein. Virology. 187(2):480–491.
68
Nikos Vasilakis and Scott C. Weaver
Men, R., Bray, M., Clark, D., Chanock, R. M., and Lai, C. J. (1996). Dengue type 4 virus mutants containing deletions in the 30 noncoding region of the RNA genome: Analysis of growth restriction in cell culture and altered viremia pattern and immunogenicity in rhesus monkeys. J. Virol. 70(6):3930–3937. Messer, W. B., Vitarana, U. T., Sivananthan, K., Elvtigala, J., Preethimala, L. D., Ramesh, R., Withana, N., Gubler, D. J., and De Silva, A. M. (2002). Epidemiology of dengue in Sri Lanka before and after the emergence of epidemic dengue hemorrhagic fever. Am. J. Trop. Med. Hyg. 66(6):765–773. Messer, W. B., Gubler, D. J., Harris, E., Sivananthan, K., and de Silva, A. M. (2003). Emergence and global spread of a dengue serotype 3, subtype III virus. Emerg. Infect. Dis. 9 (7):800–809. Metselaar, D., Grainger, C. R., Oei, K. G., Reynolds, D. G., Pudney, M., Leake, C. J., Tukei, P. M., D’Offay, R. M., and Simpson, D. I. (1980). An outbreak of type 2 dengue fever in the Seychelles, probably transmitted by Aedes albopictus (Skuse). Bull. World Health Organ. 58(6):937–943. Mitchell, C. J., and Miller, B. R. (1990). Vertical transmission of dengue viruses by strains of Aedes albopictus recently introduced into Brazil. J. Am. Mosq. Control Assoc. 6(2):251–253. Moncayo, A. C., Fernandez, Z., Ortiz, D., Diallo, M., Sall, A., Hartman, S., Davis, C. T., Coffey, L., Mathiot, C. C., Tesh, R. B., and Weaver, S. C. (2004). Dengue emergence and adaptation to peridomestic mosquitoes. Emerg. Infect. Dis. 10(10):1790–1796. Monlun, E., Zeller, H., Traore-Lamizana, M., Hervy, J. P., Adam, F., Mondo, M., and Digoutte, J. P. (1992). Caracteres cliniques et epidemiologiques de la dengue 2 au Senegal. Med. Mal. Infect. 22:718–721. Morales, A., Groot, H., Russell, P. K., and McCown, J. M. (1973). Recovery of dengue-2 virus from Aedes aegypti in Colombia. Am. J. Trop. Med. Hyg. 22(6):785–787. More, F. W. (1904). Observations on dengue fever in Singapore. J. Malaya Branch Br. Med. Assoc. 1:24–29. Moreau, J. P., Rosen, L., Saugrain, J., and Lagraulet, J. (1973). An epidemic of dengue on Tahiti associated with hemorrhagic manifestations. Am. J. Trop. Med. Hyg. 22(2):237–241. Morens, D. M. (1994). Antibody-dependent enhancement of infection and the pathogenesis of viral disease. Clin. Infect. Dis. 19(3):500–512. Morens, D. M., Rigau-Perez, J. G., Lopez-Correa, R. H., Moore, C. G., Ruiz-Tiben, E. E., Sather, G. E., Chiriboga, J., Eliason, D. A., Casta-Velez, A., and Woodall, J. P. (1986). Dengue in Puerto Rico, 1977: Public health response to characterize and control an epidemic of multiple serotypes. Am. J. Trop. Med. Hyg. 35(1):197–211. Morimoto, K., Hooper, D. C., Carbaugh, H., Fu, Z. F., Koprowski, H., and Dietzschold, B. (1998). Rabies virus quasispecies: Implications for pathogenesis. Proc. Natl. Acad. Sci. USA 95(6):3152–3156. Munoz-Jordan, J. L., Sanchez-Burgos, G. G., Laurent-Rolle, M., and Garcia-Sastre, A. (2003). Inhibition of interferon signaling by dengue virus. Proc. Natl. Acad. Sci. USA 100 (24):14333–14338. Munoz-Jordan, J. L., Laurent-Rolle, M., Ashour, J., Martinez-Sobrido, L., Ashok, M., Lipkin, W. I., and Garcia-Sastre, A. (2005). Inhibition of alpha/beta interferon signaling by the NS4B protein of flaviviruses. J. Virol. 79(13):8004–8013. Murgue, B., Roche, C., Chungue, E., and Deparis, X. (2000). Prospective study of the duration and magnitude of viraemia in children hospitalised during the 1996–1997 dengue-2 outbreak in French Polynesia. J. Med. Virol. 60(4):432–438. Myat Thu, H., Lowry, K., Jiang, L., Hlaing, T., Holmes, E. C., and Aaskov, J. (2005). Lineage extinction and replacement in dengue type 1 virus populations are due to stochastic events rather than to natural selection. Virology 336(2):163–172. Myers, R. M., Carey, D. E., and Rodriques, F. M. (1965). Experiences with the isolation of dengue virus types 1, 2 and 4 from human blood. Indian J. Med. Res. 53:191–198.
History and Evolution of Human Dengue Emergence
69
Myers, R. M., Carey, D. E., Banerjee, K., Reuben, R., and Ramamurti, D. V. (1968). Recovery of dengue type 3 virus from human serum and Aedes aegypti in South India. Indian J. Med. Res. 56(6):781–787. Navarro-Sanchez, E., Altmeyer, R., Amara, A., Schwartz, O., Fieschi, F., Virelizier, J. L., Arenzana-Seisdedos, F., and Despres, P. (2003). Dendritic-cell-specific ICAM3-grabbing non-integrin is essential for the productive infection of human dendritic cells by mosquito-cell-derived dengue viruses. EMBO Rep. 4(7):723–728. Neff, J. M., Morris, L., Gonzalez-Alcover, R., Coleman, P. H., Lyss, S. B., and Negron, H. (1967). Dengue fever in a Puerto Rican community. Am. J. Epidemiol. 86(1):162–184. Nogueira, R. M., Miagostovich, M. P., de Filippis, A. M., Pereira, M. A., and Schatzmayr, H. G. (2001). Dengue virus type 3 in Rio de Janeiro, Brazil. Mem. Inst. Oswaldo Cruz 96(7):925–926. Novella, I. S., Hershey, C. L., Escarmis, C., Domingo, E., and Holland, J. J. (1999). Lack of evolutionary stasis during alternating replication of an arbovirus in insect and mammalian cells. J. Mol. Biol. 287(3):459–465. Okuno, Y., Fukunaga, T., Srisupaluck, S., Kasemsarn, P., Dharakul, C., and Sangkawibha, N. (1980). Serological and virological studies on patients with dengue hemorrhagic fever (DHF) in Chanthaburi province, Thailand. I. Serological studies on paired sera from DHF patients by neutralization (N), hemagglutination inhibition (HI) and staining tests. Biken J. 23(3):113–121. Ooi, E.-E., Goh, K.-T., and Gubler, D. J. (2006). Dengue prevention and 35 years vector control in Singapore. Emerg. Infect. Dis. 12(6):887–893. O’Sullivan, M. A., and Killen, H. M. (1994). The differentiation state of monocytic cells affects their susceptibility to infection and the effects of infection by dengue virus. J. Gen. Virol. 75(Pt. 9):2387–2392. Padbidri, V. S., Adhikari, P., Thakare, J. P., Ilkal, M. A., Joshi, G. D., Pereira, P., Guttikar, S. N., Walhekar, B. D., Chowta, N., and Hegde, B. M. (1995). The 1993 epidemic of dengue fever in Mangalore, Karnataka state, India. Southeast Asian J. Trop. Med. Public Health 26(4):699–704. PAHO. (2007). Number of reported cases of dengue & dengue hemorrhagic fever (DHF), region of the Americas (by country and subregion). Available at http://www.paho.org/ English/AD/DPC/CD/dengue-cases-2007.htm. Pandey, B., Rai, S., Morita, K., and Kurane, I. (2004). First case of dengue virus infection in Nepal. Nepal Med. Coll. J. 6(2):157–159. Papaevangelou, G., and Halstead, S. B. (1977). Infections with two dengue viruses in Greece in the 20th century. Did dengue hemorrhagic fever occur in the 1928 epidemic? J. Trop. Med. Hyg. 80(3):46–51. Pattnaik, P., Babu, J. P., Verma, S. K., Tak, V., and Rao, P. V. (2007). Bacterially expressed and refolded envelope protein (domain III) of dengue virus type-4 binds heparan sulfate. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci. 846(1–2):184–194. Peiris, J. S., Dittus, W. P., and Ratnayake, C. B. (1993). Seroepidemiology of dengue and other arboviruses in a natural population of toque macaques (Macaca sinica) at Polonnaruwa, Sri Lanka. J. Med. Primatol. 22(4):240–245. Pepper, P. (1941). A note on David Bylon and dengue. Ann. Med. Hist. 3:363–368. Pinheiro, F. P. (1989). Dengue in the Americas. 1980–1987. Epidemiol. Bull. 10(1):1–8. Platt, K. B., Linthicum, K. J., Myint, K. S., Innis, B. L., Lerdthusnee, K., and Vaughn, D. W. (1997). Impact of dengue virus infection on feeding behavior of Aedes aegypti. Am. J. Trop. Med. Hyg. 57(2):119–125. Pryor, M. J., Carr, J. M., Hocking, H., Davidson, A. D., Li, P., and Wright, P. J. (2001). Replication of dengue virus type 2 in human monocyte-derived macrophages: Comparisons of isolates and recombinant viruses with substitutions at amino acid 390 in the envelope glycoprotein. Am. J. Trop. Med. Hyg. 65(5):427–434.
70
Nikos Vasilakis and Scott C. Weaver
Putnam, J. L., and Scott, T. W. (1995a). Blood-feeding behavior of dengue-2 virus-infected Aedes aegypti. Am. J. Trop. Med. Hyg. 52(3):225–227. Putnam, J. L., and Scott, T. W. (1995b). The effect of multiple host contacts on the infectivity of dengue-2 virus-infected Aedes aegypti. J. Parasitol. 81(2):170–174. Pybus, O. G., Rambaut, A., Belshaw, R., Freckleton, R. P., Drummond, A. J., and Holmes, E. C. (2007). Phylogenetic evidence for deleterious mutation load in RNA viruses and its contribution to viral evolution. Mol. Biol. Evol. 24(3):845–852. Qiu, F. X., Zhang, H. Y., Shao, L., Li, X. D., Luo, H. R., and Yu, Y. X. (1981). Studies on the rapid detection of Dengue virus antigen by immunofluorescence and radioimmunoassay. Chin. Med. J. (Engl.) 94(10):653–658. Ramakrishnan, S. P., Gelfand, H. M., Bose, P. N., Sehgal, P. N., and Mukherjee, R. N. (1964). The epidemic of acute haemorrhagic fever, Calcutta, 1963: Epidemiological inquiry. Indian J. Med. Res. 52:633–650. Rambaut, A. (2000). Estimating the rate of molecular evolution: Incorporating noncontemporaneous sequences into maximum likelihood phylogenies. Bioinformatics 16 (4):395–399. Rathavuth, H., Vaughn, D. W., Minn, K., Nimmannitya, S., Nisalak, A., Raengsakulrach, B., Rorabaugh, M. L., Yuvatha, K., and Sophal, O. (1997). Hemorrhagic fever in Cambodia is caused by dengue viruses: Evidence for transmission of all four serotypes. Southeast Asian J. Trop. Med. Public Health 28(1):120–125. Ravi, V., Desai, A. S., Shenoy, P. K., Satishchandra, P., Chandramuki, A., and GourieDevi, M. (1993). Persistence of Japanese encephalitis virus in the human nervous system. J. Med. Virol. 40(4):326–329. Reed, W., and Carroll, J. (1902). The etiology of yellow fever, a supplementary note. Am. Med. 3:301–305. Reed, W., Carroll, J., Agramonte, A., and Lazear, J. W. (1900). The etiology of yellow fever. A preliminary note. Philad. Med. J. 6:790–796. Repik, P. M., Dalrymple, J. M., Brandt, W. E., McCown, J. M., and Russell, P. K. (1983). RNA fingerprinting as a method for distinguishing dengue 1 virus strains. Am. J. Trop. Med. Hyg. 32(3):577–589. Reynes, J. M., Laurent, A., Deubel, V., Telliam, E., and Moreau, J. P. (1994). The first epidemic of dengue hemorrhagic fever in French Guiana. Am. J. Trop. Med. Hyg. 51(5):545–553. Rico-Hesse, R. (1990). Molecular evolution and distribution of dengue viruses type 1 and 2 in nature. Virology 174(2):479–493. Rico-Hesse, R., Harrison, L. M., Salas, R. A., Tovar, D., Nisalak, A., Ramos, C., Boshell, J., de Mesa, M. T., Nogueira, R. M., and da Rosa, A. T. (1997). Origins of dengue type 2 viruses associated with increased pathogenicity in the Americas. Virology 230(2):244–251. Robert Putnak, J., Coller, B. A., Voss, G., Vaughn, D. W., Clements, D., Peters, I., Bignami, G., Houng, H. S., Chen, R. C., Barvir, D. A., Seriwatana, J., Cayphas, S., et al. (2005). An evaluation of dengue type-2 inactivated, recombinant subunit, and live-attenuated vaccine candidates in the rhesus macaque model. Vaccine 23(35):4442–4452. Roberts, D. R., Peyton, E. L., Pinheiro, F. P., Balderrama, F., and Vargas, R. (1984). Associations of arbovirus vectors with gallery forests and domestic environments in southeastern Bolivia. Bull. Pan Am. Health Organ. 18(4):337–350. Robin, Y., Cornet, M., Heme, G., and Le Gonidec, G. (1980). Isolement du virus dela dengue au Senegal. Ann. Virol. (Institut Pasteur) 131:149–154. Rocco, I. M., Kavakama, B. B., and Santos, C. L. (2001). First isolation of dengue 3 in Brazil from an imported case. Rev. Inst. Med. Trop. Sao Paulo 43(1):55–57. Roche, J. C., Cordellier, R., Hervy, J. P., Digoutte, J. P., and Monteny, N. (1983). Isolement de 96 souches de virus Dengue 2 a partir de moustiques captures en Cote d’Ivoire et en Haute Volta. Ann. Virol. (Institut Pasteur) 134E:233–244.
History and Evolution of Human Dengue Emergence
71
Rodhain, F. (1991). The role of monkeys in the biology of dengue and yellow fever. Comp. Immunol. Microbiol. Infect. Dis. 14(1):9–19. Rodier, G. R., Gubler, D. J., Cope, S. E., Cropp, C. B., Soliman, A. K., Polycarpe, D., Abdourhaman, M. A., Parra, J. P., Maslin, J., and Arthur, R. R. (1996). Epidemic dengue 2 in the city of Djibouti 1991–1992. Trans. R. Soc. Trop. Med. Hyg. 90(3):237–240. Roehrig, J. T., Risi, P. A., Brubaker, J. R., Hunt, A. R., Beaty, B. J., Trent, D. W., and Mathews, J. H. (1994). T-helper cell epitopes on the E-glycoprotein of dengue 2 Jamaica virus. Virology 198(1):31–38. Rosen, L. (1958a). Experimental infection of New World monkeys with dengue and yellow fever viruses. Am. J. Trop. Med. Hyg. 7(4):406–410. Rosen, L. (1958b). Observations on the epidemilogy of dengue in Panama. Am. J. Hyg. 68(1):45–58. Rosen, L. (1977). The Emperor’s New Clothes revisited, or reflections on the pathogenesis of dengue hemorrhagic fever. Am. J. Trop. Med. Hyg. 26(3):337–343. Rosen, L. (1986). The pathogenesis of dengue haemorrhagic fever. A critical appraisal of current hypotheses. S. Afr. Med. J. (Suppl.): 40–42. Rosen, L. (1987). Sexual transmission of dengue viruses by Aedes albopictus. Am. J. Trop. Med. Hyg. 37(2):398–402. Rosen, L. (1988). Further observations on the mechanism of vertical transmission of flaviviruses by Aedes mosquitoes. Am. J. Trop. Med. Hyg. 39(1):123–126. Rosen, L., Rozeboom, L. E., Sweet, B. H., and Sabin, A. B. (1954). The transmission of dengue by Aedes polynesiensis marks. Am. J. Trop. Med. Hyg. 3(5):878–882. Rosen, L., Shroyer, D. A., Tesh, R. B., Freier, J. E., and Lien, J. C. (1983). Transovarial transmission of dengue viruses by mosquitoes: Aedes albopictus and Aedes aegypti. Am. J. Trop. Med. Hyg. 32(5):1108–1119. Rosen, L., Roseboom, L. E., Gubler, D. J., Lien, J. C., and Chaniotis, B. N. (1985). Comparative susceptibility of mosquito species and strains to oral and parenteral infection with dengue and Japanese encephalitis viruses. Am. J. Trop. Med. Hyg. 34(3):603–615. Rudnick, A. (1965). Studies of the ecology of dengue in Malaysia: A preliminary report. J. Med. Entomol. 2(2):203–208. Rudnick, A. (1984). ‘‘Proceedings of the International Conference on Dengue/DHF.’’ Kuala Lampur, Malaysia. Rudnick, A. (1986). Dengue virus ecology in Malaysia. Inst. Med. Res. Malays. Bull. 23:51–152. Rudnick, A., and Lim, T. W. (1986). Dengue fever studies in Malaysia. Inst. Med. Res. Malays. Bull. 23:51–152. Rudnick, A., Tan, E. E., Lucas, J. K., and Omar, M. B. (1965). Mosquito-borne haemorrhagic fever in Malaya. Br. Med. J. 5445:1269–1272. Rudnick, A., Marchette, N. J., and Garcia, R. (1967). Possible jungle dengue—recent studies and hypotheses. Jpn. J. Med. Sci. Biol. 20:69–74. Rush, A. B. (1789). An account of the bilious remitting fever, as it appeared in Philadelphia in the summer and autumn of the year 1980. In ‘‘Medical Inquiries and Observations,’’ pp. 89–100. Prichard and Hall, Philadelphia. Russell, P. K., Buescher, E. L., McCown, J. M., and Ordonez, J. (1966). Recovery of dengue viruses from patients during epidemics in Puerto Rico and East Pakistan. Am. J. Trop. Med. Hyg. 15(4):573–579. Russell, P. K., Udomsakdi, S., and Halstead, S. B. (1967). Antibody response in dengue and dengue hemorrhagic fever. Jpn. J. Med. Sci. Biol. 20:103–108. Sabin, A. B. (1952). Research on dengue during World War II. Am. J. Trop. Med. Hyg. 1(1):30–50. Sabin, A. B., and Schlesinger, M. C. (1945). Production of immunity to dengue with virus modified by propagation in mice. Science 101:640–642.
72
Nikos Vasilakis and Scott C. Weaver
Salda, L. T., Parquet, M. D., Matias, R. R., Natividad, F. F., Kobayashi, N., and Morita, K. (2005). Molecular epidemiology of dengue 2 viruses in the Philippines: Genotype shift and local evolution. Am. J. Trop. Med. Hyg. 73(4):796–802. Saleh, A. S., Hassan, A., Scott, R. M., Mellick, P. W., Oldfield, E. C., III, and Podgore, J. K. (1985). Dengue in north-east Africa. Lancet 2(8448):211–212. Saluzzo, J. F., Cornet, M., Adam, C., Eyraud, M., and Digoutte, J. P. (1986a). [Dengue 2 in eastern Senegal: Serologic survey in simian and human populations. 1974–85]. Bull. Soc. Pathol. Exot. Filiales 79(3):313–322. Saluzzo, J. F., Cornet, M., Castagnet, P., Rey, C., and Digoutte, J. P. (1986b). Isolation of dengue 2 and dengue 4 viruses from patients in Senegal. Trans. R. Soc. Trop. Med. Hyg. 80(1):5. Sanchez, I. J., and Ruiz, B. H. (1996). A single nucleotide change in the E protein gene of dengue virus 2 Mexican strain affects neurovirulence in mice. J. Gen. Virol. 77(Pt. 10):2541–2545. Saugrain, J., Rosen, L., Outin-Fabre, D., and Moreau, J. P. (1970). [A recent epidemic due to arbovirus infections of the dengue type in Tahiti. Comparative study of the 1964 epidemic]. Bull. Soc. Pathol. Exot. Filiales 63(6):636–642. Schlesinger, J. J., Chapman, S., Nestorowicz, A., Rice, C. M., Ginocchio, T. E., and Chambers, T. J. (1996). Replication of yellow fever virus in the mouse central nervous system: Comparison of neuroadapted and non-neuroadapted virus and partial sequence analysis of the neuroadapted strain. J. Gen. Virol. 77(Pt. 6):1277–1285. Sharma, S., Mathur, A., Prakash, V., Kulshreshtha, R., Kumar, R., and Chaturvedi, U. C. (1991). Japanese encephalitis virus latency in peripheral blood lymphocytes and recurrence of infection in children. Clin. Exp. Immunol. 85(1):85–89. Sharp, T. W., Wallace, M. R., Hayes, C. G., Sanchez, J. L., DeFraites, R. F., Arthur, R. R., Thornton, S. A., Batchelor, R. A., Rozmajzl, P. J., Hanson, R. K., et al. (1995). Dengue fever in U.S. troops during Operation Restore Hope, Somalia, 1992–1993. Am. J. Trop. Med. Hyg. 53(1):89–94. Shope, R. E. (2003). Epidemiology of other arthropod-borne flaviviruses infecting humans. In ‘‘The Flaviviruses: Detection, Diagnosis and Vaccine Development’’ (T. J. Chambers and T. P. Monath, Eds.), pp. 373–391. Elsevier, San Diego, CA. Shresta, S., Sharar, K. L., Prigozhin, D. M., Beatty, P. R., and Harris, E. (2006). Murine model for dengue virus-induced lethal disease with increased vascular permeability. J. Virol. 80 (20):10208–10217. Shroyer, D. A. (1990). Vertical maintenance of dengue-1 virus in sequential generations of Aedes albopictus. J. Am. Mosq. Control Assoc. 6(2):312–314. Siler, J. F., Hall, M. W., and Hitchens, A. P. (1926). Dengue: Its history, epidemiology, mechanism of transmission, etiology, clinical manifestations, immunity, and prevention. Philippine J. Sci. 29:1–252. Simmons, J. S., St John, J. H., and Reynolds, F. H. K. (1931). Experimental studies of dengue. Philippine J. Sci. 44:1–252. Siqueira, J. B., Jr., Martelli, C. M., Coelho, G. E., Simplicio, A. C., and Hatch, D. L. (2005). Dengue and dengue hemorrhagic fever, Brazil, 1981–2002. Emerg. Infect. Dis. 11 (1):48–53. Sittisombut, N., Sistayanarain, A., Cardosa, M. J., Salminen, M., Damrongdachakul, S., Kalayanarooj, S., Rojanasuphot, S., Supawadee, J., and Maneekarn, N. (1997). Possible occurrence of a genetic bottleneck in dengue serotype 2 viruses between the 1980 and 1987 epidemic seasons in Bangkok, Thailand. Am. J. Trop. Med. Hyg. 57(1):100–108. Skae, F. M. T. (1902). Dengue fever in Penang. Br. Med. J. 2:1581–1582. Smith, C. E. (1956). The history of dengue in tropical Asia and its probable relationship to the mosquito Aedes aegypti. J. Trop. Med. Hyg. 59(10):243–251.
History and Evolution of Human Dengue Emergence
73
Smith, C. E. (1958). The distribution of antibodies to Japanese Encephalitis, dengue, and yellow fever viruses in five rural communities in Malaya. Trans. R. Soc. Trop. Med. Hyg. 52 (3):237–252. Spence, L., Jonkers, A. H., and Casals, J. (1969). Dengue type 3 virus isolated from an antiguan patient during the 1963–64 Caribbean epidemic. Am. J. Trop. Med. Hyg. 18 (4):584–587. Stanton, A. T. (1919). The mosquitoes of Far Eastern ports with special reference to the prevalence of Stegomyia fasciata. Bull. Ent. Res. 10:333–344. Stanton, A. T. (1920). The mosquitoes of the Far East ports with special referenceto the prevalence of Stegomyia fasciata. Bull. Ent. Res. 10:333–346. Steadman, G. W. (1828). Some account of an anomalous disease which raged in the islands of St. Thomas and Santa Cruz, in the West Indies, during the months of September, October, November, December, and January 1827–8. Edinb. Med. Surg. J. 30(97):227–248. St George, T. D., Standfast, H. A., Doherty, R. L., Carley, J. G., Fillipich, C., and Brandsma, J. (1977). The isolation of Saumarez Reef virus, a new flavivirus, from bird ticks Ornithodoros capensis and Ixodes eudyptidis in Australia. Aust. J. Exp. Biol. Med. Sci. 55(5):493–499. Strobel, M., Jattiot, F., Boulard, F., Lamaury, I., Salin, J., Jarrige, B., Mazille, V., Vachon, F., and Goursaud, R. (1998). [Emergence of dengue hemorrhagic fever in French Antilles. 3 initial fatal cases in Guadeloupe]. Presse Med. 27(27):1376–1378. Sudiro, T. M., Zivny, J., Ishiko, H., Green, S., Vaughn, D. W., Kalayanarooj, S., Nisalak, A., Norman, J. E., Ennis, F. A., and Rothman, A. L. (2001). Analysis of plasma viral RNA levels during acute dengue virus infection using quantitative competitor reverse transcription-polymerase chain reaction. J. Med. Virol. 63(1):29–34. Sulkin, S. E., Burns, K. F., Shelton, D. F., and Wallis, C. (1962). Bat salivary gland virus: Infections of man and monkey. Tex. Rep. Biol. Med. 20:113–127. Swanepoel, R. (1994). Louping ill. In ‘‘Infectious Diseases of Livestock’’ (J. A. W. Coetzer, G. R. Thomson, and R. C. Tustin, Eds.), pp. 671–677. Oxford University Press. Swanepoel, R., and Coetzer, J. A. W. (1994). Wesselsbron disease. In ‘‘Infectious Diseases of Livestock’’ (J. A. W. Coetzer, G. R. Thomson, and R. C. Tustin, Eds.), pp. 663–670. Oxford University Press, Oxford. Tabachnick, W. J., and Powell, J. R. (1979). A world-wide survey of genetic variation in the yellow fever mosquito, Aedes aegypti. Genet. Res. 34(3):215–229. Tassaneetrithep, B., Burgess, T. H., Granelli-Piperno, A., Trumpfheller, C., Finke, J., Sun, W., Eller, M. A., Pattanapanyasat, K., Sarasombath, S., Birx, D. L., Steinman, R. M., Schlesinger, S., et al. (2003). DC-SIGN (CD209) mediates dengue virus infection of human dendritic cells. J. Exp. Med. 197(7):823–829. Tesh, R. B., Chaniotis, B. N., and Johnson, K. M. (1972). Vesicular stomatitis virus (Indiana serotype): Transovarial transmission by phlebotomine sandlies. Science 175 (29):1477–1479. Thaung, U., Ming, C. K., and Thein, M. (1975). Dengue haemorrhagic fever in Burma. Southeast Asian J. Trop. Med. Public Health 6(4):580–591. Thavara, U., Siriyasatien, P., Tawatsin, A., Asavadachanukorn, P., Anantapreecha, S., Wongwanich, R., and Mulla, M. S. (2006). Double infection of heteroserotypes of dengue viruses in field populations of Aedes aegypti and Aedes albopictus (Diptera: Culicidae) and serological features of dengue viruses found in patients in southern Thailand. Southeast Asian J. Trop. Med. Public Health 37(3):468–476. Theiler, M., and Downs, W. G. (1973). Group B viruses. In ‘‘The Arthropod-Borne Viruses of Vertebrates,’’ pp. 147–208. Yale University Press, New Haven. Theobald, F. V. (1901). In ‘‘A Monograph of the Culicidae of the World.’’ Vol. 1, British Museum of Natural History, London.
74
Nikos Vasilakis and Scott C. Weaver
Thisyakorn, U., and Nimmannitya, S. (1993). Nutritional status of children with dengue hemorrhagic fever. Clin. Infect. Dis. 16(2):295–297. Thomas, S. J., Strickman, D., and Vaughn, D. W. (2003). Dengue epidemiology: Virus epidemiology, ecology, and emergence. Adv. Virus Res. 61:235–289. Tolou, H. J., Couissinier-Paris, P., Durand, J. P., Mercier, V., de Pina, J. J., de Micco, P., Billoir, F., Charrel, R. N., and de Lamballerie, X. (2001). Evidence for recombination in natural populations of dengue virus type 1 based on the analysis of complete genome sequences. J. Gen. Virol. 82(Pt. 6):1283–1290. Traore-Lamizana, M., Zeller, H., Monlun, E., Mondo, M., Hervy, J. P., Adam, F., and Digoutte, J. P. (1994). Dengue 2 outbreak in southeastern Senegal during 1990: Virus isolations from mosquitoes (Diptera: Culicidae). J. Med. Entomol. 31(4):623–627. Trent, D. W., Manske, C. L., Fox, G. E., Chu, M. C., Kliks, S. C., and Monath, T. P. (1990). The molecular epidemiology of dengue viruses: Genetic variation and microevolution. Appl. Virol. Res. 2:293–315. Tsai, T. F., and Mitchell, C. J. (1989). St. louis encephalitis. In ‘‘The Arboviruses: Epidemiology and Ecology’’ (T. P. Monath, Ed.), pp. 431–458. CRC Press, Boca Raton, Florida. Twiddy, S. S., Farrar, J. J., Vinh Chau, N., Wills, B., Gould, E. A., Gritsun, T., Lloyd, G., and Holmes, E. C. (2002a). Phylogenetic relationships and differential selection pressures among genotypes of dengue-2 virus. Virology 298(1):63–72. Twiddy, S. S., Woelk, C. H., and Holmes, E. C. (2002b). Phylogenetic evidence for adaptive evolution of dengue viruses in nature. J. Gen. Virol. 83(Pt. 7):1679–1689. Twiddy, S. S., Holmes, E. C., and Rambaut, A. (2003). Inferring the rate and time-scale of dengue virus evolution. Mol. Biol. Evol. 20(1):122–129. Ungchusak, K., and Kunasol, P. (1988). Dengue haemorrhagic fever in Thailand, 1987. Southeast Asian J. Trop. Med. Public Health 19(3):487–490. University of Ibadan Arbovirus Research Project Annual Report (1969). Group B Arboviruses, pp. 152–162. Anonymous. Uzcategui, N. Y., Camacho, D., Comach, G., Cuello de Uzcategui, R., Holmes, E. C., and Gould, E. A. (2001). Molecular epidemiology of dengue type 2 virus in Venezuela: Evidence for in situ virus evolution and recombination. J. Gen. Virol. 82(Pt. 12):2945–2953. Uzcategui, N. Y., Comach, G., Camacho, D., Salcedo, M., Cabello de Quintana, M., Jimenez, M., Sierra, G., Cuello de Uzcategui, R., James, W. S., Turner, S., Holmes, E. C., and Gould, E. A. (2003). Molecular epidemiology of dengue virus type 3 in Venezuela. J. Gen. Virol. 84(Pt. 6):1569–1575. Varelas-Wesley, I., and Calisher, C. H. (1982). Antigenic relationships of flaviviruses with undetermined arthropod-borne status. Am. J. Trop. Med. Hyg. 31(6):1273–1284. Vasilakis, N., Holmes, E. C., Fokam, E. B., Faye, O., Diallo, M., Sall, A. A., and Weaver, S. C. (2007a). Evolutionary processes among sylvatic Dengue-2 viruses. J. Virol. 81:9591–9595. Vasilakis, N., Shell, E. J., Fokam, E. B., Mason, P. W., Hanley, K. A., Estes, D. M., and Weaver, S. C. (2007b). Potential of ancestral sylvatic dengue-2 viruses to re-emerge. Virology 358(2):402–412. Vasilakis, N., Durbin, A., Travassos da Rosa, A. P. A., Munoz-Jordan, J. L., Tesh, R. B., and Weaver, S. C. (2008a). Antigenic relationships between sylvatic and endemic dengue viruses. Am. J. Trop. Med. Hyg. 79(1):128–132. Vasilakis, N., Tesh, R. B., and Weaver, S. C. (2008b). Sylvatic dengue virus type 2 activity in humans, Nigeria, 1966. Emerg. Infect. Dis. 14(3):502–504. Vasilakis, N., Fokam, E. B., Hanson, C. T., Weinberg, E., Sall, A. A., Whitehead, S. S., Hanley, K. A., and Weaver, S. C. (2008c). Genetic and phenotypic characterization of sylvatic dengue virus type 2 strains. Virology 377(2):296–307. Vaughn, D. W., Green, S., Kalayanarooj, S., Innis, B. L., Nimmannitya, S., Suntayakorn, S., Rothman, A. L., Ennis, F. A., and Nisalak, A. (1997). Dengue in the early febrile phase: Viremia and antibody responses. J. Infect. Dis. 176(2):322–330.
History and Evolution of Human Dengue Emergence
75
Vaughn, D. W., Green, S., Kalayanarooj, S., Innis, B. L., Nimmannitya, S., Suntayakorn, S., Endy, T. P., Raengsakulrach, B., Rothman, A. L., Ennis, F. A., and Nisalak, A. (2000). Dengue viremia titer, antibody response pattern, and virus serotype correlate with disease severity. J. Infect. Dis. 181(1):2–9. Vazeille, M., Mousson, L., Rakatoarivony, I., Villeret, R., Rodhain, F., Duchemin, J. B., and Failloux, A. B. (2001). Population genetic structure and competence as a vector for dengue type 2 virus of Aedes aegypti and Aedes albopictus from Madagascar. Am. J. Trop. Med. Hyg. 65(5):491–497. Vazeille, M., Rosen, L., Mousson, L., and Failloux, A. B. (2003). Low oral receptivity for dengue type 2 viruses of Aedes albopictus from Southeast Asia compared with that of Aedes aegypti. Am. J. Trop. Med. Hyg. 68(2):203–208. Ventura, A. K., and Ehrenkranz, N. J. (1976). Endemic dengue virus infection in Hispaniola. I. Haiti. J. Infect. Dis. 134(5):436–441. Wallace, H. G., Lim, T. W., Rudnick, A., Knudsen, A. B., Cheong, W. H., and Chew, V. (1980). Dengue hemorrhagic fever in Malaysia: The 1973 epidemic. Southeast Asian J. Trop. Med. Public Health 11(1):1–13. Wang, E., Ni, H., Xu, R., Barrett, A. D., Watowich, S. J., Gubler, D. J., and Weaver, S. C. (2000). Evolutionary relationships of endemic/epidemic and sylvatic dengue viruses. J. Virol. 74(7):3227–3234. Wang, W. K., Chao, D. Y., Kao, C. L., Wu, H. C., Liu, Y. C., Li, C. M., Lin, S. C., Ho, S. T., Huang, J. H., and King, C. C. (2003). High levels of plasma dengue viral load during defervescence in patients with dengue hemorrhagic fever: Implications for pathogenesis. Virology 305(2):330–338. Wang, W. K., Chen, H. L., Yang, C. F., Hsieh, S. C., Juan, C. C., Chang, S. M., Yu, C. C., Lin, L. H., Huang, J. H., and King, C. C. (2006). Slower rates of clearance of viral load and virus containing immune complexes in patients with dengue hemorrhagic fever. Clin. Infect. Dis. 43:1023–1030. Watts, D. M., Pantuwatana, S., DeFoliart, G. R., Yuill, T. M., and Thompson, W. H. (1973). Transovarial transmission of LaCrosse virus (California encephalitis group) in the mosquito, Aedes triseriatus. Science. 182(117):1140–1141. Watts, D. M., Harrison, B. A., Pantuwatana, S., Klein, T. A., and Burke, D. S. (1985). Failure to detect natural transovarial transmission of dengue viruses by Aedes aegypti and Aedes albopictus (Diptera: Culicidae). J. Med. Entomol. 22(3):261–265. Watts, D. M., Porter, K. R., Putvatana, P., Vasquez, B., Calampa, C., Hayes, C. G., and Halstead, S. B. (1999). Failure of secondary infection with American genotype dengue 2 to cause dengue haemorrhagic fever. Lancet 354(9188):1431–1434. Weaver, S. C., Rico-Hesse, R., and Scott, T. W. (1992). Genetic diversity and slow rates of evolution in New World alphaviruses. Curr. Top. Microbiol. Immunol. 176:99–117. Weaver, S. C., Brault, A. C., Kang, W., and Holland, J. J. (1999). Genetic and fitness changes accompanying adaptation of an arbovirus to vertebrate and invertebrate cells. J. Virol. 73 (5):4316–4326. White, R. S. (1934). Three years mosquito control work in Calcutta. Bull. Ent. Res. 25:551–573. Whitehead, S. S., Hanley, K. A., Blaney, J. E., Jr., Gilmore, L. E., Elkins, W. R., and Murphy, B. R. (2003). Substitution of the structural genes of dengue virus type 4 with those of type 2 results in chimeric vaccine candidates which are attenuated for mosquitoes, mice, and rhesus monkeys. Vaccine 21(27–30):4307–4316. WHO. (2006). Trend of dengue case and CFR in SEAR countries (W.R.O.f.S.E. Asia, Ed.), Vol. 2008 WHO. WHO. (2007). ‘‘Situation of dengue/dengue haemorrhagic fever in the south-east Asia region,’’ Vol. 2008 http://www.searo.who.int/en/Section10/Section332_1098.htm. Wills, B. A., Oragui, E. E., Stephens, A. C., Daramola, O. A., Dung, N. M., Loan, H. T., Chau, N. V., Chambers, M., Stepniewska, K., Farrar, J. J., and Levin, M. (2002).
76
Nikos Vasilakis and Scott C. Weaver
Coagulation abnormalities in dengue hemorrhagic Fever: Serial investigations in 167 Vietnamese children with Dengue shock syndrome. Clin. Infect. Dis. 35(3):277–285. Wittke, V., Robb, T. E., Thu, H. M., Nisalak, A., Nimmannitya, S., Kalayanrooj, S., Vaughn, D. W., Endy, T. P., Holmes, E. C., and Aaskov, J. G. (2002). Extinction and rapid emergence of strains of dengue 3 virus during an interepidemic period. Virology 301(1):148–156. Work, T. H., Trapido, H., Murthy, D. P., Rao, R. L., Bhatt, P. N., and Kulkarni, K. G. (1957). Kyasanur forest disease. III. A preliminary report on the nature of the infection and clinical manifestations in human beings. Indian J. Med. Sci. 11(8):619–645. Worobey, M., and Holmes, E. C. (2001). Homologous recombination in GB virus C/hepatitis G virus. Mol. Biol. Evol. 18(2):254–261. Worobey, M., Rambaut, A., and Holmes, E. C. (1999). Widespread intra-serotype recombination in natural populations of dengue virus. Proc. Natl. Acad. Sci. USA 96(13):7352–7357. Wu, S. J., Grouard-Vogel, G., Sun, W., Mascola, J. R., Brachtel, E., Putvatana, R., Louder, M. K., Filgueira, L., Marovich, M. A., Wong, H. K., Blauvelt, A., Murphy, G. S., et al. (2000). Human skin Langerhans cells are targets of dengue virus infection. Nat. Med 6(7):816–820. Xu, G., Dong, H., Shi, N., Liu, S., Zhou, A., Cheng, Z., Chen, G., Liu, J., Fang, T., Zhang, H., Gu, C., Tan, X., et al. (2007). An outbreak of dengue virus serotype 1 infection in Cixi, Ningbo, People’s Republic of China, 2004, associated with a traveler from Thailand and high density of Aedes albopictus. Am. J. Trop. Med. Hyg. 76(6):1182–1188. Yang, Z., Nielsen, R., Goldman, N., and Pedersen, A. M. (2000). Codon-substitution models for heterogeneous selection pressure at amino acid sites. Genetics/Society 155(1):431–449. Yeh, W. T., Chen, R. F., Wang, L., Liu, J. W., Shaio, M. F., and Yang, K. D. (2006). Implications of previous subclinical dengue infection but not virus load in dengue hemorrhagic fever. FEMS Immunol. Med. Microbiol. 48:84–90. Yuwono, J., Suharyono, W., Koiman, I., Tsuchiya, Y., and Tagaya, I. (1984). Seroepidemiological survey on dengue and Japanese encephalitis virus infections in Asian monkeys. Southeast Asian J. Trop. Med. Public Health 15(2):194–200. Zanotto, P. M., Kallas, E. G., de Souza, R. F., and Holmes, E. C. (1999). Genealogical evidence for positive selection in the nef gene of HIV-1. Genetics 153(3):1077–1089. Zanotto, P. M., Gould, E. A., Gao, G. F., Harvey, P. H., and Holmes, E. C. (1996). Population dynamics of flaviviruses revealed by molecular phylogenies. Proc. Natl. Acad. Sci. USA 93 (2):548–553. Zhang, C., Mammen, M. P., Jr., Chinnawirotpisan, P., Klungthong, C., Rodpradit, P., Monkongdee, P., Nimmannitya, S., Kalayanarooj, S., and Holmes, E. C. (2005). Clade replacements in dengue virus serotypes 1 and 3 are associated with changing serotype prevalence. J. Virol. 79(24):15123–15130. Zivna, I., Green, S., Vaughn, D. W., Kalayanarooj, S., Stephens, H. A., Chandanayingyong, D., Nisalak, A., Ennis, F. A., and Rothman, A. L. (2002). T cell responses to an HLA-B*07restricted epitope on the dengue NS3 protein correlate with disease severity. J. Immunol. 168(11):5959–5965.
CHAPTER
2 Third-Generation Flavivirus Vaccines Based on Single-Cycle, Encapsidation-Defective Viruses Douglas G. Widman,* Ilya Frolov,* and Peter W. Mason*,†
Contents
I. Introduction A. Epidemiology of flavivirus diseases B. Flavivirus biology II. Currently Licensed Vaccines for Flavivirus Diseases A. LAV for yellow fever B. INV and LAV for Japanese encephalitis C. INV for tick-borne encephalitis III. Promising Vaccine Candidates in Development A. Vero cell-derived INV for JE B. Subunit vaccines C. Virus-vectored vaccines D. DNA vaccines E. Traditional LAVs F. Recombinant LAVs for dengue G. Dengue LAV-based chimeras H. ChimeriVax vaccines IV. Replication-Defective and Single-Cycle Virus Vaccines A. Replication-defective and single-cycle virus vaccines for other viral families B. Flavivirus single-cycle nucleic acid vaccine candidates
78 79 81 83 84 86 87 88 88 89 91 93 96 97 99 101 104 104 106
* Department of Microbiology and Immunology, University of Texas Medical Branch, Galveston, Texas 77555 {
Department of Pathology, University of Texas Medical Branch, Galveston, Texas 77555
Advances in Virus Research, Volume 72 ISSN 0065-3527, DOI: 10.1016/S0065-3527(08)00402-8
#
2008 Elsevier Inc. All rights reserved.
77
78
Douglas G. Widman et al.
C. RepliVAX: A particle-based, single-cycle flavivirus vaccine candidate D. Growth of RepliVAX using a novel twocomponent genome system V. Conclusion and Perspectives Acknowledgments References
Abstract
106 110 112 112 113
Flaviviruses are arthropod-borne pathogens that cause significant disease on all continents of the world except Antarctica. Flavivirus diseases are particularly important in tropical regions where arthropod vectors are abundant. Live-attenuated virus vaccines (LAVs) and inactivated virus vaccines (INVs) exist for some of these diseases. LAVs are economical to produce and potent, but are not suitable for use in the immunocompromised. INVs are safer, but are more expensive to produce and less potent. Despite the success of both classes of these first-generation flavivirus vaccines, problems associated with their use indicate a need for improved products. Furthermore, there are no suitable vaccines available for important emerging flavivirus diseases, notably dengue and West Nile encephalitis (WNE). To address these needs, new products, including LAVs, INVs, viral-vectored, genetically engineered LAVs, naked DNA, and subunit vaccines are in various stages of development. Here we describe the current state of these first- and secondgeneration vaccine candidates, and compare these products to our recently described single-cycle, encapsidation defective flavivirus vaccine: RepliVAX. RepliVAX can be propagated in C-expressing cells (or as a unique two-component virus) using methods similar to those used to produce today’s economical and potent LAVs. However, due to deletion of most of the gene for the C protein, RepliVAX cannot spread between normal cells, and is unable to cause disease in vaccinated animals. Nevertheless, RepliVAX is potent and efficacious in animal models for WNE and Japanese encephalitis, demonstrating its utility as a thirdgeneration flavivirus vaccine that should be potent, economical to produce, and safe in the immunocompromised.
I. INTRODUCTION Members of the Flavivirus genus of the Flaviviridae family place a tremendous burden on global public health. The most important members of this group are yellow fever virus (YFV), dengue virus (DENV), Japanese encephalitis virus (JEV), West Nile virus (WNV), and tick-borne encephalitis virus (TBEV). These produce diseases in humans ranging from mild
Third-Generation Flavivirus Vaccines
79
febrile illness to meningoencephalitis or hemorrhagic fever. While it is estimated that more than half of the world’s population is at risk of developing flavivirus diseases, first-world licensed vaccines exist for just three of these diseases. The YFV live-attenuated viral vaccine (LAV) is considered by many to be the gold standard of vaccines, but its success has not been duplicated with vaccines for other flaviviruses and there has been a recent unexplained increase of observed adverse events associated with its use. Inactivated viral vaccines (INV) have been licensed for use against JE and TBE, although these are not without limitations. An increase in adverse events has curtailed the production and use of this JE vaccine (CDC, 2008a; WHO, 2005), and lengthy vaccination schedules requiring frequent boosters along with financial and logistical issues have restricted the widespread use of the TBE INV in endemic regions outside of Central Europe (Gritsun et al., 2003; Kleiter et al., 2007). While an LAV exists to prevent JE, its use outside a handful of East Asian countries is not licensed and major hurdles exist for licensing in developed markets. The recent introduction and spread of WNV in the western hemisphere has reconfirmed the ability of flaviviruses to become endemic in hitherto naı¨ve regions, and increased the urgency with which new treatment options are being explored. As a result, a number of promising vaccine candidates are currently in development, and many of these have reached clinical trials.
A. Epidemiology of flavivirus diseases Medically important members of the Flavivirus genus are arboviruses that are maintained in nature through cycles involving hematophagous arthropod vectors, such as mosquitoes and ticks, and warm-blooded hosts, such as birds and mammals. As a result of this transmission cycle, the host-ranges of flavivirus vectors play a key role in determining the areas under greatest threat of disease. An estimated 2.5 billion people, mostly in tropical regions, are at risk for dengue and annually 50–100 million individuals develop infection with one of the 4 serotypes of this virus (Halstead, 2007). Infection with DENV can result in asymptomatic infection, mild febrile illness (dengue fever, DF), or far more severe diseases such as dengue shock syndrome (DSS) or dengue hemorrhagic fever (DHF). The determinants of such severe manifestations of DENV infection are not well understood, however, previous infection with a different serotype or waning maternal immunity in newborns are clearly risk factors for severe disease (Halstead, 2003; Rothman, 2003). These data were used to produce the theory that the presence of non- or subneutralizing antibodies from previous infection with a heterologous DENV serotype may play a role in these more severe infections by facilitating infection through Fc-receptor-dependent infection of certain types
80
Douglas G. Widman et al.
of cells, a process that is readily demonstrated in vitro (Halstead, 2003). However, alternative hypotheses exist, including those indicating that an inappropriate cellular immune response to the second serotype of DENV encountered could also play a role in the more severe diseases most often encountered in people undergoing an infection with a second serotype of DENV (Rothman, 2003). Recently dengue outbreaks have been reported in Hawaii (Effler et al., 2005) and along the Texas–Mexico border (CDC, 2007), indicating that previously naı¨ve populations are now at risk for this disease. YFV, another flavivirus capable of causing hemorrhagic disease, results in approximately 1000 reported cases each year; however, due to gaps in surveillance the actual number is likely to be much higher (WHO, 2007). Encephalitic flaviviruses present a significant public health threat to much of the world’s population. Although a licensed vaccine has been available to prevent JE for over 40 years, approximately 20,000 cases are reported annually with 6000 resulting in death (Halstead and Tsai, 2004). Unfortunately, due to gaps in surveillance, the incidence of JE is also likely to be much higher than reported (Oya and Kurane, 2007). JEV infection is often asymptomatic; however, when cases do present they are likely to be in the form of severe encephalitis. Mortality rates can reach 30% among confirmed cases, with up to one-third of survivors suffering from permanent and severe neurological sequelae (Myint et al., 2007). The endemic region of JEV is growing, and the affected area has recently expanded west into parts of Pakistan and India, and south into the northern reaches of Australia (Diagana et al., 2007). Recent WNV activity has also demonstrated that flaviviruses can rapidly spread into new environments. Since its arrival in the western hemisphere in the summer of 1999, WNV has spread throughout North America and parts of South America (Gubler, 2007). The disease presentation resulting from WNV infection is similar to JE, but case fatality rates are generally lower and outcomes are more favorable for WNV infections, although due to its recent introduction to developed nations, the longterm outcomes of infection are still unclear (Sejvar, 2007). Worldwide incidence rates are not available, but in the United States alone there have been over 27,000 cases of WNV disease since 1999, resulting in over 1000 fatalities (CDC, 2008b). TBEV infection can also result in disease manifestations ranging from febrile illness to meningoencephalitis and a poliomyelitis-like syndrome (Gritsun et al., 2003). TBE is estimated to be fatal in about 1–2% of cases, although up to 50% of survivors are left with long-term neurological complications (Kunze et al., 2007). Despite a safe and effective vaccine against TBE, there are still more than 10,000 new cases each year due to low vaccination rates in high-risk areas (presumably due to lengthy vaccination schedules and high cost), the existence of highly virulent
81
Third-Generation Flavivirus Vaccines
strains of TBEV, and increased spread of the ticks that serve as vectors for TBEV (Kunze et al., 2007).
B. Flavivirus biology Members of the genus Flavivirus share a number of common characteristics despite inducing a wide range of diseases. Flaviviruses belong to the family Flaviviridae which also contains the genera Hepacivirus and Pestivirus. The Pestivirus genus includes bovine diarrhea virus and classical swine fever virus which, while having important implications on livestock production, have little impact on human health. The lone member of the Hepacivirus genus is hepatitis C virus, a significant human pathogen that establishes a chronic infection that can lead to hepatocellular carcinoma. Flaviviruses are composed of a nucleocapsid enveloped by a host cellderived lipid bilayer studded with the viral surface proteins. Their genomes are approximately 11 kb in length, and are comprised of directly infectious single-stranded, positive-sense RNA that possesses a 50 cap but is not polyadenylated. The RNA genome encodes three structural and seven non-structural proteins and contains both 50 and 30 untranslated regions (UTRs) that play important roles in replication and translation (Lindenbach et al., 2007). The genome is translated into a single polyprotein that is co- and post-translationally cleaved by host and viral proteases (Fig. 1). The first protein produced is the highly basic capsid (C) protein, which associates with the negatively charged RNA genome to form Structural proteins s
s
Non-structural proteins s
s Protease/ helicase
5⬘UTR C
v
pr M
E
f
NS1
NS2A NS2B
? (v) v
NS3
2K
Polymerase
NS4A
v
v
NS4B
v
3⬘UTR
NS5
v
Cleavage sites: s = signal peptidase v = viral encoded NS2B/NS3 f = furin ? = unknown Infectious virion
Subviral particle (SVP)
NS1
FIGURE 1 Representation of flavivirus genome and key antigenic components. Structural genes (C, prM, E) are clustered at the 50 end of the genome while the non-structural components (NS1–5) are located at the 30 end of the genome. Proteolytic cleavage sites are indicated by arrows color-coded to correspond to the enzyme responsible for cleavage. Graphical representations of key antigenic structures [infectious virion, subviral particle (SVP), and NS1] are shown below the genome. (See Page 1 in Color Section at the back of the book.)
82
Douglas G. Widman et al.
the nucleocapsid. The membrane (M) protein is produced within cells as a precursor (prM), and during virion maturation it is cleaved by furin to produce the mature M protein. The envelope (E) protein is involved in cell binding and fusion. During assembly E associates with prM to form provirions covered with 60 trimers of prM/E heterodimers; however, upon maturation these heterodimers dissociate and E forms 90 homodimers on the surface of the mature virion. Upon infection, these homodimers undergo a pH-dependent conformational change that results in the formation of homotrimers that facilitate fusion of the viral lipid envelope with the endosome membrane, resulting in release of the nucleocapsid into the cytoplasm (Lindenbach et al., 2007). Most importantly, for flavivirus vaccine development, E contains neutralizing epitopes and important determinants for host cell tropism. The seven flavivirus non-structural proteins are involved in genome replication and polyprotein processing, as well as modulating host cell function and immune response (Fig. 1). The NS1 protein is secreted from infected cells and involved in RNA replication, although the exact nature of this role is unclear. NS1 is highly immunogenic and NS1 immunization can confer protection from lethal challenge (Schlesinger et al., 1986). NS2A is a small hydrophobic protein that plays a role in virus assembly and RNA recruitment to the genome replication complexes. Interestingly, NS2A has also been shown to act as an interferon antagonist in some mosquito-borne, but not tick-borne flaviviruses. NS2B is a required cofactor for the protease function of NS3, which is responsible for the cleavage of C and all non-structural proteins except NS1. NS3 also has a domain that acts as an RNA helicase. NS3 is followed in the polyprotein by three small membrane-associated proteins: NS4A, NS4B, and 2k, whose precise functions are unknown. The final protein produced in the flavivirus polypeptide is NS5, a multi-functional protein that contains the core subunit of the viral RNA-dependent RNA polymerase, as well as a methyltransferase that serves in capping of the viral RNA (Lindenbach et al., 2007). Many years ago, a noninfectious, slowly sedimenting, hemagglutinating particle (SHA; referred to in more recent literature as a sub-viral particle, SVP) composed of host cell membranes and prM/M and E (but lacking the viral RNA and the nucleocapsid) was discovered in density gradients used to purify viral particles recovered from flavivirus-infected cells or from brains of infected mice (Brandt et al., 1970; Smith et al., 1970; Stevens and Schlesinger, 1965) (see Fig. 1). More recently, recombinant SVPs were shown to elicit antiviral immune responses and have been the basis for a number of vaccine candidates discussed throughout this review. Additionally, truncated forms of the E protein (trE) have been engineered by removal of the carboxy-terminal transmembrane region of E, facilitating secretion as a soluble product (Allison et al., 1995; Delenda
Third-Generation Flavivirus Vaccines
83
et al., 1994; Jan et al., 1993; Men et al., 1991). These trE proteins have also been the primary antigenic component of a number of flavivirus subunit vaccine candidates, although as described herein, side-by-side studies have suggested that SVPs are more effective than trE in eliciting protective immune responses in vivo.
II. CURRENTLY LICENSED VACCINES FOR FLAVIVIRUS DISEASES The existence of efficacious vaccines for a handful of flavivirus diseases has demonstrated that flavivirus diseases can be prevented by vaccination. Licensed vaccines are commercially available for three flavivirus diseases: YF, JE, and TBE although not all are universally accepted throughout the world. The YF vaccine, based on the attenuated YFV17D strain of YFV, has been credited with preventing outbreaks of urban YF, and until recently, this vaccine was considered to be one of the safest and most efficacious LAVs in use. However, the recent discovery that acute viscerotropic disease (YEL-AVD) can be caused by this vaccine has become a concern. A formalin-inactivated JEV vaccine produced in suckling mouse brain (SMB) has been in use in Japan for over 40 years; however, its safety profile has recently come under scrutiny and this vaccine is no longer being manufactured. A live-attenuated vaccine for JE is currently licensed for use in a few select nations. However, due to its production in a cell line unacceptable for vaccine generation in developed countries, it is not used worldwide. An INV is also available for TBE and has demonstrated an excellent safety and efficacy record in Central and Eastern Europe, but is not widely used. Both the TBE and JE INVs suffer from high production costs and the requirement for multiple immunizations that make their use practical only in affluent regions of the world. Thus, despite the availability of these vaccines, there is a tremendous need for new flavivirus vaccines. The development of vaccines against dengue, arguably the medically most important of all flavivirus diseases, is complicated by the fact that low levels of cross-reactive immunity to individual DENV serotypes have been shown to be a risk factor for severe forms of the disease (see Section I.A.). Thus, there is widespread agreement that dengue vaccines must be tetravalent and must be able to simultaneously induce strong immune responses to all four serotypes of DENV. Furthermore, the recent emergence of WNV in the western hemisphere has highlighted the ability of flaviviruses to gain footholds in new environments and increased the urgency of development of treatments and vaccines against emergent flavivirus diseases.
84
Douglas G. Widman et al.
A. LAV for yellow fever Shortly after YFV was first isolated as the causative agent of YF in 1927, work began on the development of a vaccine to prevent the disease. In 1932, French investigators reported the derivation of the French neurotropic vaccine (FNV) by utilizing Max Theiler’s method to passage the French viscerotropic virus strain of YFV through mouse brain (Sellards and Langret, 1932), creating what would be the first efficacious vaccine against YF. The YFV-17D vaccine strain was concurrently derived in the 1930s by passage of the Asibi strain of YFV through mice and chicken embryonic tissues (Theiler and Smith, 1937a). Similar to FNV, YFV-17D showed acceptable levels of attenuation and potency in laboratory evaluations; however, in the early years of production a lack of standardization led to high levels of inconsistency between vaccine lots. The uncontrolled subpassaging of YFV-17D led to the overattenuation of some strains, while cases of yellow fever vaccine-associated neurotropic disease (YEL-AND) in Brazil were associated with use of other strains (Fox et al., 1942), highlighting the genetic instability of this vaccine strain and the ease with which viruses with unexpected properties could arise in manufacture. These problems prompted the WHO to develop and adopt a seed-lot standardization process for the production of YFV-17D in 1945 (WHO, 1945, 1959). This method was modified slightly many years later to reflect advances in technology (WHO, 1976), and it remains the system that still governs the production of vaccines from all three current YFV vaccine strains still in use (17D-204, 17DD, and 17D-204-WHO). Despite the high level of efficacy associated with use of FNV, it was found to produce unacceptably high rates of encephalitis in vaccinees. As a result of these complications, and the availability of the YFV-17D vaccine, FNV use was curtailed greatly after the 1950s and FNV use was suspended altogether in 1980 (Barrett, 1997b). The attenuating affects of serial passage of the YFV-17D strain were clearly demonstrated by the lack of neurotropism, viscertropism, and lethality upon intracerebral inoculation of rhesus monkeys (Theiler and Smith, 1937a) and subcutaneous vaccination of naı¨ve human subjects (Theiler and Smith, 1937b). Furthermore, the virus has demonstrated a limited level of replication in vaccinated individuals, but significant dissemination throughout the body that leads to a robust immune response (Marchevsky et al., 2003). Despite the use of YFV-17D worldwide for over 70 years, the underlying mechanisms of attenuation have only recently been evaluated, and are still not fully understood. The three different YFV-17D strains share 48 nucleotide differences compared to the parental Asibi strain, of which 22 are non-synonymous (dos Santos et al., 1995). Of particular interest, among these 22 amino acid changes, are four located within a surface-exposed loop in domain III (DIII) of the
Third-Generation Flavivirus Vaccines
85
E protein which is believed to be responsible for the inability of YFV-17D to replicate in mosquitoes and also thought to alter receptor binding and cellular tropism (Galler et al., 1997). Although other amino acid changes have been observed throughout the polyprotein (particularly in the nonstructural genes) and 30 UTR, their exact contribution to the attenuated phenotype of YFV-17D is still unclear and these likely function in a synergistic manner (Monath, 2004). With an estimated 200 million doses administered as of 2003 (Marchevsky et al., 2003), YFV-17D was generally considered to have an outstanding safety profile, although there have been a number of reports of severe adverse events associated with vaccination in recent years. The recent reports of adverse events may be due to better surveillance and understanding of symptoms or a higher prevalence of host factors such as thymus disease and advanced age (Barwick, 2004; Hayes, 2007; Khromava et al., 2005; Martin et al., 2001b) that contribute to the development of vaccine-induced disease. Cases of YEL-AND (in many cases associated with specific strains of YFV-17D; see above) have been exceedingly rare, with only 23 cases being reported worldwide since 1945 (Cetron et al., 2002). More recently YEL-AVD, a new syndrome associated with YFV-17D vaccination, has come to light. First recognized in 2001 (CDC, 2002; Chan et al., 2001; Martin et al., 2001a; Vasconcelos et al., 2001), YEL-AVD is believed to occur in one of every 200,000 vaccinations, with a case fatality rate of 60% (Monath, 2008). Disease presentation is similar to that seen with jungle yellow fever, with a clinical spectrum of disease ranging from moderate disease punctuated by liver dysfunction to severe multiorgan failure (Cetron et al., 2002; Hayes, 2007). As a result of the recognition of YEL-AND and YEL-AVD, in 2002 the Advisory Committee on Immunization Practices issued revised guidelines for the administration of YFV-17D. It recommended that very young (<9 months), elderly (>65 years), pregnant, nursing, and immunocompromised individuals, and those with a known hypersensitivity to chicken egg products not be vaccinated with YFV-17D unless there is an eminent risk of being exposed to YFV (Cetron et al., 2002). Thus, despite the longstanding success of YFV-17D, a large and ever-increasing portion of the population is unable to benefit from vaccination, highlighting the need for alternative vaccines to control YF. Despite the steps taken by the WHO in 1945 to standardize the production of YFV-17D vaccine lots, different strains of the vaccine still demonstrate variable degrees of attenuation and thus manufacture requires constant evaluation to ensure safety. As recently as 1998, the WHO has felt the need to reinforce the required use of non-human primates (NHPs) to assess the attenuation and potency of new master and seed stocks of YFV-17D (WHO, 1998). As is seen routinely with other RNA viruses, heterogeneity exists both among and within the
86
Douglas G. Widman et al.
substrains of YFV-17D. Multiple plaque variants can be isolated from a single lot of YFV-17D (Liprandi, 1981) and variants from vaccine preparations of YFV-17D have demonstrated differing levels of attenuation in mice (Gould et al., 1986; Liprandi, 1981). The existence of these variants highlights a problem shared by all classically derived LAVs; the collection of point mutations that are responsible for the attenuating phenotype of the virus are highly susceptible to reversions or compensating mutations that serve to decrease the attenuation and jeopardize the safety of the product. Thus the development of new vaccines whose attenuating characteristics remain stable throughout manufacture of the product is desirable.
B. INV and LAV for Japanese encephalitis A number of different vaccines are available to prevent JE and these have demonstrated an excellent record of efficacy throughout their history. The vaccine that has been in use the longest is the INV prepared from JEVinfected mouse brains. Two different strains of JEV, Nakayama-NIH or Beijing-1, have been used as the seed virus to produce these vaccines although due to a higher antigen yield in manufacture and a stronger neutralizing antibody response upon immunization, the Beijing-1 strain was preferentially used in the late 1990s and early twenty-first century (WHO, 2006). After amplification in SMBs, the virus undergoes rigorous purification including protamine sulfate precipitation and ultracentrifugation before a final formalin-inactivation step (Barrett, 1997a). This vaccine has been used extensively in East Asia since the 1960s to control JE, and is widely used throughout the world to immunize travelers who visit endemic areas. It has been the subject of numerous field studies which have determined its protective efficacy to be 80–90% in JEVendemic regions (Hoke et al., 1988; Hsu et al., 1971). Despite its widespread use and a strong association of vaccination with decline in disease rates, this vaccine is not without drawbacks. The product requires a threedose vaccination schedule in order to induce protective immunity and this, along with the recommendation of boosters every 2–3 years (CDC, 1993), leads to an expensive and time-consuming regimen that makes its widespread use prohibitive, particularly in developing countries. Furthermore, there have been a number of reports of severe adverse reactions associated with use of this JE INV. These include allergic reactions believed to be caused by bovine and porcine gelatin stabilizers used in vaccine formulation or from residual mouse-derived antigens that have been carried over from the production of the virus (Plesner, 2003; Robinson et al., 1991; Sakaguchi et al., 2001; WHO, 2006). Far more serious complications including severe neurological disorders such as acute disseminated encephalomyelitis, reported to be associated with
Third-Generation Flavivirus Vaccines
87
vaccination with this product, have recently drawn the attention of the global health community (Fukuda et al., 1997; Ohtaki et al., 1992, 1995; Plesner et al., 1996). As a result of concerns over these adverse events, in 2005 the Health, Labor and Welfare Ministry of Japan halted recommendation for vaccination of children with this INV (WHO, 2005) and, in 2006, production for use in other developed countries was halted (CDC, 2008a). In addition to this JE INV, an LAV based on the attenuated SA14-14-2 strain has been used as a vaccine in China since 1989 (Oya and Kurane, 2007). The SA14-14-2 strain was derived by virus passage through weanling mice and primary hamster kidney (PHK) cells followed by cloning in chick embryo cells and subpassaging in mice and hamsters (Halstead and Tsai, 2004). This vaccine is produced in PHK cells, and has an excellent record of efficacy in China, Nepal, and Korea (Bista et al., 2001; Sohn et al., 1999). This JE LAV has recently been introduced in India, but except these four nations its use elsewhere in the world is not approved because of concerns over cell substrate certification and licensing (Chang et al., 2004). At the present time two doses of the vaccine are recommended to develop protective immunity, although there is evidence to suggest that a single dose is adequate to achieve comparable efficacy (Bista et al., 2001). The adoption of a single-dose regimen could be an important step in limiting the cost of this vaccine and increasing its chances for more widespread use, assuming production could be shifted to a vaccine-compatible substrate without the loss of potency. This was attempted in the late 1980s when SA14-14-2 was successfully adapted to primary dog kidney (PDK) cells (Eckels et al., 1988). However, in a Phase 1 trial immunogenicity of this PDK-grown LAV was very low, with only 30–50% of vaccinees seroconverting (Halstead and Tsai, 2004). As a result of this, the PDK strain was deemed over-attenuated and development was discontinued. Thus, despite the availability of existing vaccines for JE, there is still a need for new vaccines that have the ability to gain universal acceptance and use.
C. INV for tick-borne encephalitis Similar to JE, an INV exists for the prevention of TBE. Based on the prototype Neudorfl strain of TBEV, manufacture of this vaccine involves propagation of the virus in chick embryo fibroblast cells followed by formalin-inactivation and centrifugal purification (Barrett et al., 2003). A number of different European manufacturers produce variations of the TBE INV. Although there were safety concerns and adverse events associated with early formulations of this INV, the removal of additives such as gelatin has greatly improved the safety profile of these products (Barrett et al., 2003). While these vaccine have an excellent record of efficacy, including the near eradication of TBEV-associated disease in Austria (Barrett, 2001; Gritsun et al., 2003), the lengthy vaccination
88
Douglas G. Widman et al.
schedule (requiring boosters up to one year after initial vaccination to develop protective immunity) has led to the development of disease in vaccinees who had not yet received all necessary boosters and were therefore incompletely immunized (Plisek et al., 2008). Furthermore, use of this vaccine in Eastern Europe and Asia is limited (likely due to financial and logistical issues), leaving a large population living in an endemic area vulnerable to disease (Gritsun et al., 2003). Thus there remains work to be done in order to produce more useful vaccines to prevent disease associated with tick-borne flaviviruses.
III. PROMISING VACCINE CANDIDATES IN DEVELOPMENT To address the lack of vaccines for flavivirus diseases, research has focused on the development of multiple new products. A new INV for JE prepared from virus propagated in Vero cells has recently completed a modified Phase 3 study and seems poised to replace the widely used mouse brain-derived vaccine that has recently come under scrutiny. ChimeriVax, a vaccine platform based on the chimerization of the YFV17D genome with structural genes of other flaviviruses, has been aggressively evaluated in preclinical and clinical tests. Recently ChimeriVax-JE has successfully completed Phase 3 trials and preliminary reports indicate that ChimeriVax-WN and ChimeriVax-Dengue have performed well in Phase 2 evaluations. Although these candidates have been most extensively characterized, they do not represent the whole of the research being performed on flavivirus vaccines. A number of naturally and genetically engineered live-attenuated candidates, particularly against dengue, have begun clinical evaluation, as has a DNA vaccine against WNV. Other vaccines based on subunit, live-vectored, and replicon-based technologies have performed well in preclinical testing and some appear poised for clinical trials. Based on these encouraging results, it seems likely that the next decade will see the licensure of a number of new vaccines to prevent flavivirus diseases.
A. Vero cell-derived INV for JE Due to an ever-increasing intolerance to vaccine-induced side effects, even in a very small percentage of the vaccinated population, efforts have been underway to develop better vaccines for JE. Concern over reactions to gelatin and neural tissue in the mouse brain-derived INV has led to the suspension of its manufacture (see Section II.B.) and a recommendation by the WHO to develop alternative methods of JE vaccine production (Tsai, 2000). One promising candidate for this type of first-generation INV is being developed by Intercell. This product
Third-Generation Flavivirus Vaccines
89
utilizes the SA14-14-2 strain of JEV used in the Chinese LAV as the source of inactivated antigen; however, the virus has been adapted to grow well in vaccine-certified Vero cells and is administered in a highly purified inactivated form similar to the traditional JE INV (Srivastava et al., 2001). Pre-clinical testing of this Vero cell-derived INV demonstrated a level of efficacy comparable to that of the existing JE vaccine (Srivastava et al., 2001) and Phase 1 trials indicated an acceptable level of safety although only 50% of vacinees developed an immune response to vaccination (Lyons et al., 2007). A Phase 2 study demonstrated a better safety profile, higher geometric mean titers, and longer-term seroconversion rates than those achieved with the currently licensed JE vaccine (Lyons et al., 2007). The clear superiority of the Vero cell-derived product set the stage for non-inferiority Phase 3 trials which were mandated by the existence of the proven efficacious SMB-derived INV (Markoff, 2000). A recent report outlining the results of this large-scale Phase 3 non-inferiority study comparing the safety and potency (as a surrogate for efficacy) of the Intercell vaccine to the current SMB-derived INV was encouraging. This study revealed PRNT50 titers obtained from a two-dose immunization schedule with the Intercell vaccine that were comparable to those obtained from the traditional three-dose schedule with the licensed SMB-INV (Tauber et al., 2007). Moreover the Intercell vaccine demonstrated an outstanding safety profile (Tauber et al., 2007). However, since only a few thousand individuals have thus far received the vaccine, no definitive statements attesting the safety of this product have yet been reported (Fischer et al., 2007).
B. Subunit vaccines The development of subunit vaccines containing purified antigenic components to prevent flavivirus diseases has been investigated extensively. Early studies using Kunjin virus demonstrated that vaccination with purified SHA or fragments of E purified from virus-infected cells was capable of inducing a humoral immune response in rabbits, although contamination of the SHA with infectious virus complicated interpretation of these data (Della-Porta and Westaway, 1977). Later studies showed that immunization with the purified secreted NS1 of YFV was capable of protecting NHPs from lethal challenge as well as reducing the magnitude of viremia in these animals (Schlesinger et al., 1986). Since that time, flavivirus vaccine candidates have been created using a number of different expression systems, including viral (Konishi et al., 1992b, 1997b; Qiao et al., 2004), bacterial (Chu et al., 2007; Martina et al., 2008; Simmons et al., 1998, 2006; Wang et al., 2001; Wu et al., 2003), and eukaryotic cells (Konishi and Fujii, 2002; Ledizet et al., 2005; Mota et al., 2005; Mutoh et al., 2004; Robert Putnak et al., 2005; Watts et al., 2007), primarily to drive the
90
Douglas G. Widman et al.
expression of SVPs or portions of the E protein. Some of the first attempts at the development of subunit vaccines involved the use of recombinant viral vectors such as baculoviruses or vaccinia viruses to drive the expression of flavivirus antigens which were then purified and used for immunization. SVPs of JEV produced from recombinant vaccinia virus-infected cells were shown to induce neutralizing antibody responses and long-lasting T cell responses in mice, although use of multiple vaccinations or inclusion of adjuvanting agents was required in order to induce protective immunity (Konishi et al., 1992b, 1997b). Recombinant baculoviruses have been utilized to express the structural proteins of DENV (Delenda et al., 1994; Putnak et al., 1991; Zhang et al., 1988) and WNV (Qiao et al., 2004), although these studies highlighted the difficulty of obtaining purified flavivirus antigens from insect cells and the need for multiple vaccinations (or vaccination with large amounts of antigen) in order to achieve efficacy. Bacterial expression systems were shown to be useful for producing flavivirus antigens over 20 years ago (Mason et al., 1987) and since that time these inexpensive-to-produce proteins have been tested extensively as subunit vaccine candidates. Recombinant E proteins of DENV (Simmons et al., 1998; Srivastava et al., 1995), JEV (Wu et al., 2003), and WNV (Chu et al., 2007; Martina et al., 2008) have been evaluated for efficacy in rodent models of flavivirus disease; however, the inclusion of unlicensed adjuvants, use lengthy vaccination schedules with unreasonably large amounts of antigen and variable levels of efficacy demonstrate the obstacles that will be faced in developing subunit vaccines from bacterially produced flavivirus proteins. Recently a study using DENV-2 was performed in NHPs evaluating various combinations of a recombinant subunit vaccine containing DIII of E, a DNA vaccine expressing prM and E, and an INV, all using a three-dose schedule. In this study, the subunit vaccine induced higher levels of neutralizing antibodies than the DNA vaccine; however, both of these levels were lower than those produced by INV immunization. When challenged, monkeys that had received three doses of the INV were protected from viremia, whereas none of the monkeys receiving the subunit vaccine were protected despite administration of three doses of 1.5 mg of the bacterially produced E antigen fragment (Simmons et al., 2006). Eukaryotic expression systems can also be used to obtain flavivirus antigens that are useful for vaccine production, but the lower yields of antigen obtained from these systems is certainly a drawback to their use in subunit vaccine development. Mammalian cell lines continuously expressing SVPs of JEV (Konishi et al., 2001) and DENV (Konishi and Fujii, 2002) were established from CHO cells and these cell lines were able to express flavivirus proteins for more than 20 cell passages, allowing long-term repeated harvest of antigen, although antigen yield was low,
Third-Generation Flavivirus Vaccines
91
especially in the case of the DENV antigen. A promising step forward was made with the establishment of a rabbit kidney cell line capable of producing large amounts of JEV SVPs (Kojima et al., 2003) and immunization of mice with equal amounts of this antigen or SMB-INV yielded equivalent efficacy in mice (Mutoh et al., 2004). One of the most promising flavivirus subunit vaccine candidates is a WNV product containing purified trE and NS1 proteins that is being developed by Hawaii Biotech. This product is based on combining a drosophila cell-expressed trE with a drosophila cell-expressed NS1 protein and an experimental adjuvant (Lieberman et al., 2007). The high levels of proteins produced by these cells (as high as 25 mg per liter of culture) indicate that this product can be economically manufactured (Lieberman et al., 2007) and evaluation of this vaccine candidate in a hamster model for WNE has shown protection from lethal WNV challenge for up to one year after immunization (Watts et al., 2007). Subunit vaccine candidates such as this one could prove useful in populations such as the elderly and immunocompromised for which application of LAVs is contraindicated.
C. Virus-vectored vaccines A number of different viral vectors have been explored for flavivirus vaccine development. While these products have generally demonstrated high levels of efficacy and safety, prior immunity to the viral vector that prevents immunization to the flavivirus antigens has prevented these vaccine platforms from becoming widely accepted. The choice of viral vector is an important consideration when designing these vaccines, as they need to demonstrate tolerance for the insertion of foreign genes, and the vectors themselves are the primary determinants of infectivity, attenuation, and modulation of the host immune response. Based on these considerations, poxviruses were among the first vectors used to develop flavivirus vaccine candidates. It was first demonstrated with DENV-4 that a single immunization of mice with vaccinia virus recombinants expressing various forms of flavivirus structural and non-structural proteins was capable of eliciting a protective immune response (Bray et al., 1989) and these findings were later duplicated in rabbits using vaccinia recombinants expressing prM and E of JEV (Yasuda et al., 1990). Vaccinia vectors that expressed prM, E, and NS1 of JEV were shown to produce secreted M- and E-containing SVPs, and were efficacious at one dose and uniformly protective at two doses (Mason et al., 1991). Based on these studies, a similar strategy was employed to produce a highly attenuated NYVAC strain of vaccinia virus expressing JEV glycoproteins (referred to below as NYVAC-JEV) with similarly safety and efficacy properties in mice (Konishi et al., 1992a). Because of concerns over vaccinia-induced disease and pre-existing vaccinia immunity, a canarypox-vectored vaccine
92
Douglas G. Widman et al.
for JEV (ALVAC-JEV) was developed simultaneously with NYVAC-JEV using the same strategy and this product demonstrated similar safety and efficacy as its vaccinia counterpart in preclinical evaluation (Konishi et al., 1994, 1997a, 1998a). In rhesus macaques, both NYVAC-JEV and ALVACJEV were shown to be safe and immunogenic. However, it was shown that only NYVAC-JEV was efficacious in preventing encephalitis (Raengsakulrach et al., 1999). Unfortunately development of these vaccine candidates was halted after a Phase 1 trial revealed that ALVAC-JEV was not immunogenic enough to warrant further study and NYVAC-JEV was unable to elicit immune responses in individuals with prior immunity to vaccinia (Kanesa et al., 2000). Another line of research that is currently being pursued involves the use of replication-defective adenovirus (Ad) vectors to drive the expression of flavivirus antigens. Adenovirus vectors were first applied to flavivirus vaccine development in the early 1990s when they were used as expression vectors to produce NS1 of TBEV (Jacobs et al., 1992). Vaccination with this vector was capable of eliciting a protective immune response in 50–75% of challenged mice (Jacobs et al., 1992, 1994), a finding that was not surprising in light of work done with JEV-vaccinia virus recombinants demonstrating that expression of prM and E elicited higher levels of protective immunity than expression of NS1 (Konishi et al., 1991). Due to the inherent obstacles in the development of a safe and effective dengue vaccine, most of the recent work involving adenovirus vectors has been focused on dengue and involves two different strategies. The first utilized the Ad vector to express trE of DENV-2. This vaccine candidate was capable of inducing neutralizing antibodies in mice using a two- or three-dose schedule (Jaiswal et al., 2003); however, later studies utilizing only DIII of E (in hopes of shortening the length of the foreign gene insert in the Ad vector to permit development of a single tetravalent dengue vaccine) demonstrated lower PRNT50 titers despite administration of a DNA plasmid boost (Khanam et al., 2006). Development has moved forward and recently it was reported that a bivalent DENV-2/4 Ad vector encoding DIII of E was capable of eliciting slightly higher virus-specific neutralizing antibody titers than the candidate expressing DIII of only DENV-2, although these studies also employed a DNA boost (Khanam et al., 2007). Another strategy for developing Ad-vectored dengue vaccine candidates more consistent with previous studies has been their use in the expression of SVPs of DENV. With this approach, the prM, E, and NS1 genes of DENV-1 and -2, incorporated into an Ad vector, were shown to elicit robust virus-specific neutralizing antibody titers. However, vaccination also resulted in the production of non-neutralizing cross-reactive antibodies against all four DENV serotypes (Raja et al., 2007). A similar vaccine composed of two Ad vectors that collectively express prM and E of all four DENV serotypes elicited neutralizing antibody responses
Third-Generation Flavivirus Vaccines
93
against all serotypes, but the production of non-neutralizing crossreactive antibodies was again noted (Holman et al., 2007). This vector has also been used to express WNV antigens with promising results (Schepp-Berglind et al., 2007). Despite these advances, the wide prevalence of adenovirus immunity in man draws into question the utility of such vaccine candidates in light of the ‘‘vector immunity’’ problem observed in the above-cited vaccinia-based vaccine trial. A number of new recombinant viral vectors have recently been evaluated for their capacity to elicit protective immune responses against flavivirus diseases. One group has recently evaluated a lentiviral vector expressing trE of WNV and demonstrated complete protection from lethal WNV challenge after only a single vaccination with as few as 50 particles (Iglesias et al., 2006). Unfortunately, the use of viral vectors that incorporate into the host cell genome will face an uphill battle that will require a great deal of refinement before reaching clinical trials. A recombinant virus vector based on the measles virus (MV) vaccine Schwarz strain was developed to express trE of WNV and preclinical evaluation in immunocompromised mice demonstrated detectable PRNT90 titers after a single immunization and robust titers after two doses (Despres et al., 2005). Furthermore, all mice receiving two doses survived lethal WNV challenge. Previous work from this group using a recombinant MV-vectored HIV vaccine in NHPs indicated that pre-existing immunity to measles did not impair the immune response to HIV (Lorin et al., 2004). The encouraging results of these studies warrant further evaluation of this MV-based vaccine vector, to demonstrate that vector immunity does not prevent immunization in the context of an MV-vectored flavivirus vaccine.
D. DNA vaccines Research has recently focused on the development of DNA-based vaccine candidates for flavivirus diseases, due to both the inherent safety of such products and the fact that DNA vaccines circumvent vector immunity. Progress has been slowed by low levels of immunogenicity that have required lengthy immunization schedules using extremely large amounts of DNA. However, the recent advancement of a WNV DNA vaccine to Phase 1 clinical trials (Martin et al., 2007) has ushered in a new era in this field of DNA vaccinology. The first flavivirus DNA vaccine to elicit protective immunity, albeit partial, was a naked DNA plasmid encoding the prM and E genes of St. Louis encephalitis virus, although vaccineinduced neutralizing antibodies were undetectable (Phillpotts et al., 1996). Immunization of mice with plasmids encoding prM and trE of DENV2 was shown to elicit PRNT50 in all animals that had received three 200-mg doses. However, despite this seroconversion, vaccinated mice were not
94
Douglas G. Widman et al.
protected from lethal challenge (Kochel et al., 1997). A DENV-2 DNA vaccine encoding prM and the entire E gene was able to elicit detectable PRNT90 titers after only two 100-mg doses (Konishi et al., 2000), indicating the expression of full-length E (likely resulting in SVP formation) was a more effective way to elicit an immune response. This supposition was fully supported by a study with TBEV which carefully compared DNA vaccines expressing various lengths of E (with or without prM) and clearly demonstrated that candidates that produced SVPs were superior in potency and efficacy to those producing trE (Aberle et al., 1999). This, along with previous in vitro analyses (Allison et al., 1995), demonstrated the requirement of prM and a full-length E protein for the secretion of SVPs and the usefulness of such prM/E cassettes in producing DNA vaccines. Work on DNA vaccines to prevent JE has helped to better characterize the immunological responses required for effective vaccination. Two immunizations, 80 mg each, of DNA encoding prM and E of a Taiwanese strain of JEV was found to confer 70% protection in mice, while inclusion of the NS1 gene in this construct protected 90% of mice from JE (Lin et al., 1998). Another group was able to demonstrate a single 100-mg dose of a plasmid encoding the prM and E genes of the Nakayama strain of JEV was able to induce protective immunity in 80% of challenged mice, although neutralizing antibodies were undetectable without administration of a second dose (Konishi et al., 1998b). A single dose of this vaccine was also shown to induce virus-specific memory cytotoxic T lymphocytes (CTLs) and B cells up to 6 months after immunization (Konishi et al., 1998b). Single administration of a third DNA vaccine candidate utilizing the prM-E region of the SA14 strain of JEV elicited a neutralizing antibody profile similar to that of a two-dose regimen of the currently licensed JE INV and this response was found to be more durable in the DNA vaccine group (Chang et al., 2000). It should be noted, however, that current guidelines for the use of the JE INV recommend a three-dose schedule in humans (CDC, 1993). A potential step forward in addressing the limited immunogenicity of DNA vaccines came when it was shown that simultaneous administration of plasmid-encoded prM and E of JEV along with purified SVPs induced a higher neutralizing antibody response than administration of the individual components (Konishi et al., 2003). A similar strategy utilizing WNV prM-E expressing DNA co-administered with a formalin-inactivated WNV vaccine was undertaken and again a synergistic effect on neutralizing antibody titers was observed (Ishikawa et al., 2007). These results lend the possibility of using increasingly smaller doses of DNA and/or protein in future vaccine formulations, making such vaccines more cost effective. It is thought that DNA vaccines hold the potential to induce protective immunity to all four DENV serotypes simultaneously without
Third-Generation Flavivirus Vaccines
95
immunological interference that, in the case of LAVs, could produce immune responses that are skewed to a subset of serotypes. Unfortunately, lack of an adequate animal model for dengue has made efficacy studies difficult to perform. Much of the current data on these vaccine candidates is serological and difficult to correlate with protection from disease. A tetravalent vaccine preparation containing plasmid DNAs encoding DIII of E of each of the four DENV serotypes was only capable of eliciting a 1:10 neutralizing antibody titer in mice vaccinated three times with 400 mg per dose (Mota et al., 2005). A study using plasmids encoding prM and E of the four DENV serotypes (in order to drive SVP expression) produced more encouraging results, eliciting neutralizing antibodies against all four serotypes with no detectable decrease in titer for up to 30 weeks after vaccination indicating that interference among the four components was not a problem (Konishi et al., 2006). Another study was unable to protect NHPs from DENV-2 viremia using a DNA vaccine, while a dengue INV was able to protect against the same challenge (Simmons et al., 2006). Yet another approach to the development of a tetravalent dengue DNA vaccine has been the use of gene shuffling to create a synthetic E protein derived from fragments of E from all four DENV serotypes. Preliminary reports from this approach indicate that candidate synthetic E proteins can be created and expressed as soluble protein or SVPs and appear to be capable of eliciting immunological responses against all four serotypes (Apt et al., 2006). While administration of a number of these shuffled constructs to rhesus macaques was shown to elicit neutralizing antibodies to all four DENV serotypes, these immune responses decreased significantly by 32 weeks post-vaccination (Raviprakash et al., 2006). Although only DENV-1 and DENV-2 were used for challenge in this study, no protection from DENV-2 viremia was observed in any of the monkeys. It is also unclear whether using a single antigenic component to induce neutralizing antibodies against multiple DENV serotypes will have the unwanted effect of producing crossreactive antibodies that may participate in antibody-mediated enhancement of disease that could result in the development of DHF or DSS. Furthermore, there is also evidence to suggest that such cross-reactive antibody responses are not beneficial in improving disease outcomes from infection with DENV-1 or -2 (Endy et al., 2004). Taken together, these results indicate that DNA vaccines may not be the best strategy for induction of long-lasting protective immunity against dengue. A DNA vaccine encoding the WNV prM and E proteins has been shown to be efficacious in mouse and horse models of WNV disease (Davis et al., 2001) and in 2005 this product became the first DNA vaccine licensed for use in animals when it was approved for use in horses (USDA, 2005). Following these encouraging developments, a Phase 1 clinical trial was initiated using a plasmid construct analogous to the
96
Douglas G. Widman et al.
licensed equine vaccine. While all subjects receiving a two- or three-dose schedule of the vaccine (administered at 4 mg per dose) demonstrated seroconversion, an individual receiving a single vaccination did not (Martin et al., 2007). The two subjects who were given two doses had barely detectible PRNT50 titers of 1:2 and 1:4 respectively, and most of the remaining 12 volunteers who received the full three-dose schedule displayed a decrease in antibody titer at 32 weeks post-vaccination (Martin et al., 2007). While this is an important step forward in flavivirus DNA vaccine development, these results highlight the need for long-term studies that will examine long-lasting immunity and for further research into optimized formulations that will increase immunogenicity in order to decrease the amount of material required for eliciting protective immunity.
E. Traditional LAVs The development of safe and effective LAVs for YF prompted several attempts to produce other traditional LAV candidates, including the SA14-14-2 LAV for JE. Other early attempts to produce LAVs for flavivirus diseases included unsuccessful attempts to produce a dengue LAV performed by Albert Sabin and his military colleagues in the 1940s (Sabin, 1952). Later, a series of four vaccine candidates for dengue were developed in the 1970s and 1980s using classical passage in vitro, although the behavior of these viruses once again highlighted the challenges facing development of these types of empirically derived products (for discussion on YFV, see Section II.A.). LAVs for all four DENV serotypes were developed and clearly shown to exhibit well-accepted markers of attenuation such as small plaque phenotypes, temperature sensitivity, decreased mouse neurovirulence, and diminished viremia in monkeys as compared to their parental viruses (Bancroft et al., 1984; Eckels et al., 1984; Innis et al., 1988; McKee et al., 1987). When DENV-2 and -4 candidates chosen for their attenuated phenotypes were evaluated in Phase 1 trials, they were found to be inadequately immunogenic in terms of seroconversion and duration of immunity, yet despite this, those individuals who did seroconvert experienced a high incidence of dengue-like symptoms (Bancroft et al., 1984; Eckels et al., 1984). Furthermore, the DENV-4 virus recovered from vaccinees demonstrated a large plaque phenotype indicating the genotypic/phenotypic instability of this product. DENV-1 and-3 candidates, despite their attenuated phenotype in laboratory studies, were found to be underattenuated in humans, as they both caused apparent DF in vaccinated volunteers (with viremia from the DENV-1 LAV detectible up to 19 days post-vaccination) (Innis et al., 1988; McKee et al., 1987). While the DENV-1 virus recovered from vaccinees demonstrated an altered phenotype from the vaccine strain (a sign of genetic instability),
Third-Generation Flavivirus Vaccines
97
the DENV-3 product remained stable indicating that the phenotypic markers used to select this latter LAV candidate were inadequate. Despite the fact that these results clearly indicate that markers such as small-plaque and temperature-sensitive phenotypes are not suitable for predicting the attenuation of a vaccine product in humans, these markers have continued to be used as benchmarks in the development of new LAVs. In the early 1980s this field was advanced when, under contract from the U.S. Army, a team led by S. Halstead at the University of Hawaii discovered that DENV could be serially propagated in PDK cells and in the process the virus developed an attenuated phenotype (Halstead et al., 1984a,b,c,d). This strategy was transferred to Mahidol University in Thailand which continued to develop candidate LAVs of three DENV serotypes (DEN-1 PDK13, DEN-2 PDK53, and DEN-4 PDK48) in this manner [summarized in (Bhamarapravati and Yoksan, 1998)]. Because it was discovered that DENV-3 did not replicate in PDK cells it was attenuated by passage in primary green monkey kidney (PGMK) cells and fetal rhesus lung (FRhL) cells and referred to as DEN-3 PGMK 30/FRhL3. Monovalent vaccines consisting of these strains demonstrated a tolerable safety profile in Phase 1 trials, although the tetravalent formulation was unable to consistently produce neutralizing antibodies against all four serotypes of DENV (Kanesa et al., 2001; Sabchareon et al., 2002, 2004). At the same time, researchers at The Walter Reed Army Institute of Research developed a set of attenuated DENV for a tetravalent vaccine in a similar manner (Kanesa-Thasan et al., 2003) that demonstrated a similar safety profile. However, this vaccine failed to induce neutralizing antibody responses to DENV-4 in most vaccinees (Edelman et al., 2003; Sun et al., 2003, 2006). More recent studies have focused on further refinement of dosage and vaccination schedule in NHP models (Koraka et al., 2007; Sun et al., 2006). At the current time, the status of these most promising classically attenuated flavivirus LAV candidates is unclear and after many years spent on their development it remains uncertain how close they are to becoming a licensed vaccine.
F. Recombinant LAVs for dengue A promising rationally attenuated dengue LAV is based on the purposeful deletion of a region in the 30 UTR found to be important for efficient replication of the viral genome. Early work on a recombinant DENV-4 with this 30nt deletion (referred to as rDEN4D30) demonstrated that while the growth of the virus was severely impaired as a result of the deletion, the virus was still capable of safely eliciting a neutralizing antibody response in NHPs and humans (Durbin et al., 2001; Men et al., 1996). It was hoped that a combination of four serotypes of DENV containing the same attenuating deletion could be used as a safe and
98
Douglas G. Widman et al.
efficacious tetravalent vaccine candidate. Based on this hypothesis, the identical 30nt deletion was incorporated into three other DENV serotypes with varied results (Blaney et al., 2004a,b; Whitehead et al., 2003a). While the recombinant DENV-1 mutant (rDEN1D30) demonstrated an acceptable level of attenuation and efficacy in preclinical evaluation (Whitehead et al., 2003a), the DENV-2 (rDEN2D30) and DENV-3 (rDEN3D30) constructs were under-attenuated and thus excluded from clinical evaluation (Blaney et al., 2004a,b). As a result of these findings, a chimeric construction strategy (discussed in Section III.G.) was undertaken as an alternative approach to develop LAV candidates for serotypes 2 and 3 by replacing the structural genes of the well-attenuated rDEN4D30 with those of DENV-2 and -3 (Blaney et al., 2004a; Whitehead et al., 2003b). Although rDEN4D30 backbones containing the C-prM-E cassettes of DENV-2 were found to be over-attenuated, those containing the C of DENV-4 and the prM and E genes of the two other respective serotypes (referred to as rDEN2/4D30 and rDEN3/4D30) were adequately attenuated and safely elicited neutralizing antibody responses in NHPs (Blaney et al., 2004a; Whitehead et al., 2003b). Furthermore, despite success with the rDEN1D30 virus, a recombinant DENV-1 chimeric virus (rDEN1/4D30) has also been engineered and evaluated in NHPs for possible inclusion in a tetravalent vaccine (Blaney et al., 2007). A tetravalent vaccine candidate containing rDEN1D30, rDEN2/4D30, rDEN3/4D30, and rDEN4D30 was evaluated in the NHP model of disease. Since chimerization per se may attenuate virulence (Bray and Lai, 1991), it was not surprising that antibody responses to the two chimeric viruses were much lower than to the DENV-1 and -4 components, and less than 20% of the rhesus monkeys that received a single vaccination with this tetravalent vaccine seroconverted to DENV-2 and -3. DENV-2 challenge experiments revealed that these singly vaccinated monkeys were not protected from DENV-2 viremia (Blaney et al., 2005). However, a cohort of animals given a second dose of this tetravalent vaccine 4 months after initial vaccination displayed a marked increase in antibody titers against all four serotypes and all of these monkeys were completely protected from DENV-2-induced viremia (Blaney et al., 2005). As a result of these encouraging preclinical findings, the four recombinant dengue vaccine candidates have begun individual evaluation in clinical trials. The results of individual Phase 1 trials for rDEN1D30, rDEN2/4D30, and rDEN4D30 have indicated that administration of a single dose containing 1000 pfu of attenuated virus demonstrates an acceptable safety profile and induces seroconversion (defined by a fourfold rise in serum neutralizing antibody titer over pre-vaccination levels) in 95–100% of volunteers tested (Durbin et al., 2005, 2006a,b). Phase 1 evaluation of the chimeric rDEN3/4D30 is currently ongoing (Blaney et al., 2008), but results have yet to be reported. While these results are
Third-Generation Flavivirus Vaccines
99
encouraging, the potential for immunological interference between the four viruses included in a tetravalent vaccine presents a potential roadblock that has yet to be evaluated for these vaccine candidates in humans. In particular, because three of the four vaccine candidates (rDEN2/4D30, rDEN3/4D30, and rDEN4D30) all share the same DENV-4-derived NS1 gene, the development of immunity to this NS1 could limit the duration of antigen presentation and confound the induction of an immune response to the structural components of the chimeric viruses, providing incomplete immunity.
G. Dengue LAV-based chimeras One of the interesting features of flavivirus biology is the ability to swap structural genes of one virus with those of another within the genus. As indicated above, this has led to the development of a number of chimeric dengue vaccine candidates, in which the structural genes of the virus for which the vaccine is being developed are placed into the backbone of an attenuated flavivirus. While the replicational machinery is comprised of components of the backbone virus, the antigenic structures on the surface of the chimeric virions are those of the virus of interest and participate in the induction of immunity. In most cases, this act of chimerization serves to attenuate the phenotype of the resulting virus and has provided an attractive option for vaccine development. This strategy was first demonstrated by Bray and Lai, when they successfully cloned the C-prM-E regions of DENV-1 or DENV-2 into a DENV-4 backbone and recovered live chimeras that in the case of the DENV-2/4 virus was more attenuated than either parental virus in vitro, however, still retained the neurovirulence of the parental viruses in mice (Bray and Lai, 1991). Based on these findings that chimerization per se attenuates these viruses (at least in vitro), a similar chimeric strategy was utilized to develop an attenuated vaccine candidate by inserting the prM and E genes of TBEV into the DENV-4 backbone. This chimera was found to be viable in vitro and vaccination with it was able to completely protect mice from lethal TBEV challenge. However, not surprisingly, it partially retained the neurovirulent phenotype of TBEV (Pletnev et al., 1992). To address this problem, mutations that ablated the prM furin cleavage site or the glycosylation sites of E or NS1 were identified that reduced (although did not ablate) neurovirulence of the chimera without compromising vaccine efficacy (Pletnev et al., 1993) and as an alternative strategy a new DENV-4 chimera was developed utilizing the prM and E genes of the closely related naturally attenuated Langat virus (LGT) in hopes of generating a vaccine capable of cross-protecting against TBEV (Pletnev and Men, 1998). Although this LGT/DEN4 chimera was found to be safe in preclinical evaluation and induced protective immune responses in NHPs and mice against
100
Douglas G. Widman et al.
homologous (LGT) and heterologous (TBEV) virus (Pletnev and Men, 1998; Pletnev et al., 2000, 2001, 2006), a Phase 1 trial recently conducted revealed that despite an acceptable level of attenuation the vaccine was unable to effectively elicit an immune response against TBEV and will likely need to be redesigned (Wright et al., 2008). Although the LGT/DEN4 chimera was unable to stimulate immunity against a heterologous virus in man, it did demonstrate the feasibility of chimerization as a means of developing candidate LAVs. Based on this, the WNV prM/E cassette was used to create a chimera with wild type (wt) DENV-4 (Pletnev et al., 2002). This WN/DEN4 chimeric virus demonstrated an attenuated phenotype and has been shown to confer complete protection from WNV in mice and NHPs at doses as low as 100,000 ffu (Pletnev et al., 2002, 2003). The simultaneous development of a WN vaccine candidate using the rationally attenuated DENV-4 backbone containing a 30nt deletion in the 30 UTR (reviewed in Section III.F.) that has demonstrated similar efficacy with a higher degree of attenuation (Pletnev et al., 2003, 2006); however, make it unlikely that the chimera based on the wt DENV-4 backbone will be evaluated further. A similar chimerization strategy has also been utilized by a group from the CDC which inserted the C-prM-E region of DENV-1 into the attenuated DEN-2 PDK53 virus backbone (Huang et al., 2000) in hopes of developing a LAV candidate for DENV-1 based on the safe and highly immunogenic DEN-2 PDK53 strain selected at Mahidol University using the PDK passaging system (reviewed in Section III.E.). This chimera retained the attenuated phenotype of the parental PDK53 virus and elicited protective immune responses comparable to those seen from Mahidol University’s classically attenuated DEN-1 PDK13 (Huang et al., 2000). A tetravalent vaccine containing three chimeras expressing prM-E of heterologous DENV serotypes in the context of a DEN-2 PDK53 backbone (DEN-2/1, DEN-2/3, and DEN-2/4) and the parental DEN-2 PDK53 virus have shown acceptable levels of attenuation in vitro and in vivo and were capable of eliciting neutralizing antibody titers against all four DENV serotypes in mice. However, as had been seen in previous studies with LAV tetravalent candidates (see Section III.E.), titers to DENV-4 were much lower than to other serotypes and challenge experiments were not conducted (Huang et al., 2003). Recently, a WN/DEN-2 PDK53 chimera has been shown to be safe and efficacious in a murine model of WNV disease (Huang et al., 2005), indicating that chimeras based on the DEN-2 PDK53 may be useful as vaccine candidates for other flavivirus diseases. Because these vaccine candidates rely on classically attenuated DENV backbones, they are subject to the same production and safety issues as traditional LAVs. Characterization of the both the tetravalent dengue vaccine components (Huang et al., 2003) and the WN/DEN-2 PDK53
Third-Generation Flavivirus Vaccines
101
chimera (Huang et al., 2005) has been performed using classical markers of attenuation (i.e., small plaque phenotype, temperature sensitivity, and decreased mouse neurovirulence) that in the case of DENV LAVs, have clearly been inadequate indicators of attenuation and immunogenicity in humans (see Section III.E.). Furthermore reversion at an attenuation locus involved in reduced mouse neurovirulence and decreased replication in mosquito cells has been observed at rates as high as 92% in components of the tetravalent dengue product (Butrapet et al., 2006). The results of these studies emphasize the need for new well-characterized stable LAV candidates.
H. ChimeriVax vaccines One promising set of vaccine candidates for flavivirus diseases has been built on the ChimeriVax vaccine platform that is based on chimeras produced by insertion of various flavivirus structural genes (specifically prM and E) into the YFV-17D backbone. The long-standing history of safety and efficacy of YFV-17D made it an attractive option for use in chimera construction. To date ChimeriVax vaccines prepared in this manner have been produced for JE, WN, and DEN and candidates for all of these have reached clinical trials. These vaccines have demonstrated repeatedly high levels of safety and potency and appear closer to achieving licensure for human use than any other flavivirus LAV candidates in development.
1. ChimeriVax JE
The demonstration by Chambers et al. that viable chimeric viruses encoding the prM and E genes of JEV in the context of the YFV-17D backbone could be produced (Chambers et al., 1999) led to the development of the ChimeriVax platform of vaccines. Although viruses encoding the prM-E proteins of the virulent Nakayama strain or the LAV SA14-14-2 vaccine strain (reviewed in Section II.B.) were produced, only those containing the SA14-14-2 prM and E genes were considered sufficiently attenuated in terms of neurovirulence, and these chimeras were therefore used for all future studies (Chambers et al., 1999; Monath et al., 1999). The SA14-142 product, renamed ChimeriVax-JE, performed well in preclinical evaluation demonstrating a better safety profile than YFV-17D in NHP studies and providing complete protection from JE with one dose of as little as 100 pfu (Monath et al., 2000). When evaluated in humans, this vaccine demonstrated a tolerability profile similar to YFV-17D vaccination, and was immunogenic with a single dose of as little as 10,000 pfu (Monath et al., 2002). In Phase 2 clinical trials the vaccine demonstrated high seroconversion rates against circulating wt viruses and these same studies demonstrated that previous vaccination with YFV-17D did not interfere
102
Douglas G. Widman et al.
with JEV-specific immunity elicited by ChimeriVax-JE (Monath et al., 2003). Analysis of T cell epitopes of ChimeriVax-JE predicted that it is likely that vaccination would provide cross-reaction against wt virus strains (De Groot et al., 2007) and passive antibody transfer experiments also demonstrated the expected cross-reactivity to multiple JEV strains (Beasley et al., 2004). Results of Phase 3 trials demonstrated an excellent safety profile and a homologous virus seroconversion rate of >99% for ChimeriVax-JE compared to 75% for the existing JE INV (Anonymous, 2007a). Thus, ChimeriVax-JE has consistently demonstrated high levels of safety and potency in numerous human trials, although it has still only been administered to less than 5000 people. Larger studies are still needed to better assess the frequency of adverse events to vaccination and protective efficacy of ChimeriVax-JE.
2. ChimeriVax-dengue Simultaneous with its development of ChimeriVax-JE, Acambis began construction of ChimeriVax products to protect against dengue. Utilizing the same YFV-17D backbone and chimerization strategy as employed for JE, recombinant YFV-17D chimeras were developed for all four DENV serotypes (Guirakhoo et al., 2000, 2001). When administered to NHPs, tetravalent ChimeriVax-Dengue was well-tolerated, induced neutralizing antibody responses against all four DENV serotypes (Guirakhoo et al., 2002, 2004) and most animals displayed protection from viremia when challenged with wt DENV (Guirakhoo et al., 2004). Phase 1 studies performed on ChimeriVax-Dengue-2 demonstrated >90% seroconversion even in the face of pre-existing YF immunity and interestingly those YFimmune subjects developed neutralizing antibody titers to all four DENV serotypes (Guirakhoo et al., 2006). This latter finding, which was even found surprising by the authors of this study, stands in contrast to an earlier non-human primate study that reported that YF immunity reduced potency and efficacy of a similar YFV-17D-based DENV-2 chimera (Galler et al., 2005). Phase 2 trials recently completed on the tetravalent ChimeriVax-Dengue vaccine candidate, now licensed to Sanofi Pasteur, resulted in a 100% seroconversion rate to all four DENV serotypes (Anonymous, 2007b), although details have yet to be published in peer-reviewed literature. Unfortunately problems with immunological interference have been observed with ChimeriVax-Dengue, but initial attempts to balance these responses by altering the makeup of the tetravalent product, have improved antibody responses to the four DENV serotypes (Guirakhoo et al., 2001, 2004). Taken together these results closely mirror those seen for ChimeriVax-JE, indicating this platform of vaccine development could be capable of producing consistently safe and efficacious vaccines against a number of flavivirus diseases.
Third-Generation Flavivirus Vaccines
103
3. ChimeriVax-WN The ChimeriVax vaccine platform has also been used to develop a vaccine candidate for WN known as ChimeriVax-WN. Initial studies were performed using a chimeric YFV-17D virus containing the prM-E of WNV NY99 which displayed a measurable level of neurovirulence in mice, albeit lower than the parental YFV-17D virus (Monath, 2001). To further attenuate this chimeric LAV, three mutations corresponding to a set of those distinguishing the E protein of the SA14-14-2 strain of JEV from to its parental virus were engineered into the WNV E. This mutated ChimeriVax-WN (ChimeriVax-WN02) was selected for all subsequent NHP and human studies. These NHP studies confirmed the safety of ChimeriVax-WN02 by demonstrating that intracranial inoculation of NHPs with this product resulted in significantly less neurovirulence than YFV-17D and that viremia from ChimeriVax-WN02 was within WHO specifications outlined for YFV-17D (Arroyo et al., 2004). ChimeriVax-WN02 also demonstrated efficacy, as all immunized monkeys were completely protected from WNV viremia following intracranial challenge (Arroyo et al., 2004). Based on these findings ChimeriVax-WN02 was evaluated in Phase 1 clinical trials where it stimulated robust neutralizing antibody titers and T cell responses against WNV after a single vaccination (Monath et al., 2006). Initial reports of results from Phase 2 trials have indicated a 97% seroconversion rate from one dose (Anonymous, 2006); however, these data have yet to be peer-reviewed. Thus, similar to its sister vaccines for JE and DEN, ChimeriVax-WN02 has demonstrated an acceptable safety and potency profile through all preclinical and clinical studies, and warrants further investigation as a potential LAV for WNV. Despite the rapid progress of ChimeriVax products through the research and development pipeline, some questions remain about the safety and stability of these vaccine candidates. Mutations have been observed in nearly all of the ChimeriVax products as a result of either in vitro or in vivo passaging and the overall contribution of these mutations to the attenuated phenotypes of the viruses is still largely unknown (Arroyo et al., 2004; Guirakhoo et al., 2004; Monath et al., 1999). The vaccine lot of ChimeriVax-WN02 produced using good manufacturing practices (GMP) was found to contain an equal mixture of large- and small-plaque viruses (Arroyo et al., 2004) and serial passage of the GMP vaccine lots of all four ChimeriVax-Dengue selected mutations in the DENV-3 and -4 chimeras that resulted in a large-plaque phenotype and higher viral yields (Guirakhoo et al., 2004). Mutations selected for in passaging of the ChimeriVax-JE vaccine have also been identified though not wellcharacterized (Monath et al., 1999), underscoring the genetic instability of these products. Although these mutations do not appear to have an effect on mouse neurovirulence, it has been established that, at least in
104
Douglas G. Widman et al.
the case of DEN LAVs, in vitro and murine models are not always a reliable indicator of attenuation in humans (see Section III.E.). Furthermore the overall genetic instability of ChimeriVax products would seem to indicate that the safe use of these vaccines (especially in the immunocompromised) may require additional evaluation and the rigorous implementation of quality control measures.
IV. REPLICATION-DEFECTIVE AND SINGLE-CYCLE VIRUS VACCINES Replication-defective virus vaccines, which utilize mutant virus strains that contain defects that render them incapable of replicating their viral genomes, or single-cycle virus vaccines (also referred to as single-round infectious particles and pseudoinfectious viruses) which utilize mutant virus strains that are unable to assemble and release progeny virus particles, are a new area of vaccine development being explored for a number of diseases. These classes of vaccines (particularly single-cycle viruses) combine the replicative capacity of LAVs with the safety of INV products without the concern for residual virulence and/or reversion to virulence associated with traditional LAVs, or the difficult production methods (requiring large-scale production and removal of contaminants that can produce adverse reactions) and low potency of INVs. Both replicationdefective and single-cycle viruses are propagated in complementing or packaging cell lines designed to express the defective gene(s) of the deliberately mutated virus, allowing for efficient genome replication and packaging, respectively. When these viruses infect normal cells (in a vaccinee for instance) genome replication (replication-defective) or production of infectious progeny (single-cycle) does not occur, but instead the intact functions of the virus serve to drive gene expression. When used as vaccines this gene expression results in the production of viral antigens that can induce protective immune responses without cell-to-cell spread of the virus.
A. Replication-defective and single-cycle virus vaccines for other viral families Replication-defective and single-cycle virus vaccines have been developed from herpes simplex viruses (HSV). Initial studies with these DNA viruses demonstrated that vaccination with replication-defective HSV-1 viruses could safely protect mice from lethal challenge (Morrison and Knipe, 1994; Nguyen et al., 1992). Studies using a single-cycle HSV-1 that was unable to synthesize a surface-exposed glycoprotein necessary for infectivity demonstrated that it was possible to propagate such a virus in
Third-Generation Flavivirus Vaccines
105
packaging cell lines (Forrester et al., 1992) and these defective viruses could be used as vaccines (Farrell et al., 1994). Replication-defective adenoviruses have been explored for their utility as vectors to produce vaccines against diseases caused by several agents (Dudek and Knipe, 2006) including flaviviruses (discussed along with other viral expression systems in Section III.C.). It is likely that in the case of positive-stranded RNA viruses, single-cycle virus vaccines would be advantageous over replication-defective products because multiple copies of the viral genome produced by single-cycle viruses would lead to high-level expression of viral proteins needed to elicit a protective immune response (the small number of genome copies and associated viral proteins produced by cells infected with replicationdefective positive-strand viruses would be unlikely to elicit strong immune responses). Single-cycle viruses composed of packaged replicons encoding the intact nonstructural polyprotein of alphaviruses as well as the antigenic proteins of heterologous pathogens under control of the alphavirus subgenomic promoter have been extensively studied. Current state-of-the-art in packaging of these replicon genomes requires simultaneous electroporation of three different RNA species (encoding the replicon genome encoding the foreign antigen, a helper RNA to supplying the alphavirus capsid protein, and a helper RNA supplying the alphavirus glycoproteins) into the same cells. This strategy has been used to develop vaccine candidates for diseases caused by several viruses including influenza virus (Pushko et al., 1997), Marburg virus (Hevey et al., 1998; Lee et al., 2006), respiratory syncytial virus (Elliott et al., 2007; Mok et al., 2007), and DENV (White et al., 2007). The technology is currently being developed by several companies including Alphavax, which recently reported results from Phase 1 influenza trials (Anonymous, 2007b). Although elegant, the method is not without drawbacks that may make these types of vaccines technically difficult to produce and expensive. These problems include the fact that the vaccines are prepared directly from cultures of electroporated cells. Thus there is no amplification step of the type used to produce traditional LAVs; all vaccine is recovered from cells that have been directly electroporated with three synthetic RNAs that need to be repeatedly synthesized following the very high manufacturing standards needed to produce a product for use in man. Furthermore, the simultaneous replication of three alphavirus RNAs within the same cell has led to concerns about recombination between the genomes that could produce a disease-causing virus capable of developing a spreading infection, requiring additional quality control steps. Finally, repeated vaccinations with packaged alphavirus replicons can lead to the development of adaptive immune response to the alphavirus envelope (Davis et al., 2000). Thus, the problems cited above for vector-immunity to other types of viral-vectored flavivirus vaccines (see Section III.C.) could have a significant impact on the utility of alphavirus replicon particles as vaccines.
106
Douglas G. Widman et al.
B. Flavivirus single-cycle nucleic acid vaccine candidates Because the non-structural components of flaviviruses are required for efficient genome replication and the preservation of prM and E is crucial for production and secretion of highly immunogenic SVPs from infected cells, the capsid gene was targeted for development of single-cycle flavivirus mutants. The utility of this approach was first demonstrated by Kofler et al. using TBEV (Kofler et al., 2002, 2004). These authors reported that deletion of a large internal portion of the C gene completely ablated production of infectious progeny without significantly impacting RNA replication or translation. Cells transfected with in vitro-synthesized RNA with several of these deletions in the C gene (in some cases augmented by specific mutations added to optimize the signal sequence at the start of prM) released large amounts of E protein structurally and antigenically identical to recombinant SVPs. Furthermore, synthetic TBEV RNAs containing these C gene-deletions were found to be highly attenuated in suckling mice (no infectious progeny or disease were detected after intracranial inoculation) yet still capable of eliciting protective immune responses in adult mice that were comparable to those produced by the licensed inactivated TBEV vaccine (Aberle et al., 2005; Kofler et al., 2004). In addition, the RNA vaccine induced strong cellular immune responses not produced by vaccination with the INV (Aberle et al., 2005). Although this RNA-based vaccine candidate was ground-breaking, since it was the first report of a single-cycle flavivirus, the challenges that all RNA-based vaccines face [including stability, production, and limited potency (Cannon and Weissman, 2002)] may prevent single-cycle C gene-deleted RNAs from being developed into a viable vaccine candidates. A similar approach has been taken by creating a C gene-deleted genome of WNV for delivery in a DNA vaccine format (Seregin et al., 2006). When taken into cells, this DNA drives the production of viral RNA that initiates the infectious cycle and leads to genome replication and secretion of SVPs and NS1. This vaccine candidate was capable of eliciting detectible neutralizing antibody titers and protecting mice from lethal WNV challenge, although protection was only examined after two DNA injections (Seregin et al., 2006), highlighting another challenge (limited potency – see Section III.D.) in nucleic acid vaccine development.
C. RepliVAX: A particle-based, single-cycle flavivirus vaccine candidate Since replicons can be packaged into particles, a logical extension of these C gene-deleted nucleic acid vaccines was the trans-complementation of their C gene-deleted genomes using a packaging technology similar to that utilized to create single-cycle flavivirus replicon particles [also
Third-Generation Flavivirus Vaccines
107
known as virus-like particles (VLPs) or pseudoinfectious particles (PIPs)] that have been used to study aspects of flavivirus assembly and/or infection (Bourne et al., 2007; Gehrke et al., 2003; Jones et al., 2005; Khromykh et al., 1998; Scholle et al., 2004; Yoshii et al., 2008). RepliVAX, the name we have given to our version of this type of vaccine, is a singlecycle flavivirus that consists of a replicationally active, C gene-deleted genome that is encapsidated into an infectious particle by packaging cells producing the missing C protein (Fig. 2A). Our initial studies demonstrated that the genomes of YFV or WNV engineered to lack most of the C gene (a portion the C gene was retained to maintain a short RNA structure required for genome replication; see below) could be readily packaged into high-titer stocks of infectious particles in stable cell lines expressing the missing C protein from a non-cytopathic Venezuelan equine encephalitis virus replicon (VEErep) (Mason et al., 2006). The distinguishing feature of this packaging technology is that in the packaging cells, the C-deficient virus develops productive, spreading infection and the titers of infectious particles, containing defective genomes, approach the levels of wt virus grown in similar cell lines. Therefore, unlike other technologies (notably the alphavirus replicon particles mentioned in Section IV. A.), RepliVAX vaccine production does not require repeated RNA transfections, can be easily standardized, and since the single-cycle virus is severely attenuated, the high biocontainment conditions required to propagate virus (such as those needed to produce INVs) are also not needed. RepliVAX particles released from C-expressing cells are infectious, but are capable of performing only a single round of replication in cells that do not express C (Fig. 2B). However, infection of these cells results in the efficient release of SVPs and NS1, products that are known to be potent and efficacious vaccine components when delivered by other methods (see above). RepliVAX has been shown to be highly attenuated in baby mice (at least 1,000,000-fold less virulent than wt viruses, with no death detected in pups given as much as two million infectious units (iu)) (Mason et al., 2006). Thus, as expected, RepliVAX WN was shown to be safe in adult mice and was also capable of eliciting neutralizing antibody responses and protecting these animals from lethal WNV challenge after a single dose (Mason et al., 2006). As shown in Fig. 2A, C gene-deleted RepliVAX genomes can be packaged into infectious particles by transencapsidation in cells lines that continuously express the missing C protein in the context of a noncytopathic VEErep. To enhance the safety of this system over our initial report (Mason et al., 2006), we generated a VEErep with multiple modifications designed to enhance its safe utilization. First, the WNV C gene has been modified to prevent intergenomic recombination by utilizing a form of C that was systematically altered in its first 30 codons to have the
108
Douglas G. Widman et al.
A
RepliVAX production cell RepliVAX genome (C-deleted replicon)
VEEreplicon encoding the C protein (+) RNA
(+) RNA
(−) RNA (−) RNA
Translation of C
Genome replication
Translation of prM/E
Genome replication (+) RNA
(+) RNA
Nucleus Packaging of RepliVAX genome
B
Processing and release of RepliVAX and NS1
RepliVAX-infected cell RepliVAX genome (C-deleted replicon) (+) RNA Transcription
(−) RNA Genome replication (+) RNA Translation
Nucleus
Processing and release of SVPs and NS1
FIGURE 2 (A) RepliVAX production in C-expressing cells. C gene-deleted RepliVAX genomes are introduced by electroporation into cells expressing the missing C gene from a non-cytopathic VEErep. RepliVAX produces all of the components necessary for genome replication and expression of the C gene by the VEErep allows for packaging of RepliVAX genomes into infectious particles that are released [along with NS1 and SVPs (not shown)] via the exocytic pathway. Alternatively, mature RepliVAX infectious particles can be used (in place of electroporated genetic material) to initiate the infection process for large-scale cultivation. (B) RepliVAX infection of normal cells in culture or in a vaccinated individual. Infection of cells with RepliVAX mimics the early events of natural flavivirus infection. The RepliVAX genome is replicated in the cell, and antigenic components [including released products and peptides that can be loaded onto MHC molecules (not shown)] are produced. The lack of C prevents RepliVAX genomes from being packaged into infectious particles, but does not interfere with secretion of other antigenic components. Non-infectious SVPs and NS1 are released from RepliVAXinfected cells, permitting them to elicit protective humoral immunity in vaccinated individuals. (See Page 2 in Color Section at the back of the book.)
Third-Generation Flavivirus Vaccines
109
maximum number of nucleotide differences with the RepliVAX-encoded C fragment (Widman et al., 2008). Second, included in these mutations were changes to the cyclization sequence (CS), an RNA structure that is required for genome replication and which must be complementary to a 30 CS to function (Lindenbach et al., 2007). Thus, a recombination event between the cell-encoded C gene and RepliVAX would yield an RNA species with non-complementary CSs that would be unable to replicate. Third, to further reduce the chances of recombination, the C gene in this VEErep expression construct contained a stop codon at the NS2B/NS3 cleavage site (Widman et al., 2008), even though we had previously shown that slightly higher titers of infectious particles could be produced from C gene-deleted genomes if the packaging construct contained the entire C gene and portions of the prM gene (Mason et al., 2006). The fourth feature to this C gene-expression cassette was the direct fusion of the modified C gene, via a ubiquitin gene linker, to the puromycin acetyl transferase gene (used to maintain the VEEreps in these cells) (Widman et al., 2008). BHK cells were able to carry the resulting modified C expression construct for dozens of passages without loss of packaging ability and studies with vaccine-certified Vero cells expressing this C construct produced high titers of infectious particles for long periods permitting repeated harvests (Widman et al., 2008). Importantly, repeated passaging of RepliVAX WN in cells expressing the modified C construct failed to detect any evidence of recombination, even following 30 sequential passages. Interestingly, these sequential passages selected a better-growing version of RepliVAX WN which was used to re-engineer a secondgeneration product with superior growth properties (Widman et al., 2008), allowing for more efficient production without a loss of attenuation. This highlights a key advantage of RepliVAX over traditional and chimeric LAVs; since the mechanism of attenuation (namely the C gene-deletion preventing packaging) is completely defined, mutations that alter growth properties can be used to improve production without the possibility of having an effect on virulence. This property stands in marked contrast to all other LAVs, where the precise mechanism of attenuation is unclear. This places a tremendous burden on quality control in manufacture of other types of LAVs, because changes to in vitro properties often encountered during vaccine production (e.g., changes in temperature sensitivity or plaque size; see Section III.E.) could result in changes in the ability of these LAVs to produce disease, especially in immunocompromised hosts. Using mouse and hamster models for WNE we were able to demonstrate that a single inoculation of RepliVAX WN at the lowest dose tested (40,000 iu in mice and 200,000 iu in hamsters) provided complete protection from lethal challenge (Widman et al., 2008). Further studies have demonstrated complete protection of hamsters using a 40,000 iu dose
110
Douglas G. Widman et al.
(Widman et al., unpublished work), and protection of both mice and hamsters at this dose have also been achieved using either intraperitoneal or subcutaneous (s.c.) inoculations (Widman et al., 2008, unpublished work). As with the TBE RNA vaccines (see Section IV.B.), RepliVAX WN elicits T cell responses, and RepliVAX-induced memory responses are similar to those produced by wt WNV infection (Brien et al., unpublished work). Evaluation of RepliVAX WN in the rhesus macaque model of WN disease is currently ongoing, but initial results have shown that a single s.c. inoculation of RepliVAX WN is capable of inducing neutralizing antibody titers in these animals (Widman et al., unpublished work). The utility of RepliVAX as a vaccine platform has also been demonstrated by the development of a chimeric RepliVAX to prevent JE (Ishikawa et al., 2008). This RepliVAX JE genome encodes the WNV replicative machinery (NS1–5), JEV prM/E, and the small fragment of WNV C (containing the CS) found in our original RepliVAX WN. RepliVAX JE grew to high titers in precisely the same WNV C-expressing cell line used to produce RepliVAX WN (see above and (Widman et al., 2008), although an adaptive mutation (similar to those found in RepliVAX WN) needed to be added to RepliVAX JE to produce a product which displayed enhanced growth in vitro (Ishikawa et al., 2008). Interestingly, in the case of RepliVAX JE, we were able to document that this adaptive mutation increased both infectious titer (in C-expressing cells) and SVP yield (in normal cells) in vitro (Ishikawa et al., 2008). Evaluation in rodent models has demonstrated complete safety and efficacy of RepliVAX JE in homologous (JE) and heterologous (WNE) models (Ishikawa et al., 2008).
D. Growth of RepliVAX using a novel two-component genome system Recently we have shown that RepliVAX can be efficiently packaged in normal cells if these cells are co-transfected (or co-infected) with a second flavivirus genome expressing C, but not the structural glycoproteins (Shustov et al., 2007). In this novel system, the flavivirus genetic information essential for viral packaging has been split into two separate replication-competent genomes (Fig. 3). This system provides the advantage over the ‘‘packaging’’ cell system (see Section IV.C.) in that any type of cell (including currently available vaccine production-grade cells) could be used for large-scale RepliVAX production. Initially, both C- and prM/ E-encoding in vitro-synthesized genomes are delivered into the cells via co-transfection and presence of both replication-competent RNAs in the same cell leads to synthesis of the complete set of viral structural proteins and release of both defective genomes, packaged individually into infectious viral particles (Fig. 3A). These particles can be further propagated
Third-Generation Flavivirus Vaccines
111
RepliVAX two-component production cell
A
Helper genome (prM/E-deleted replicon)
RepliVAX genome (C-deleted replicon)
(+) RNA
(+) RNA
Translation of C
(−) RNA Genome replication
(−) RNA Translation of prM/E
Genome replication (+) RNA
(+) RNA
Nucleus Packaging of RepliVAX genome Processing and release of two-component RepliVAX and NS1
RepliVAX two-component infection
B RepliVAX genome (C-deleted replicon)
Helper genome (prM/E-deleted replicon) (+) RNA
(−) RNA
Genome replication
(+) RNA
(+) RNA
(−) RNA
Genome replication
(+) RNA Translation
Translation
Production of intracellular C Nucleus
Nucleus Processing and release of SVPs and NS1
Release of NS1
FIGURE 3 (A) Two-component RepliVAX production by high multiplicity infection in normal cells in culture. C gene-deleted RepliVAX genomes and prM/E-deleted helper genomes are simultaneously introduced by electroporation into normal cells. Both genomes produce all of the components necessary for genome replication. Expression of prM/E by the RepliVAX genome in concert with expression of the C gene by the helper genome allows for packaging of both genomes into individual infectious particles that are released [along with NS1 and SVPs (not shown)] via the exocytic pathway. Alternatively, these mature RepliVAX and helper infectious particles can also be used (in place of electroporated genetic material) to initiate this process when infection is carried out at MOIs greater than 1. (B) Two-component RepliVAX infection of normal cells in culture or in vaccinated individuals. Infection of cells with either RepliVAX or helper particles mimics the early events of natural flavivirus infection. This type of infection occurs in vivo and in vitro at MOIs less than 1. Left: The RepliVAX genome is replicated in the cell, producing SVPs and NS1. The lack of C prevents RepliVAX genomes from being packaged into infectious particles, but does not interfere with secretion of SVPs or NS1. Right: The helper genome is replicated in the cell, and NS1 is produced. The lack of prM and E prevents helper genomes from being packaged into infectious particles. (See Page 3 in Color Section at the back of the book.)
by co-infecting naı¨ve cells at a multiplicity of infection (MOI) above 1, but are incapable of developing productive infection either at lower MOIs in vitro or in vivo, because under these conditions C- and prM/E-encoding genomes do not replicate in the same cell (Fig. 3B). Replication of both genomes in the same cell raises a concern about the possibility of intermolecular recombination that might lead to the
112
Douglas G. Widman et al.
formation of infectious, replication-competent virus. Therefore, additional safety features have been employed such as use of sequences in C of the helper genome that utilize different codons to minimize similarity, thus reducing the potential for homologous recombination. Moreover, in our initial constructs the functional C coding region was separated from the fragment of C encoding the CS, reducing the chance that homologous recombination events could produce a replication-competent RNA (Shustov, Mason, and Frolov, 2007). Thus, with these constructs, the generation of viable pathogenic viruses from these RNAs would require several rounds of non-homologous recombination events, and to date such recombination has never been detected despite extensive passaging of these two-component viruses (Shustov and Frolov, unpublished work).
V. CONCLUSION AND PERSPECTIVES The RepliVAX platform is still under investigation, but studies cited above demonstrate its potential for large-scale production at low cost in low biocontainment conditions, and its potency/efficacy in animals. The inability of RepliVAX to produce spreading infection in vitro, and its resulting safety in vivo (even in immuno-incompetent baby mice) suggests that unlike other LAVs, RepliVAX will be suitable for use in highly immunocompromised individuals. Moreover, unlike virus-vectored vaccines (see Section III.C.) and alphavirus replicon particles (see Section IV. A.), immunity to the proteins encapsidating RepliVAX is unlikely to interfere with its utility, since it is not vectored by another virus which is subject to neutralization by heterologous antibodies. The surprising potency of RepliVAX suggests that this unique LAV is targeted to sites where effective innate immune stimulation and efficient antigen presentation are quickly engaged, permitting the production of an efficacious immune response. Learning more about these aspects of RepliVAX mechanisms of action and confirming the utility of this vaccine candidate in NHPs appear to be the next steps in the development of RepliVAX into a vaccine that can be tested in humans. In summary, RepliVAX represents a promising technology for development of new flavivirus vaccines that combine the potency, efficacy and economy of LAVs and safety of INVs and subunit vaccines.
ACKNOWLEDGMENTS We thank James Brien and Janko Nikolich-Zugich (University of Arizona, Tucson, AZ) and Ricardo Carrion (SWFBR, San Antonio, TX) for sharing unpublished data. We thank Franz Heinz, University of Vienna for providing helpful information on TBE vaccination. PWM and IF are supported by grants from the NIH, and DGW is supported by a James W. McLaughlin fellowship.
Third-Generation Flavivirus Vaccines
113
REFERENCES Aberle, J. H., Aberle, S. W., Allison, S. L., Stiasny, K., Ecker, M., Mandl, C. W., Berger, R., and Heinz, F. X. (1999). A DNA immunization model study with constructs expressing the tick-borne encephalitis virus envelope protein E in different physical forms. J. Immunol. 163(12):6756–6761. Aberle, J. H., Aberle, S. W., Kofler, R. M., and Mandl, C. W. (2005). Humoral and cellular immune response to RNA immunization with flavivirus replicons derived from tickborne encephalitis virus. J. Virol. 79(24):15107–15113. Allison, S. L., Stadler, K., Mandl, C. W., Kunz, C., and Heinz, F. X. (1995). Synthesis and secretion of recombinant tick-borne encephalitis virus protein E in soluble and particulate form. J. Virol. 69(9):5816–5820. Anonymous. (2006). Acambis announces encouraging data from Phase 2 trial of West Nile virus vaccine Acambis. Cambridge, Massachusetts, Cambridge, UK. Anonymous. (2007a). Acambis’ JE vaccine meets and exceeds primary endpoint in pivotal Phase 3 efficacy trial Acambis. Cambridge, Massachusetts, Cambridge, UK. Anonymous. (2007b). Sanofi Pasteur presents positive results of tetravalent dengue candidate vaccine. Sanofi Pasteur, Lyon, France. Apt, D., Raviprakash, K., Brinkman, A., Semyonov, A., Yang, S., Skinner, C., Diehl, L., Lyons, R., Porter, K., and Punnonen, J. (2006). Tetravalent neutralizing antibody response against four dengue serotypes by a single chimeric dengue envelope antigen. Vaccine 24 (3):335–344. Arroyo, J., Miller, C., Catalan, J., Myers, G. A., Ratterree, M. S., Trent, D. W., and Monath, T. P. (2004). ChimeriVax-West Nile virus live-attenuated vaccine: preclinical evaluation of safety, immunogenicity, and efficacy. J. Virol. 78(22):12497–12507. Bancroft, W. H., Scott, R. M., Eckels, K. H., Hoke, C. H., Jr., Simms, T. E., Jesrani, K. D., Summers, P. L., Dubois, D. R., Tsoulos, D., and Russell, P. K. (1984). Dengue virus type 2 vaccine: Reactogenicity and immunogenicity in soldiers. J. Infect. Dis. 149(6):1005–1010. Barrett, A. D. (1997a). Japanese encephalitis and dengue vaccines. Biologicals 25(1):27–34. Barrett, A. D. (1997b). Yellow fever vaccines. Biologicals 25(1):17–25. Barrett, A. D. (2001). Current status of flavivirus vaccines. Ann. N. Y. Acad. Sci. 951:262–271. Barrett, P. N., Schober-Bendixen, S., and Ehrlich, H. J. (2003). History of TBE vaccines. Vaccine 21(Suppl 1):S41–S49. Barwick, R. (2004). History of thymoma and yellow fever vaccination. Lancet 364(9438):936. Beasley, D. W., Li, L., Suderman, M. T., Guirakhoo, F., Trent, D. W., Monath, T. P., Shope, R. E., and Barrett, A. D. (2004). Protection against Japanese encephalitis virus strains representing four genotypes by passive transfer of sera raised against ChimeriVax-JE experimental vaccine. Vaccine 22(27–28):3722–3726. Bhamarapravati, N., and Yoksan, S. (1998). Live attenuated tetravalent dengue vaccine. In ‘‘Dengue and Dengue Hemorrhagic Fever’’ (D. Gubler and G. Kuno, eds.), pp. 367–377. CAB International, Wallingford. Bista, M. B., Banerjee, M. K., Shin, S. H., Tandan, J. B., Kim, M. H., Sohn, Y. M., Ohrr, H. C., Tang, J. L., and Halstead, S. B. (2001). Efficacy of single-dose SA 14-14-2 vaccine against Japanese encephalitis: a case control study. Lancet 358(9284):791–795. Blaney, J. E., Jr., Hanson, C. T., Firestone, C. Y., Hanley, K. A., Murphy, B. R., and Whitehead, S. S. (2004a). Genetically modified, live attenuated dengue virus type 3 vaccine candidates. Am. J. Trop. Med. Hyg. 71(6):811–821. Blaney, J. E., Jr., Hanson, C. T., Hanley, K. A., Murphy, B. R., and Whitehead, S. S. (2004b). Vaccine candidates derived from a novel infectious cDNA clone of an American genotype dengue virus type 2. BMC Infect. Dis. 4:39. Blaney, J. E., Jr., Matro, J. M., Murphy, B. R., and Whitehead, S. S. (2005). Recombinant, liveattenuated tetravalent dengue virus vaccine formulations induce a balanced, broad, and
114
Douglas G. Widman et al.
protective neutralizing antibody response against each of the four serotypes in rhesus monkeys. J. Virol. 79(9):5516–5528. Blaney, J. E., Jr., Sathe, N. S., Hanson, C. T., Firestone, C. Y., Murphy, B. R., and Whitehead, S. S. (2007). Vaccine candidates for dengue virus type 1 (DEN1) generated by replacement of the structural genes of rDEN4 and rDEN4Delta30 with those of DEN1. Virol. J. 4:23. Blaney, J. E., Jr., Sathe, N. S., Goddard, L., Hanson, C. T., Romero, T. A., Hanley, K. A., Murphy, B. R., and Whitehead, S. S. (2008). Dengue virus type 3 vaccine candidates generated by introduction of deletions in the 30 untranslated region (30 -UTR) or by exchange of the DENV-3 30 -UTR with that of DENV-4. Vaccine 26(6):817–828. Bourne, N., Scholle, F., Silva, M. C., Rossi, S. L., Dewsbury, N., Judy, B., De Aguiar, J. B., Leon, M. A., Estes, D. M., Fayzulin, R., and Mason, P. W. (2007). Early production of type i interferon during West Nile virus infection: Role for lymphoid tissues in IRF3-independent interferon production. J. Virol. 81(17):9100–9108. Brandt, W. E., Cardiff, R. D., and Russell, P. K. (1970). Dengue virions and antigens in brain and serum of infected mice. J. Virol. 6(4):500–506. Bray, M., and Lai, C. J. (1991). Construction of intertypic chimeric dengue viruses by substitution of structural protein genes. Proc. Natl. Acad. Sci. USA 88(22):10342–10346. Bray, M., Zhao, B. T., Markoff, L., Eckels, K. H., Chanock, R. M., and Lai, C. J. (1989). Mice immunized with recombinant vaccinia virus expressing dengue 4 virus structural proteins with or without nonstructural protein NS1 are protected against fatal dengue virus encephalitis. J. Virol. 63(6):2853–2856. Butrapet, S., Kinney, R. M., and Huang, C. Y. (2006). Determining genetic stabilities of chimeric dengue vaccine candidates based on dengue 2 PDK-53 virus by sequencing and quantitative TaqMAMA. J. Virol. Methods 131(1):1–9. Cannon, G., and Weissman, D. (2002). RNA based vaccines. DNA Cell Biol. 21(12):953–961. CDC. (1993). Inactivated Japanese encephalitis virus vaccine. Recommendations of the Advisory Committee on Immunization Practices (ACIP). MMWR Recomm. Rep. 42(RR-1):1–15. CDC. (2002). Adverse events associated with 17D-derived yellow fever vaccination--United States, 2001–2002. MMWR Morb. Mortal. Wkly. Rep. 51(44):989–993. CDC. (2007). Dengue hemorrhagic fever–U.S.-Mexico border, 2005. MMWR Morb. Mortal. Wkly. Rep. 56(31):785–789. CDC. (2008a). Health Information for International Travel 2008. Atlanta: US Department of Health and Human Services, Public Health Service. CDC. (2008b). West Nile Virus - Statistics, Surveillance, and Control - Case Counts.. Cetron, M. S., Marfin, A. A., Julian, K. G., Gubler, D. J., Sharp, D. J., Barwick, R. S., Weld, L. H., Chen, R., Clover, R. D., Deseda-Tous, J., Marchessault, V., Offit, P. A., et al. (2002). Yellow fever vaccine. Recommendations of the Advisory Committee on Immunization Practices (ACIP), 2002. MMWR Recomm. Rep. 51(RR-17):1–11; quiz CE1–4. Chambers, T. J., Nestorowicz, A., Mason, P. W., and Rice, C. M. (1999). Yellow fever/ Japanese encephalitis chimeric viruses: Construction and biological properties. J. Virol. 73(4):3095–3101. Chan, R. C., Penney, D. J., Little, D., Carter, I. W., Roberts, J. A., and Rawlinson, W. D. (2001). Hepatitis and death following vaccination with 17D-204 yellow fever vaccine. Lancet North Am. Ed. 358(9276):121–122. Chang, G. J., Kuno, G., Purdy, D. E., and Davis, B. S. (2004). Recent advancement in flavivirus vaccine development. Expert Rev. Vaccines 3(2):199–220. Chang, G. J. J., Hunt, A. R., and Davis, B. (2000). A single intramuscular injection of recombinant plasmid DNA induces protective immunity and prevents Japanese encephalitis in mice. J. Virol. 74(9):4244–4252.
Third-Generation Flavivirus Vaccines
115
Chu, J. H., Chiang, C. C., and Ng, M. L. (2007). Immunization of flavivirus West Nile recombinant envelope domain III protein induced specific immune response and protection against West Nile virus infection. J. Immunol. 178(5):2699–2705. Davis, N. L., Caley, I. J., Brown, K. W., Betts, M. R., Irlbeck, D. M., McGrath, K. M., Connell, M. J., Montefiori, D. C., Frelinger, J. A., Swanstrom, R., Johnson, P. R., and Johnston, R. E. (2000). Vaccination of macaques against pathogenic simian immunodeficiency virus with Venezuelan equine encephalitis virus replicon particles. J. Virol. 74 (1):371–378. Davis, B. S., Chang, G. J. J., Cropp, B., Roehrig, J. T., Martin, D. A., Mitchell, C. J., Bowen, R., and Bunning, M. L. (2001). West Nile virus recombinant DNA vaccine protects mouse and horse from virus challenge and expresses in vitro a noninfectious recombinant antigen that can be used in enzyme-linked immunosorbent assays. J. Virol. 75 (9):4040–4047. De Groot, A. S., Martin, W., Moise, L., Guirakhoo, F., and Monath, T. (2007). Analysis of ChimeriVax Japanese Encephalitis Virus envelope for T-cell epitopes and comparison to circulating strain sequences. Vaccine 25(47):8077–8084. Delenda, C., Frenkiel, M. P., and Deubel, V. (1994). Protective efficacy in mice of a secreted form of recombinant dengue-2 virus envelope protein produced in baculovirus infected insect cells. Arch. Virol. 139(1–2):197–207. Della-Porta, A. J., and Westaway, E. G. (1977). Immune response in rabbits to virion and nonvirion antigens of the Flavivirus kunjin. Infect. Immun. 15(3):874–882. Despres, P., Combredet, C., Frenkiel, M. P., Lorin, C., Brahic, M., and Tangy, F. (2005). Live measles vaccine expressing the secreted form of the West Nile virus envelope glycoprotein protects against West Nile virus encephalitis. J. Infect. Dis. 191(2):207–214. Diagana, M., Preux, P. M., and Dumas, M. (2007). Japanese encephalitis revisited. J. Neurol. Sci. 262(1–2):165–170. dos Santos, C. N., Post, P. R., Carvalho, R., Ferreira, C. M., II, Rice, C. M., and Galler, R. (1995). Complete nucleotide sequence of yellow fever virus vaccine strains 17DD and 17D-213. Virus Res. 35(1):35–41. Dudek, T., and Knipe, D. M. (2006). Replication-defective viruses as vaccines and vaccine vectors. Virology 344(1):230–239. Durbin, A. P., Karron, R. A., Sun, W., Vaughn, D. W., Reynolds, M. J., Perreault, J. R., Thumar, B., Men, R., Lai, C. J., Elkins, W. R., Chanock, R. M., Murphy, B. R., et al. (2001). Attenuation and immunogenicity in humans of a live dengue virus type-4 vaccine candidate with a 30 nucleotide deletion in its 30 -untranslated region. Am. J. Trop. Med. Hyg. 65(5):405–413. Durbin, A. P., Whitehead, S. S., McArthur, J., Perreault, J. R., Blaney, J. E., Jr., Thumar, B., Murphy, B. R., and Karron, R. A. (2005). rDEN4delta30, a live attenuated dengue virus type 4 vaccine candidate, is safe, immunogenic, and highly infectious in healthy adult volunteers. J. Infect. Dis. 191(5):710–718. Durbin, A. P., McArthur, J., Marron, J. A., Blaney, J. E., Jr., Thumar, B., Wanionek, K., Murphy, B. R., and Whitehead, S. S. (2006a). The live attenuated dengue serotype 1 vaccine rDEN1Delta30 is safe and highly immunogenic in healthy adult volunteers. Hum. Vaccin. 2(4):167–173. Durbin, A. P., McArthur, J. H., Marron, J. A., Blaney, J. E., Thumar, B., Wanionek, K., Murphy, B. R., and Whitehead, S. S. (2006b). rDEN2/4Delta30(ME), a live attenuated chimeric dengue serotype 2 vaccine is safe and highly immunogenic in healthy denguenaive adults. Hum. Vaccin. 2(6):255–260. Eckels, K. H., Scott, R. M., Bancroft, W. H., Brown, J., Dubois, D. R., Summers, P. L., Russell, P. K., and Halstead, S. B. (1984). Selection of attenuated dengue 4 viruses by serial passage in primary kidney cells. V. Human response to immunization with a candidate vaccine prepared in fetal rhesus lung cells. Am. J. Trop. Med. Hyg. 33(4):684–689.
116
Douglas G. Widman et al.
Eckels, K. H., Yu, Y. X., Dubois, D. R., Marchette, N. J., Trent, D. W., and Johnson, A. J. (1988). Japanese encephalitis virus live-attenuated vaccine, Chinese strain SA14142; adaptation to primary canine kidney cell cultures and preparation of a vaccine for human use. Vaccine 6(6):513–518. Edelman, R., Wasserman, S. S., Bodison, S. A., Putnak, R. J., Eckels, K. H., Tang, D., KanesaThasan, N., Vaughn, D. W., Innis, B. L., and Sun, W. (2003). Phase I trial of 16 formulations of a tetravalent live-attenuated dengue vaccine. Am. J. Trop. Med. Hyg. 69(6 Suppl.):48–60. Effler, P. V., Pang, L., Kitsutani, P., Vorndam, V., Nakata, M., Ayers, T., Elm, J., Tom, T., Reiter, P., Rigau-Perez, J. G., Hayes, J. M., Mills, K., et al. (2005). Dengue fever, Hawaii, 2001–2002. Emerg. Infect. Dis. 11(5):742–749. Elliott, M. B., Chen, T., Terio, N. B., Chong, S. Y., Abdullah, R., Luckay, A., Egan, M. A., Boutilier, L. A., Melville, K., Lerch, R. A., Long, D., Eldridge, J. H., et al. (2007). Alphavirus replicon particles encoding the fusion or attachment glycoproteins of respiratory syncytial virus elicit protective immune responses in BALB/c mice and functional serum antibodies in rhesus macaques. Vaccine 25(41):7132–7144. Endy, T. P., Nisalak, A., Chunsuttitwat, S., Vaughn, D. W., Green, S., Ennis, F. A., Rothman, A. L., and Libraty, D. H. (2004). Relationship of preexisting dengue virus (DV) neutralizing antibody levels to viremia and severity of disease in a prospective cohort study of DV infection in Thailand. J. Infect. Dis. 189(6):990–1000. Farrell, H. E., McLean, C. S., Harley, C., Efstathiou, S., Inglis, S., and Minson, A. C. (1994). Vaccine potential of a herpes simplex virus type 1 mutant with an essential glycoprotein deleted. J. Virol. 68(2):927–932. Fischer, M., Casey, C., and Chen, R. T. (2007). Promise of new Japanese encephalitis vaccines. Lancet 370(9602):1806–1808. Forrester, A., Farrell, H., Wilkinson, G., Kaye, J., Davis-Poynter, N., and Minson, T. (1992). Construction and properties of a mutant of herpes simplex virus type 1 with glycoprotein H coding sequences deleted. J. Virol. 66(1):341–348. Fox, J. P., Lennette, E. H., Manso, C., and Aguiar, J. R. S. (1942). Encephalitis in man following vaccination with 17 D yellow fever virus. Am. J. Hyg. 36(2):117–142. Fukuda, H., Umehara, F., Kawahigashi, N., Suehara, M., and Osame, M. (1997). Acute disseminated myelitis after Japanese B encephalitis vaccination. J. Neurol. Sci. 148 (1):113–115. Galler, R., Freire, M. S., Jabor, A. V., and Mann, G. F. (1997). The yellow fever 17D vaccine virus: Molecular basis of viral attenuation and its use as an expression vector. Braz. J. Med. Biol. Res. 30(2):157–168. Galler, R., Marchevsky, R. S., Caride, E., Almeida, L. F., Yamamura, A. M., Jabor, A. V., Motta, M. C., Bonaldo, M. C., Coutinho, E. S., and Freire, M. S. (2005). Attenuation and immunogenicity of recombinant yellow fever 17D-dengue type 2 virus for rhesus monkeys. Braz. J. Med. Biol. Res. 38(12):1835–1846. Gehrke, R., Ecker, M., Aberle, S. W., Allison, S. L., Heinz, F. X., and Mandl, C. W. (2003). Incorporation of tick-borne encephalitis virus replicons into virus-like particles by a packaging cell line. J. Virol. 77(16):8924–8933. Gould, E. A., Buckley, A., Barrett, A. D., and Cammack, N. (1986). Neutralizing (54K) and non-neutralizing (54K and 48K) monoclonal antibodies against structural and nonstructural yellow fever virus proteins confer immunity in mice. J. Gen. Virol. 67(Pt 3): 591–595. Gritsun, T. S., Lashkevich, V. A., and Gould, E. A. (2003). Tick-borne encephalitis. Antiviral Res. 57(1–2):129–146. Gubler, D. J. (2007). The continuing spread of West Nile virus in the western hemisphere. Clin. Infect. Dis. 45(8):1039–1046. Guirakhoo, F., Weltzin, R., Chambers, T. J., Zhang, Z. X., Soike, K., Ratterree, M., Arroyo, J., Georgakopoulos, K., Catalan, J., and Monath, T. P. (2000). Recombinant chimeric yellow
Third-Generation Flavivirus Vaccines
117
fever-dengue type 2 virus is immunogenic and protective in nonhuman primates. J. Virol. 74(12):5477–5485. Guirakhoo, F., Arroyo, J., Pugachev, K. V., Miller, C., Zhang, Z. X., Weltzin, R., Georgakopoulos, K., Catalan, J., Ocran, S., Soike, K., Ratterree, M., and Monath, T. P. (2001). Construction, safety, and immunogenicity in nonhuman primates of a chimeric yellow fever-dengue virus tetravalent vaccine. J. Virol. 75(16):7290–7304. Guirakhoo, F., Pugachev, K., Arroyo, J., Miller, C., Zhang, Z. X., Weltzin, R., Georgakopoulos, K., Catalan, J., Ocran, S., Draper, K., and Monath, T. P. (2002). Viremia and immunogenicity in nonhuman primates of a tetravalent yellow fever-dengue chimeric vaccine: Genetic reconstructions, dose adjustment, and antibody responses against wild-type dengue virus isolates. Virology 298(1):146–159. Guirakhoo, F., Pugachev, K., Zhang, Z., Myers, G., Levenbook, I., Draper, K., Lang, J., Ocran, S., Mitchell, F., Parsons, M., Brown, N., Brandler, S., et al. (2004). Safety and efficacy of chimeric yellow Fever-dengue virus tetravalent vaccine formulations in nonhuman primates. J. Virol. 78(9):4761–4775. Guirakhoo, F., Kitchener, S., Morrison, D., Forrat, R., McCarthy, K., Nichols, R., Yoksan, S., Duan, X., Ermak, T. H., Kanesa-Thasan, N., Bedford, P., Lang, J., et al. (2006). Live attenuated chimeric yellow fever dengue type 2 (ChimeriVax-DEN2) vaccine: Phase I clinical trial for safety and immunogenicity: Effect of yellow fever pre-immunity in induction of cross neutralizing antibody responses to all 4 dengue serotypes. Hum. Vaccin. 2(2):60–67. Halstead, S. B. (2003). Neutralization and antibody-dependent enhancement of dengue viruses. Adv. Virus Res. 60:421–467. Halstead, S. B. (2007). Dengue. Lancet 370(9599):1644–1652. Halstead, S. B., and Tsai, T. F. (2004). Japanese encephalitis vaccine. In (‘‘Vaccine’’) (S. Plotkin and W. A. Orenstein, eds.), 5th Ed., pp. 919–957. W.B. Saunders Company, Philadelphia, PA. Halstead, S. B., Diwan, A. R., Marchette, N. J., Palumbo, N. E., and Srisukonth, L. (1984a). Selection of attenuated dengue 4 viruses by serial passage in primary kidney cells. I. Attributes of uncloned virus at different passage levels. Am. J. Trop. Med. Hyg. 33 (4):654–665. Halstead, S. B., Eckels, K. H., Putvatana, R., Larsen, L. K., and Marchette, N. J. (1984b). Selection of attenuated dengue 4 viruses by serial passage in primary kidney cells. IV. Characterization of a vaccine candidate in fetal rhesus lung cells. Am. J. Trop. Med. Hyg. 33 (4):679–683. Halstead, S. B., Marchette, N. J., Diwan, A. R., Palumbo, N. E., and Putvatana, R. (1984c). Selection of attenuated dengue 4 viruses by serial passage in primary kidney cells. II. Attributes of virus cloned at different dog kidney passage levels. Am. J. Trop. Med. Hyg. 33 (4):666–671. Halstead, S. B., Marchette, N. J., Diwan, A. R., Palumbo, N. E., Putvatana, R., and Larsen, L. K. (1984d). Selection of attenuated dengue 4 viruses by serial passage in primary kidney cells. III. Reversion to virulence by passage of cloned virus in fetal rhesus lung cells. Am. J. Trop. Med. Hyg. 33(4):672–678. Hayes, E. B. (2007). Acute viscerotropic disease following vaccination against yellow fever. Trans. R. Soc. Trop. Med. Hyg. 101(10):967–971. Hevey, M., Negley, D., Pushko, P., Smith, J., and Schmaljohn, A. (1998). Marburg virus vaccines based upon alphavirus replicons protect guinea pigs and nonhuman primates. Virology 251(1):28–37. Hoke, C. H., Nisalak, A., Sangawhipa, N., Jatanasen, S., Laorakapongse, T., Innis, B. L., Kotchasenee, S., Gingrich, J. B., Latendresse, J., Fukai, K., and Burke, D. S. (1988). Protection against Japanese encephalitis by inactivated vaccines. N. Engl. J. Med. 319 (10):608–614.
118
Douglas G. Widman et al.
Holman, D. H., Wang, D., Raviprakash, K., Raja, N. U., Luo, M., Zhang, J., Porter, K. R., and Dong, J. Y. (2007). Two complex, adenovirus-based vaccines that together induce immune responses to all four dengue virus serotypes. Clin. Vaccine Immunol. 14 (2):182–189. Hsu, T. C., Chow, L. P., Wei, H. Y., Chen, C. L., and Hsu, S. T. (1971). A controlled field trial for an evaluation of effectiveness of mouse-brain Japanese encephalitis vaccine. Taiwan Yi Xue Hui Za Zhi 70(2):55–62. Huang, C. Y., Butrapet, S., Tsuchiya, K. R., Bhamarapravati, N., Gubler, D. J., and Kinney, R. M. (2003). Dengue 2 PDK-53 virus as a chimeric carrier for tetravalent dengue vaccine development. J. Virol. 77(21):11436–11447. Huang, C. Y., Silengo, S. J., Whiteman, M. C., and Kinney, R. M. (2005). Chimeric dengue 2 PDK-53/West Nile NY99 viruses retain the phenotypic attenuation markers of the candidate PDK-53 vaccine virus and protect mice against lethal challenge with West Nile virus. J. Virol. 79(12):7300–7310. Huang, C. Y. H., Butrapet, S., Pierro, D. J., Chang, G. J. J., Hunt, A. R., Bhamarapravati, N., Gubler, D. J., and Kinney, R. M. (2000). Chimeric dengue type 2 (vaccine strain PDK-53)/ dengue type 1 virus as a potential candidate dengue type 1 virus vaccine. J. Virol. 74 (7):3020–3028. Iglesias, M. C., Frenkiel, M. P., Mollier, K., Souque, P., Despres, P., and Charneau, P. (2006). A single immunization with a minute dose of a lentiviral vector-based vaccine is highly effective at eliciting protective humoral immunity against West Nile virus. J. Gene Med. 8 (3):265–274. Innis, B. L., Eckels, K. H., Kraiselburd, E., Dubois, D. R., Meadors, G. F., Gubler, D. J., Burke, D. S., and Bancroft, W. H. (1988). Virulence of a live dengue virus vaccine candidate: A possible new marker of dengue virus attenuation. J. Infect. Dis. 158 (4):876–880. Ishikawa, T., Takasaki, T., Kurane, I., Nukuzuma, S., Kondo, T., and Konishi, E. (2007). Coimmunization with West Nile DNA and inactivated vaccines provides synergistic increases in their immunogenicities in mice. Microbes Infect. 9(9):1089–1095. Ishikawa, T., Widman, D. G., Bourne, N., Konishi, E., and Mason, P. W. (2008). Construction and evaluation of a chimeric pseudovirus vaccine to prevent Japanese encephalitis. Vaccine (In Press). Jacobs, S. C., Stephenson, J. R., and Wilkinson, G. W. (1992). High-level expression of the tickborne encephalitis virus NS1 protein by using an adenovirus-based vector: Protection elicited in a murine model. J. Virol. 66(4):2086–2095. Jacobs, S. C., Stephenson, J. R., and Wilkinson, G. W. (1994). Protection elicited by a replication-defective adenovirus vector expressing the tick-borne encephalitis virus non-structural glycoprotein NS1. J. Gen. Virol. 75(Pt 9):2399–2402. Jaiswal, S., Khanna, N., and Swaminathan, S. (2003). Replication-defective adenoviral vaccine vector for the induction of immune responses to dengue virus type 2. J. Virol. 77 (23):12907–12913. Jan, L. R., Yang, C. S., Henchal, L. S., Sumiyoshi, H., Summers, P. L., Dubois, D. R., and Lai, C. J. (1993). Increased immunogenicity and protective efficacy in outbred and inbred mice by strategic carboxyl-terminal truncation of Japanese encephalitis virus envelope glycoprotein. Am. J. Trop. Med. Hyg. 48(3):412–423. Jones, C. T., Patkar, C. G., and Kuhn, R. J. (2005). Construction and applications of yellow fever virus replicons. Virology 331(2):247–259. Kanesa, T. N., Smucny, J. J., Hoke, C. H., Jr., Marks, D. H., Konishi, E., Kurane, I., Tang, D. B., Vaughn, D. W., Mason, P. W., and Shope, R. E. (2000). Safety and immunogenicity of NYVAC-JEV and ALVAC-JEV attenuated recombinant Japanese encephalitis virus: Poxvirus vaccines in vaccinia-nonimmune and vaccinia-immune humans. Vaccine 19 (4–5):483–491.
Third-Generation Flavivirus Vaccines
119
Kanesa, T. N., Sun, W., Kim, A. G., Van, A. S., Putnak, J. R., King, A., Raengsakulsrach, B., Christ, S. H., Gilson, K., Zahradnik, J. M., Vaughn, D. W., Innis, B. L., et al. (2001). Safety and immunogenicity of attenuated dengue virus vaccines (Aventis Pasteur) in human volunteers. Vaccine 19(23–24):3179–3188. Kanesa-Thasan, N., Edelman, R., Tacket, C. O., Wasserman, S. S., Vaughn, D. W., Coster, T. S., Kim-Ahn, G. J., Dubois, D. R., Putnak, J. R., King, A., Summers, P. L., Innis, B. L., et al. (2003). Phase 1 studies of Walter Reed Army Institute of Research candidate attenuated dengue vaccines: Selection of safe and immunogenic monovalent vaccines. Am. J. Trop. Med. Hyg. 69(6 Suppl.):17–23. Khanam, S., Khanna, N., and Swaminathan, S. (2006). Induction of neutralizing antibodies and T cell responses by dengue virus type 2 envelope domain III encoded by plasmid and adenoviral vectors. Vaccine 24(42–43):6513–6525. Khanam, S., Rajendra, P., Khanna, N., and Swaminathan, S. (2007). An adenovirus prime/ plasmid boost strategy for induction of equipotent immune responses to two dengue virus serotypes. BMC Biotechnol. 7:10. Khromava, A. Y., Eidex, R. B., Weld, L. H., Kohl, K. S., Bradshaw, R. D., Chen, R. T., and Cetron, M. S. (2005). Yellow fever vaccine: An updated assessment of advanced age as a risk factor for serious adverse events. Vaccine 23(25):3256–3263. Khromykh, A. A., Varnavski, A. N., and Westaway, E. G. (1998). Encapsidation of the flavivirus kunjin replicon RNA by using a complementation system providing Kunjin virus structural proteins in trans. J. Virol. 72(7):5967–5977. Kleiter, I., Jilg, W., Bogdahn, U., and Steinbrecher, A. (2007). Delayed humoral immunity in a patient with severe tick-borne encephalitis after complete active vaccination. Infection 35 (1):26–29. Kochel, T., Wu, S. J., Raviprakash, K., Hobart, P., Hoffman, S., Porter, K., and Hayes, C. (1997). Inoculation of plasmids expressing the dengue-2 envelope gene elicit neutralizing antibodies in mice. Vaccine 15(5):547–552. Kofler, R. M., Heinz, F. X., and Mandl, C. W. (2002). Capsid protein C of tick-borne encephalitis virus tolerates large internal deletions and is a favorable target for attenuation of virulence. J Virol 76(7):3534–3543. Kofler, R. M., Aberle, J. H., Aberle, S. W., Allison, S. L., Heinz, F. X., and Mandl, C. W. (2004). Mimicking live flavivirus immunization with a noninfectious RNA vaccine. Proc. Natl. Acad. Sci. USA 101(7):1951–1956. Kojima, A., Yasuda, A., Asanuma, H., Ishikawa, T., Takamizawa, A., Yasui, K., and Kurata, T. (2003). Stable high-producer cell clone expressing virus-like particles of the Japanese encephalitis virus e protein for a second-generation subunit vaccine. J. Virol. 77 (16):8745–8755. Konishi, E., and Fujii, A. (2002). Dengue type 2 virus subviral extracellular particles produced by a stably transfected mammalian cell line and their evaluation for a subunit vaccine. Vaccine 20(7–8):1058–1067. Konishi, E., Pincus, S., Fonseca, B. A., Shope, R. E., Paoletti, E., and Mason, P. W. (1991). Comparison of protective immunity elicited by recombinant vaccinia viruses that synthesize E or NS1 of Japanese encephalitis virus. Virology 185(1):401–410. Konishi, E., Pincus, S., Paoletti, E., Laegreid, W. W., Shope, R. E., and Mason, P. W. (1992a). A highly attenuated host range-restricted vaccinia virus strain, NYVAC, encoding the prM, E, and NS1 genes of Japanese encephalitis virus prevents JEV viremia in swine. Virology 190(1):454–458. Konishi, E., Pincus, S., Paoletti, E., Shope, R. E., Burrage, T., and Mason, P. W. (1992b). Mice immunized with a subviral particle containing the Japanese encephalitis virus prM/M and E proteins are protected from lethal JEV infection. Virology 188(2):714–720.
120
Douglas G. Widman et al.
Konishi, E., Pincus, S., Paoletti, E., Shope, R. E., and Wason, P. W. (1994). Avipox virusvectored Japanese encephalitis virus vaccines: Use as vaccine candidates in combination with purified subunit immunogens. Vaccine 12(7):633–638. Konishi, E., Kurane, I., Mason, P. W., Shope, R. E., and Ennis, F. A. (1997a). Poxvirus-based Japanese encephalitis vaccine candidates induce JE virus- specific CD8 þ cytotoxic T lymphocytes in mice. Virology 227(2):353–360. Konishi, E., Win, K. S., Kurane, I., Mason, P. W., Shope, R. E., and Ennis, F. A. (1997b). Particulate vaccine candidate for Japanese encephalitis induces long-lasting virusspecific memory T lymphocytes in mice. Vaccine 15(3):281–286. Konishi, E., Kurane, I., Mason, P. W., Shope, R. E., Kanesa-Thasan, N., Smucny, J. J., Hoke, C. H., Jr., and Ennis, F. A. (1998a). Induction of Japanese encephalitis virus-specific cytotoxic T lymphocytes in humans by poxvirus-based JE vaccine candidates. Vaccine 16 (8):842–849. Konishi, E., Yamaoka, M., Khin Sane, W., Kurane, I., and Mason, P. W. (1998b). Induction of protective immunity against Japanese encephalitis in mice by immunization with a plasmid encoding Japanese encephalitis virus premembrane and envelope genes. J. Virol. 72(6):4925–4930. Konishi, E., Yamaoka, M., Kurane, I., and Mason, P. W. (2000). A DNA vaccine expressing dengue type 2 virus premembrane and envelope genes induces neutralizing antibody and memory B cells in mice. Vaccine 18(11–12):1133–1139. Konishi, E., Fujii, A., and Mason, P. W. (2001). Generation and characterization of a mammalian cell line continuously expressing Japanese encephalitis virus subviral particles. J. Virol. 75(5):2204–2212. Konishi, E., Terazawa, A., and Imoto, J. (2003). Simultaneous immunization with DNA and protein vaccines against Japanese encephalitis or dengue synergistically increases their own abilities to induce neutralizing antibody in mice. Vaccine 21(17–18):1826–1832. Konishi, E., Kosugi, S., and Imoto, J. (2006). Dengue tetravalent DNA vaccine inducing neutralizing antibody and anamnestic responses to four serotypes in mice. Vaccine 24 (12):2200–2207. Koraka, P., Benton, S., van Amerongen, G., Stittelaar, K. J., and Osterhaus, A. D. (2007). Efficacy of a live attenuated tetravalent candidate dengue vaccine in naive and previously infected cynomolgus macaques. Vaccine 25(29):5409–5416. Kunze, M., Suss, J., Baumhackl, U., and Freund, C. (2007). International Scientific Working Group on Tick-Borne Encephalitis. TBE: From Epidemiology to Vaccination Recommendations in 2007: New Issues – Best Practices. Ledizet, M., Kar, K., Foellmer, H. G., Wang, T., Bushmich, S. L., Anderson, J. F., Fikrig, E., and Koski, R. A. (2005). A recombinant envelope protein vaccine against West Nile virus. Vaccine 23(30):3915–3924. Lee, J. S., Groebner, J. L., Hadjipanayis, A. G., Negley, D. L., Schmaljohn, A. L., Welkos, S. L., Smith, L. A., and Smith, J. F. (2006). Multiagent vaccines vectored by Venezuelan equine encephalitis virus replicon elicits immune responses to Marburg virus and protection against anthrax and botulinum neurotoxin in mice. Vaccine 24(47–48):6886–6892. Lieberman, M. M., Clements, D. E., Ogata, S., Wang, G., Corpuz, G., Wong, T., Martyak, T., Gilson, L., Coller, B. A., Leung, J., Watts, D. M., Tesh, R. B., et al. (2007). Preparation and immunogenic properties of a recombinant West Nile subunit vaccine. Vaccine 25 (3):414–423. Lin, Y. L., Chen, L. K., Liao, C. L., Yeh, C. T., Ma, S. H., Chen, J. L., Huang, Y. L., Chen, S. S., and Chiang, H. Y. (1998). DNA immunization with Japanese encephalitis virus nonstructural protein NS1 elicits protective immunity in mice. J. Virol. 72(1):191–200. Lindenbach, B. D., Thiel, H. J., and Rice, C. M. (2007). Flaviviridae: The viruses and their replication. In ‘‘Fields Virology’’ (B. N. Fields, D. M. Knipe, and P. M. Howley, eds.). 5th Ed., Wolters Kluwer Health/Lippincott Williams & Wilkins, Philadelphia.
Third-Generation Flavivirus Vaccines
121
Liprandi, F. (1981). Isolation of plaque variants differing in virulence from the 17D strain of yellow fever virus. J. Gen. Virol. 56(Pt 2):363–370. Lorin, C., Mollet, L., Delebecque, F., Combredet, C., Hurtrel, B., Charneau, P., Brahic, M., and Tangy, F. (2004). A single injection of recombinant measles virus vaccines expressing human immunodeficiency virus (HIV) type 1 clade B envelope glycoproteins induces neutralizing antibodies and cellular immune responses to HIV. J. Virol. 78(1):146–157. Lyons, A., Kanesa-thasan, N., Kuschner, R. A., Eckels, K. H., Putnak, R., Sun, W., Burge, R., Towle, A. C., Wilson, P., Tauber, E., and Vaughn, D. W. (2007). A Phase 2 study of a purified, inactivated virus vaccine to prevent Japanese encephalitis. Vaccine 25 (17):3445–3453. Marchevsky, R. S., Freire, M. S., Coutinho, E. S., and Galler, R. (2003). Neurovirulence of yellow fever 17DD vaccine virus to rhesus monkeys. Virology 316(1):55–63. Markoff, L. (2000). Points to consider in the development of a surrogate for efficacy of novel Japanese encephalitis virus vaccines. Vaccine 18(Suppl. 2):26–32. Martin, M., Tsai, T. F., Cropp, B., Chang, G. J. J., Holmes, D. A., Tseng, J., Shieh, W. J., Zaki, S. R., Al, S. I., Cutrona, A. F., Ray, G., Weld, L. H., et al. (2001a). Fever and multisystem organ failure associated with 17D-204 yellow fever vaccination: A report of four cases. Lancet North Am. Ed. 358(9276):98–104. Martin, M., Weld, L. H., Tsai, T. F., Mootrey, G. T., Chen, R. T., Niu, M., and Cetron, M. S. (2001b). Advanced age a risk factor for illness temporally associated with yellow fever vaccination. Emerg. Infect. Dis. 7(6):945–951. Martin, J. E., Pierson, T. C., Hubka, S., Rucker, S., Gordon, I. J., Enama, M. E., Andrews, C. A., Xu, Q., Davis, B. S., Nason, M., Fay, M., Koup, R. A., et al. (2007). A West Nile virus DNA vaccine induces neutralizing antibody in healthy adults during a phase 1 clinical trial. J. Infect. Dis. 196(12):1732–1740. Martina, B. E., Koraka, P., van den Doel, P., van Amerongen, G., Rimmelzwaan, G. F., and Osterhaus, A. D. (2008). Immunization with West Nile virus envelope domain III protects mice against lethal infection with homologous and heterologous virus. Vaccine 26 (2):153–157. Mason, P. W., McAda, P. C., Dalrymple, J. M., Fournier, M. J., and Mason, T. L. (1987). Expression of Japanese encephalitis virus antigens in Escherichia coli. Virology 158 (2):361–372. Mason, P. W., Pincus, S., Fournier, M. J., Mason, T. L., Shope, R. E., and Paoletti, E. (1991). Japanese encephalitis virus-vaccinia recombinants produce particulate forms of the structural membrane proteins and induce high levels of protection against lethal JEV infection. Virology 180(1):294–305. Mason, P. W., Shustov, A. V., and Frolov, I. (2006). Production and characterization of vaccines based on flaviviruses defective in replication. Virology 351(2):432–443. McKee, K. T., Jr., Bancroft, W. H., Eckels, K. H., Redfield, R. R., Summers, P. L., and Russell, P. K. (1987). Lack of attenuation of a candidate dengue 1 vaccine (45AZ5) in human volunteers. Am. J. Trop. Med. Hyg. 36(2):435–442. Men, R. H., Bray, M., and Lai, C. J. (1991). Carboxy-terminally truncated dengue virus envelope glycoproteins expressed on the cell surface and secreted extracellularly exhibit increased immunogenicity in mice. J. Virol. 65(3):1400–1407. Men, R., Bray, M., Clark, D., Chanock, R. M., and Lai, C. J. (1996). Dengue type 4 virus mutants containing deletions in the 30 noncoding region of the RNA genome: analysis of growth restriction in cell culture and altered viremia pattern and immunogenicity in rhesus monkeys. J. Virol. 70(6):3930–3937. Mok, H., Lee, S., Utley, T. J., Shepherd, B. E., Polosukhin, V. V., Collier, M. L., Davis, N. L., Johnston, R. E., and Crowe, J. E., Jr. (2007). Venezuelan equine encephalitis virus replicon particles encoding respiratory syncytial virus surface glycoproteins induce protective mucosal responses in mice and cotton rats. J. Virol. 81(24):13710–13722.
122
Douglas G. Widman et al.
Monath, T. P. (2001). Prospects for development of a vaccine against the West Nile virus. Ann. N Y Acad. Sci. 951:1–12. Monath, T. (2004). Yellow fever vaccine. In ‘‘Vaccine’’ (S. Plotkin and W. A. Orenstein, eds.), 5th Ed., pp. 1095–1176. W.B. Saunders Company, Philadelphia, PA. Monath, T. P. (2008). Treatment of yellow fever. Antiviral Res. 78(1):116–124. Monath, T. P., Soike, K., Levenbook, I., Zhang, Z. X., Arroyo, J., Delagrave, S., Myers, G., Barrett, A. D., Shope, R. E., Ratterree, M., Chambers, T. J., and Guirakhoo, F. (1999). Recombinant, chimaeric live, attenuated vaccine (ChimeriVax) incorporating the envelope genes of Japanese encephalitis (SA14-14-2) virus and the capsid and nonstructural genes of yellow fever (17D) virus is safe, immunogenic and protective in non-human primates. Vaccine 17(15–16):1869–1882. Monath, T. P., Levenbook, I., Soike, K., Zhang, Z. X., Ratterree, M., Draper, K., Barrett, A. D. T., Nichols, R., Weltzin, R., Arroyo, J., and Guirakhoo, F. (2000). Chimeric yellow fever virus 17D-Japanese encephalitis virus vaccine: Dose-response effectiveness and extended safety testing in rhesus monkeys. J. Virol. 74(4):1742–1751. Monath, T. P., McCarthy, K., Bedford, P., Johnson, C. T., Nichols, R., Yoksan, S., Marchesani, R., Knauber, M., Wells, K. H., Arroyo, J., and Guirakhoo, F. (2002). Clinical proof of principle for ChimeriVax: Recombinant live, attenuated vaccines against flavivirus infections. Vaccine 20(7–8):1004–1018. Monath, T. P., Guirakhoo, F., Nichols, R., Yoksan, S., Schrader, R., Murphy, C., Blum, P., Woodward, S., McCarthy, K., Mathis, D., Johnson, C., and Bedford, P. (2003). Chimeric live, attenuated vaccine against Japanese encephalitis (ChimeriVax-JE): Phase 2 clinical trials for safety and immunogenicity, effect of vaccine dose and schedule, and memory response to challenge with inactivated Japanese encephalitis antigen. J. Infect. Dis. 188 (8):1213–1230. Monath, T. P., Liu, J., Kanesa-Thasan, N., Myers, G. A., Nichols, R., Deary, A., McCarthy, K., Johnson, C., Ermak, T., Shin, S., Arroyo, J., Guirakhoo, F., et al. (2006). A live, attenuated recombinant West Nile virus vaccine. Proc. Natl. Acad. Sci. USA 103(17):6694–6699. Morrison, L. A., and Knipe, D. M. (1994). Immunization with replication-defective mutants of herpes simplex virus type 1: Sites of immune intervention in pathogenesis of challenge virus infection. J. Virol. 68(2):689–696. Mota, J., Acosta, M., Argotte, R., Figueroa, R., Mendez, A., and Ramos, C. (2005). Induction of protective antibodies against dengue virus by tetravalent DNA immunization of mice with domain III of the envelope protein. Vaccine 23(26):3469–3476. Mutoh, E., Ishikawa, T., Takamizawa, A., Kurata, T., Sata, T., and Kojima, A. (2004). Japanese encephalitis subunit vaccine composed of virus-like envelope antigen particles purified from serum-free medium of a high-producer J12#26 cell clone. Vaccine 22(20):2599–2608. Myint, K. S., Gibbons, R. V., Perng, G. C., and Solomon, T. (2007). Unravelling the neuropathogenesis of Japanese encephalitis. Trans. R. Soc. Trop. Med. Hyg. 101(10):955–956. Nguyen, L. H., Knipe, D. M., and Finberg, R. W. (1992). Replication-defective mutants of herpes simplex virus (HSV) induce cellular immunity and protect against lethal HSV infection. J. Virol. 66(12):7067–7072. Ohtaki, E., Murakami, Y., Komori, H., Yamashita, Y., and Matsuishi, T. (1992). Acute disseminated encephalomyelitis after Japanese B encephalitis vaccination. Pediatr. Neurol. 8(2):137–139. Ohtaki, E., Matsuishi, T., Hirano, Y., and Maekawa, K. (1995). Acute disseminated encephalomyelitis after treatment with Japanese B encephalitis vaccine (Nakayama-Yoken and Beijing strains). J. Neurol. Neurosurg. Psychiatr. 59(3):316–317. Oya, A., and Kurane, I. (2007). Japanese encephalitis for a reference to international travelers. J. Travel Med. 14(4):259–268.
Third-Generation Flavivirus Vaccines
123
Phillpotts, R. J., Venugopal, K., and Brooks, T. (1996). Immunisation with DNA polynucleotides protects mice against lethal challenge with St. Louis encephalitis virus. Arch. Virol. 141(3–4):743–749. Plesner, A. M. (2003). Allergic reactions to Japanese encephalitis vaccine. Immunol. Allergy Clin. North Am. 23(4):665–697. Plesner, A. M., Arlien-Soborg, P., and Herning, M. (1996). Neurological complications and Japanese encephalitis vaccination. Lancet 348(9021):202–203. Pletnev, A. G., and Men, R. (1998). Attenuation of the Langat tick-borne flavivirus by chimerization with mosquito-borne flavivirus dengue type 4. Proc. Natl. Acad. Sci. USA 95(4):1746–1751. Pletnev, A. G., Bray, M., Huggins, J., and Lai, C. J. (1992). Construction and characterization of chimeric tick-borne encephalitis/dengue type 4 viruses. Proc. Natl. Acad. Sci. USA 89 (21):10532–10536. Pletnev, A. G., Bray, M., and Lai, C. J. (1993). Chimeric tick-borne encephalitis and dengue type 4 viruses: Effects of mutations on neurovirulence in mice. J. Virol. 67(8):4956–4963. Pletnev, A. G., Karganova, G. G., Dzhivanyan, T. I., Lashkevich, V. A., and Bray, M. (2000). Chimeric Langat/dengue viruses protect mice from heterologous challenge with the highly virulent strains of tick-borne encephalitis virus. Virology 274(1):26–31. Pletnev, A. G., Bray, M., Hanley, K. A., Speicher, J., and Elkins, R. (2001). Tick-borne Langat/ mosquito-borne dengue flavivirus chimera, a candidate live attenuated vaccine for protection against disease caused by members of the tick-borne encephalitis virus complex: Evaluation in rhesus monkeys and in mosquitoes. J. Virol. 75(17):8259–8267. Pletnev, A. G., Putnak, R., Speicher, J., Wagar, E. J., and Vaughn, D. W. (2002). West Nile virus/dengue type 4 virus chimeras that are reduced in neurovirulence and peripheral virulence without loss of immunogenicity or protective efficacy. Proc. Natl. Acad. Sci. USA 99(5):3036–3041. Pletnev, A. G., Claire, M. S., Elkins, R., Speicher, J., Murphy, B. R., and Chanock, R. M. (2003). Molecularly engineered live-attenuated chimeric West Nile/dengue virus vaccines protect rhesus monkeys from West Nile virus. Virology 314(1):190–195. Pletnev, A. G., Swayne, D. E., Speicher, J., Rumyantsev, A. A., and Murphy, B. R. (2006). Chimeric West Nile/dengue virus vaccine candidate: Preclinical evaluation in mice, geese and monkeys for safety and immunogenicity. Vaccine 24(40–41):6392–6404. Plisek, S., Honegr, K., and Beran, J. (2008). TBE infection in an incomplete immunized person at-risk who lives in a high-endemic area-Impact on current recommendations for immunization of high-risk groups. Vaccine 26(3):301–304. Pushko, P., Parker, M., Ludwig, G. V., Davis, N. L., Johnston, R. E., and Smith, J. F. (1997). Replicon-helper systems from attenuated Venezuelan equine encephalitis virus: Expression of heterologous genes in vitro and immunization against heterologous pathogens in vivo. Virology 239(2):389–401. Putnak, R., Feighny, R., Burrous, J., Cochran, M., Hackett, C., Smith, G., and Hoke, C. (1991). Dengue-1 virus envelope glycoprotein gene expressed in recombinant baculovirus elicits virus-neutralizing antibody in mice and protects them from virus challenge. Am. J. Trop. Med. Hyg. 45(2):159–167. Qiao, M., Ashok, M., Bernard, K. A., Palacios, G., Zhou, Z. H., Lipkin, W. I., and Liang, T. J. (2004). Induction of sterilizing immunity against West Nile Virus (WNV), by immunization with WNV-like particles produced in insect cells. J. Infect. Dis. 190(12):2104–2108. Raengsakulrach, B., Nisalak, A., Gettayacamin, M., Thirawuth, V., Young, G. D., Myint, K. S., Ferguson, L. M., Hoke, C. H., Jr., Innis, B. L., and Vaughn, D. W. (1999). Safety, immunogenicity, and protective efficacy of NYVAC-JEV and ALVAC-JEV recombinant Japanese encephalitis vaccines in rhesus monkeys. Am. J. Trop. Med. Hyg. 60(3):343–349. Raja, N. U., Holman, D. H., Wang, D., Raviprakash, K., Juompan, L. Y., Deitz, S. B., Luo, M., Zhang, J., Porter, K. R., and Dong, J. Y. (2007). Induction of bivalent immune responses by
124
Douglas G. Widman et al.
expression of dengue virus type 1 and type 2 antigens from a single complex adenoviral vector. Am. J. Trop. Med. Hyg. 76(4):743–751. Raviprakash, K., Apt, D., Brinkman, A., Skinner, C., Yang, S., Dawes, G., Ewing, D., Wu, S. J., Bass, S., Punnonen, J., and Porter, K. (2006). A chimeric tetravalent dengue DNA vaccine elicits neutralizing antibody to all four virus serotypes in rhesus macaques. Virology 353 (1):166–173. Robert Putnak, J., Coller, B. A., Voss, G., Vaughn, D. W., Clements, D., Peters, I., Bignami, G., Houng, H. S., Chen, R. C., Barvir, D. A., Seriwatana, J., Cayphas, S., et al. (2005). An evaluation of dengue type-2 inactivated, recombinant subunit, and live-attenuated vaccine candidates in the rhesus macaque model. Vaccine 23(35):4442–4452. Robinson, H. C., Russell, M. L., and Csokonay, W. M. (1991). Japanese encephalitis vaccine and adverse effects among travellers. Can. Dis. Wkly. Rep. 17(32):173–174177. Rothman, A. L. (2003). Immunology and immunopathogenesis of dengue disease. Adv. Virus Res. 60:397–419. Sabchareon, A., Lang, J., Chanthavanich, P., Yoksan, S., Forrat, R., Attanath, P., Sirivichayakul, C., Pengsaa, K., Pojjaroen-Anant, C., Chokejindachai, W., Jagsudee, A., Saluzzo, J. F., et al. (2002). Safety and immunogenicity of tetravalent live-attenuated dengue vaccines in Thai adult volunteers: role of serotype concentration, ratio, and multiple doses. Am. J. Trop. Med. Hyg. 66(3):264–272. Sabchareon, A., Lang, J., Chanthavanich, P., Yoksan, S., Forrat, R., Attanath, P., Sirivichayakul, C., Pengsaa, K., Pojjaroen-Anant, C., Chambonneau, L., Saluzzo, J. F., and Bhamarapravati, N. (2004). Safety and immunogenicity of a three dose regimen of two tetravalent live-attenuated dengue vaccines in five- to twelve-year-old Thai children. Pediatr. Infect. Dis. J. 23(2):99–109. Sabin, A. B. (1952). Research on dengue during World War II. Am. J. Trop. Med. Hyg. 1:30–50. Sakaguchi, M., Miyazawa, H., and Inouye, S. (2001). Specific IgE and IgG to gelatin in children with systemic cutaneous reactions to Japanese encephalitis vaccines. Allergy 56 (6):536–539. Schepp-Berglind, J., Luo, M., Wang, D., Wicker, J. A., Raja, N. U., Hoel, B. D., Holman, D. H., Barrett, A. D., and Dong, J. Y. (2007). Complex adenovirus-mediated expression of West Nile virus C, PreM, E, and NS1 proteins induces both humoral and cellular immune responses. Clin. Vaccine Immunol. 14(9):1117–1126. Schlesinger, J. J., Brandriss, M. W., Cropp, C. B., and Monath, T. P. (1986). Protection against yellow fever in monkeys by immunization with yellow fever virus nonstructural protein NS1. J. Virol. 60(3):1153–1155. Scholle, F., Girard, Y. A., Zhao, Q., Higgs, S., and Mason, P. W. (2004). Trans-packaged West Nile virus-like particles: Infectious properties in vitro and in infected mosquito vectors. J. Virol. 78:11605–11614. Sejvar, J. J. (2007). The long-term outcomes of human West Nile virus infection. Clin. Infect. Dis. 44(12):1617–1624. Sellards, A. W., and Langret, J. (1932). Vaccination de l’homme contre la fievre jaune. C. R. Acad. Sci. 194:1609–1611. Seregin, A., Nistler, R., Borisevich, V., Yamshchikov, G., Chaporgina, E., Kwok, C. W., and Yamshchikov, V. (2006). Immunogenicity of West Nile virus infectious DNA and its noninfectious derivatives. Virology 356(1–2):115–125. Shustov, A. V., Mason, P. W., and Frolov, I. (2007). Production of pseudoinfectious yellow fever virus with a two-component genome. J. Virol. 81(21):11737–11748. Simmons, M., Nelson, W. M., Wu, S. J., and Hayes, C. G. (1998). Evaluation of the protective efficacy of a recombinant dengue envelope B domain fusion protein against dengue 2 virus infection in mice. Am. J. Trop. Med. Hyg. 58(5):655–662.
Third-Generation Flavivirus Vaccines
125
Simmons, M., Porter, K. R., Hayes, C. G., Vaughn, D. W., and Putnak, R. (2006). Characterization of antibody responses to combinations of a dengue virus type 2 DNA vaccine and two dengue virus type 2 protein vaccines in rhesus macaques. J. Virol. 80(19):9577–9585. Smith, T. J., Brandt, W. E., Swanson, J. L., McCown, J. M., and Buescher, E. L. (1970). Physical and biological properties of dengue-2 virus and associated antigens. J. Virol. 5(4):524–532. Sohn, Y. M., Park, M. S., Rho, H. O., Chandler, L. J., Shope, R. E., and Tsai, T. F. (1999). Primary and booster immune responses to SA14-14-2 Japanese encephalitis vaccine in Korean infants. Vaccine 17(18):2259–2264. Srivastava, A. K., Putnak, J. R., Warren, R. L., and Hoke, C. H., Jr. (1995). Mice immunized with a dengue type 2 virus E and NS1 fusion protein made in Escherichia coli are protected against lethal dengue virus infection. Vaccine 13(13):1251–1258. Srivastava, A. K., Putnak, J. R., Lee, S. H., Hong, S. P., Moon, S. B., Barvir, D. A., Zhao, B., Olson, R. A., Kim, S. O., Yoo, W. D., Towle, A. C., Vaughn, D. W., et al. (2001). A purified inactivated Japanese encephalitis virus vaccine made in Vero cells. Vaccine 19(31): 4557–4565. Stevens, T. M., and Schlesinger, R. W. (1965). Studies on the nature of dengue viruses. I. Correlation of particle density, infectivity, and RNA content of type 2 virus. Virology 27 (1):103–112. Sun, W., Edelman, R., Kanesa-Thasan, N., Eckels, K. H., Putnak, J. R., King, A. D., Houng, H. S., Tang, D., Scherer, J. M., Hoke, C. H., Jr., and Innis, B. L. (2003). Vaccination of human volunteers with monovalent and tetravalent live-attenuated dengue vaccine candidates. Am. J. Trop. Med. Hyg. 69(6 Suppl.):24–31. Sun, W., Nisalak, A., Gettayacamin, M., Eckels, K. H., Putnak, J. R., Vaughn, D. W., Innis, B. L., Thomas, S. J., and Endy, T. P. (2006). Protection of Rhesus monkeys against dengue virus challenge after tetravalent live attenuated dengue virus vaccination. J. Infect. Dis. 193(12):1658–1665. Tauber, E., Kollaritsch, H., Korinek, M., Rendi-Wagner, P., Jilma, B., Firbas, C., Schranz, S., Jong, E., Klingler, A., Dewasthaly, S., and Klade, C. S. (2007). Safety and immunogenicity of a Vero-cell-derived, inactivated Japanese encephalitis vaccine: a non-inferiority, phase III, randomised controlled trial. Lancet 370(9602):1847–1853. Theiler, M., and Smith, H. H. (1937a). The effect of prolonged cultivation in vitro upon the pathogenicity of yellow fever virus. J. Exp. Med. 65(6):767–786. Theiler, M., and Smith, H. H. (1937b). The use of yellow fever virus modified by in vitro cultivation for human immunization. J. Exp. Med. 65(6):787–800. Tsai, T. F. (2000). New initiatives for the control of Japanese encephalitis by vaccination: Minutes of a WHO/CVI meeting, Bangkok, Thailand, 13–15 October 1998. Vaccine 18 (Suppl. 2):1–25. USDA. (2005). USDA Issues License for West Nile Virus DNA Vaccine for Horses USDA Animal and Plant Health Inspection Service, Washington, DC. Vasconcelos, P. F. C., Luna, E. J., Galler, R., Silva, L. J., Coimbra, T. L., Barros, V. L. R. S., Monath, T. P., Rodigues, S. G., Laval, C., Costa, Z. G., Vilela, M. F. G., Santos, C. L. S., et al. (2001). Serious adverse events associated with yellow fever 17DD vaccine in Brazil: A report of two cases. Lancet North Am. Ed. 358(9276):91–97. Wang, T., Anderson, J. F., Magnarelli, L. A., Wong, S. J., Koski, R. A., and Fikrig, E. (2001). Immunization of mice against West Nile virus with recombinant envelope protein. J. Immunol. 167(9):5273–5277. Watts, D. M., Tesh, R. B., Siirin, M., Rosa, A. T., Newman, P. C., Clements, D. E., Ogata, S., Coller, B. A., Weeks-Levy, C., and Lieberman, M. M. (2007). Efficacy and durability of a recombinant subunit West Nile vaccine candidate in protecting hamsters from West Nile encephalitis. In ‘‘Vaccine’’, Vol. 25, pp. 2913–2918. White, L. J., Parsons, M. M., Whitmore, A. C., Williams, B. M., de Silva, A., and Johnston, R. E. (2007). An immunogenic and protective alphavirus replicon particle-based dengue
126
Douglas G. Widman et al.
vaccine overcomes maternal antibody interference in weanling mice. J. Virol. 81 (19):10329–10339. Whitehead, S. S., Falgout, B., Hanley, K. A., Blaney, J. E., Jr., Markoff, L., and Murphy, B. R. (2003a). A live, attenuated dengue virus type 1 vaccine candidate with a 30-nucleotide deletion in the 30 untranslated region is highly attenuated and immunogenic in monkeys. J. Virol. 77(2):1653–1657. Whitehead, S. S., Hanley, K. A., Blaney, J. E., Jr., Gilmore, L. E., Elkins, W. R., and Murphy, B. R. (2003b). Substitution of the structural genes of dengue virus type 4 with those of type 2 results in chimeric vaccine candidates which are attenuated for mosquitoes, mice, and rhesus monkeys. Vaccine 21(27–30):4307–4316. WHO. (1945). Standards for the manufacture and control of yellow fever vaccine. Epidemiol. Inf. Bull. 1(365). WHO. (1959). Requirements for biological substances. 3. Requirements for yellow fever vaccine. 4. Requirements for cholera vaccine. Report of a study group. World Health Organ. Tech. Rep. Ser. 179:1–46. WHO. (1976). WHO Expert Committee on Biological Standardization. Twenty-seventh report. World Health Organ. Tech. Rep. Ser. (594):23–49. WHO. (1998). WHO Expert Committee on Biological Standardization. Forty-sixth Report. World Health Organ. Tech. Rep. Ser. 872(i–vii):1–90. WHO. (2005). Global Advisory Committee on Vaccine Safety, 9–10 June 2005. Wkly Epidemiol Rec 80(28):242–247. WHO. (2006). Japanese encephalitis vaccines. Wkly. Epidemiol. Rec. 81:331–340. WHO. (2007). Epidemiological trends and current situation of yellow fever. Widman, D. G., Ishikawa, T., Fayzulin, R., Bourne, N., and Mason, P. W. (2008). Construction and characterization of a second-generation pseudoinfectious West Nile virus vaccine propagated using a new cultivation system. Vaccine 26:2762–2771. Wright, P. F., Ankrah, S., Henderson, S. E., Durbin, A. P., Speicher, J., Whitehead, S. S., Murphy, B. R., and Pletnev, A. G. (2008). Evaluation of the Langat/dengue 4 chimeric virus as a live attenuated tick-borne encephalitis vaccine for safety and immunogenicity in healthy adult volunteers. Vaccine 26(7):882–890. Wu, S. C., Yu, C. H., Lin, C. W., and Chu, I. M. (2003). The domain III fragment of Japanese encephalitis virus envelope protein: Mouse immunogenicity and liposome adjuvanticity. Vaccine 21(19–20):2516–2522. Yasuda, A., Kimura-Kuroda, J., Ogimoto, M., Miyamoto, M., Sata, T., Sato, T., Takamura, C., Kurata, T., Kojima, A., and Yasui, K. (1990). Induction of protective immunity in animals vaccinated with recombinant vaccinia viruses that express PreM and E glycoproteins of Japanese encephalitis virus. J. Virol. 64(6):2788–2795. Yoshii, K., Goto, A., Kawakami, K., Kariwa, H., and Takashima, I. (2008). Construction and application of chimeric virus-like particles of tick-borne encephalitis virus and mosquitoborne Japanese encephalitis virus. J. Gen. Virol. 89(Pt 1):200–211. Zhang, Y. M., Hayes, E. P., McCarty, T. C., Dubois, D. R., Summers, P. L., Eckels, K. H., Chanock, R. M., and Lai, C. J. (1988). Immunization of mice with dengue structural proteins and nonstructural protein NS1 expressed by baculovirus recombinant induces resistance to dengue virus encephalitis. J. Virol. 62(8):3027–3031.
CHAPTER
3 Swine Influenza Viruses: A North American Perspective Amy L. Vincent,* Wenjun Ma,*,†,‡ Kelly M. Lager,* Bruce H. Janke,† and Ju¨rgen A. Richt*,‡
Contents
Abstract
I. Introduction to Influenza A Viruses A. The Virus B. Influenza A Virus Infection of Pigs II. Evolution of North American SI Viruses of the H1 and H3 Subtype III. Cross-Species Transmission of Influenza A Viruses and Novel Subtypes in North American Swine A. SI Infections of Humans B. Novel SI Isolates in North America IV. Vaccination of Pigs Against SI V. Conclusions and Outlook Acknowledgments References
128 128 130 132 136 136 137 139 147 148 148
Influenza is a zoonotic viral disease that represents a health and economic threat to both humans and animals worldwide. Swine influenza (SI) was first recognized clinically in pigs in the Midwestern U.S., in 1918, coinciding with the human influenza pandemic known as the Spanish flu. Since that time SI has remained of importance to the swine industry throughout the world. In this
* Virus and Prion Diseases of Livestock Research Unit, National Animal Disease Center, USDA-ARS, Ames,
Iowa 50010 Veterinary Diagnostic and Production Animal Medicine, College of Veterinary Medicine, Iowa State University, Ames, Iowa 50011 { Diagnostic Medicine and Pathobiology Department, College of Veterinary Medicine, Kansas State University, Manhattan, KS 66506 {
Advances in Virus Research, Volume 72 ISSN 0065-3527, DOI: 10.1016/S0065-3527(08)00403-X
#
2008 Elsevier Inc. All rights reserved.
127
128
Amy L. Vincent et al.
review, the epidemiology of swine influenza virus (SIV) infection in North American pigs is described in detail. The first 80 years of SI remained relatively static, whereas the last decade has become dynamic with the establishment of many emerging subtypes. With the increasing number of novel subtypes and genetic variants, the control of SI has become increasingly difficult and innovative strategies to combat this economically important zoonotic disease are critical. Therefore, protective immune responses against influenza virus infections as well as new paradigms of vaccine development in pigs are discussed in the review. It is expected that the dynamic evolutionary changes of SIVs in North American pigs will continue, making currently available prophylactic approaches of limited use to control the spread and economic losses associated with this important swine pathogen.
I. INTRODUCTION TO INFLUENZA A VIRUSES Influenza is a zoonotic viral disease that represents a health and economic threat to both humans and animals worldwide. Influenza A viruses infect a wide variety of species and exhibit only a partial restriction of their host range, that is, there is occasional transmission from one species to another. Annual epidemics/epizootics in humans and animals and occasional influenza pandemics in humans depend on the continued molecular evolution of influenza viruses giving rise to new antigenic variants. The surface hemagglutinin (HA) and neuraminidase (NA) antigens undergo two types of variation called antigenic drift and antigenic shift. Antigenic drift involves minor changes in the HA and NA, whereas antigenic shift involves major changes in these molecules resulting from replacement of the entire gene segment. The segmented nature of the influenza virus genome is a key feature of influenza viruses and supports antigenic shift or reassortment. In the event that cells are infected with two (or more) different influenza viruses, exchange of RNA segments between the viruses allows the generation of progeny viruses containing a novel combination of genes. In mammals, influenza viruses replicate mainly in the respiratory tract, usually accompanied with clinical signs, whereas in avian species, the major replication site is the intestinal tract without clinical signs (Webster, 2002). In aquatic birds, influenza viruses are generally highly host-adapted and show low evolutionary rates (Webby and Webster, 2001), whereas in mammalian species the evolutionary rate is much greater (Buonagurio et al., 1986).
A. The Virus Influenza viruses are members of the family Orthomyxoviridae comprising five genera: Influenza A, B and C viruses, Thogotovirus, and Isavirus (Knipe et al., 2007). Of these, only influenza A viruses are true zoonotic
SIV: A North American Perspective
129
Lipid bilayer PB2 PB1
M2 NA
PA HA
M1
NP NA M1 + M2
HA
NS1 + NS2
FIGURE 1 Diagram of an influenza A virion with the viral envelope and the eight RNA gene segments.
agents. Influenza B and C viruses are primarily human pathogens; influenza C can occasionally infect pigs and dogs (Ohwada et al., 1987). Influenza A viruses are 80–120 nm enveloped viruses with segmented, single-stranded, negative-sense RNA genomes (Fig. 1). The eight RNA segments within the viral genome, varying in length between 890 and 2341 nucleotides, encode 10 and in some cases 11 proteins. Segment 7 (Matrix, M) and segment 8 (Nonstructural, NS) encode two proteins (M1/ M2 and NS1/NS2; Knipe et al., 2007) due to differentially spliced transcripts, and in some virus strains segment 2 (polymerase basic 1, PB1) encodes a second short protein, called PB1-F2, from an additional openreading frame (Conenello and Palese, 2007). The RNA fragments are bound and protected by the viral nucleoprotein (NP; Compans et al., 1972). The trimeric RNA polymerase complex (PB1, polymerase basic 2, PB2 and polymerase acidic, PA) binds to the 50 and 30 ends of the viral RNA forming a noncovalent circular complex (Klumpp et al., 1997). The complex consisting of viral RNA, the polymerase complex, and the NP is called the ribonucleoprotein (RNP) complex. Influenza A viruses are typed according to their surface glycoproteins, HA and NA. The HA and NA are also the main targets of the host humoral immune response. Host immune pressure is the driving force in selecting mutant viruses with amino acid substitutions, a process called antigenic drift. The HA serves as the viral receptor-binding protein and mediates fusion of the
130
Amy L. Vincent et al.
virus envelope with the host cell membrane (Skehel and Wiley, 2000). The HA binds to N-acetylneuraminic acid-2,3-galactose linkage or N-acetylneuraminic acid-2,6-galactose linkage on sialyloligosaccharides for avian and mammalian viruses, respectively (Rogers and Paulson, 1983). The NA is responsible for cleaving terminal sialic acid residues from carbohydrate moieties on the surfaces of the host cell and virus (Gottschalk, 1957), thus assisting in virus cell entry by mucus degradation (Matrosovich et al., 2004) and the release and spread of progeny virions (Palese et al., 1974). Like the HA, the NA undergoes substantial antigenic variation in response to immune pressure. The M2 protein, the third envelope glycoprotein present in the influenza virion, serves as an ion channel (Wang et al., 1993). The M1 protein is the most abundant protein present in the influenza virion and lies beneath the lipid envelope (Fig. 1). Influenza viruses encode two nonstructural (NS) proteins, NS1 and NS2. While the NS2 or nuclear export protein (NEP) was originally thought to be a nonstructural protein; it has since been found to be a part of the influenza virion (Richardson and Akkina, 1991). In contrast, although NS1 is abundantly present in infected cells during virus replication, the protein is not incorporated into the progeny virions (Palese et al., 1999).
B. Influenza A Virus Infection of Pigs Swine influenza (SI) was first recognized clinically in pigs in the Midwestern U.S. in summer/fall of 1918 (Koen, 1919), coinciding with the human influenza pandemic known as the Spanish flu (Webster, 2002). Since then SI has been of importance to the swine industry throughout the world (Olsen, 2002). The first SI virus (SIV) isolated from pigs in 1930 (Shope, 1931) belonged to the H1N1 lineage of SIVs. Clinical signs of influenza in pigs are similar to those observed in humans, making it an important model to study influenza pathogenesis in a natural host. Specifically, SIV infections are manifested as acute respiratory disease characterized by fever, inactivity, decreased food intake, respiratory distress, coughing, sneezing, conjunctivitis, and nasal discharge (Alexander and Brown, 2000; McQueen et al., 1968; Richt et al., 2003). Although the severity is affected by many factors, including viral strain, the onset of disease is typically sudden. The disease incubation period is between 1 and 3 days with rapid recovery beginning 4–7 days after onset. SI is a herd disease characterized by high morbidity (approaching 100%) and generally low mortality (<1%) rates. Macroscopically, SIV-infected lungs display a purple-red, multifocal to coalescing consolidation of predominantly cranio-ventral portions of the lung (Fig. 2A). Microscopic changes in the lung consist of necrosis of bronchiolar epithelial cells and sloughing of these cells into airway lumen, which often contains cellular debris, proteinaceous fluid and a few leukocytes (Fig. 2B). This necrosis
SIV: A North American Perspective
131
FIGURE 2 Macroscopic and microscopic pneumonia associated with swine H1N1 influenza virus infection of pigs. (A) Macroscopic pneumonia characterized as purple-red consolidation located primarily in cranial and middle lung lobes. (B) Necrotizing bronchiolitis in a pig 3 days postinoculation with SIV. Necrosis and reactive proliferation of epithelial cells is occurring with sloughing of these cells into the airway lumen which contains cellular debris, proteinaceous fluid and a few leukocytes.
is accompanied by peribronchiolar lymphocytic infiltration and interstitial pneumonia of variable severity. In recovery, bronchiolar epithelium becomes proliferative and lymphocytic cuffing becomes more prominent. Influenza viruses are part of the porcine respiratory disease complex (PRDC), acting in concert with other pathogens such as Mycoplasma
132
Amy L. Vincent et al.
hyopneumoniae, Actinobacillus pleuropneumonia, Pasteurella multocida, porcine reproductive and respiratory syndrome virus (PRRSV), and porcine circovirus type 2 (PCV-2; Ellis et al., 2004; Thacker et al., 2001). Current human influenza viruses are believed to have arisen by genetic reassortment between pre-existing human influenza viruses and nonhuman primarily avian influenza viruses. Swine have been considered a potential ‘‘mixing vessel’’ (Scholtissek, 1995), because they have receptors for both avian and human influenza viruses (Ito, 2000; Ito et al., 1998). Therefore, they can serve as hosts for viruses from either birds or humans.
II. EVOLUTION OF NORTH AMERICAN SI VIRUSES OF THE H1 AND H3 SUBTYPE Historically, SI in the United States had a predictable pattern with an epizootic in the late fall and early winter months similar to that in humans. Prior to 1998, this acute respiratory disease was almost exclusively caused by viruses of the classical-swine H1N1 lineage (cH1N1; Easterday and van Reeth, 1999). The cH1N1 virus, first isolated and identified in North America in 1930 (Shope, 1931), is believed to have been introduced into the U.S. pig population during the 1918 Spanish influenza pandemic since a concurrent disease similar to that of people was described in the pig population (Fig. 3A). For nearly 70 years, SIV in North America was relatively stable with the cH1N1 as the only predominant subtype. However, serological evidence indicated that human subtype H3 influenza viruses were circulating at a low frequency in U.S. pigs (Chambers et al., 1991), but failed to establish a stable lineage (Fig. 3A). In 1998, a severe influenza-like illness was observed in pigs on a farm in North Carolina with additional outbreaks in swine herds in Minnesota, Iowa, and Texas. The causative agents for these outbreaks were identified as influenza viruses of the subtype H3N2. Genetic analysis of these H3N2 viruses showed that at least two different genotypes were present (Fig. 3A). The initial North Carolina isolate contained gene segments similar to those of the human (HA, NA, PB1) and classical-swine (NS, NP, M, PB2, PA) lineages (double reassortant), whereas the isolates from Minnesota, Iowa, and Texas contained genes from the human (HA, NA, PB1), swine (NS, NP, M), and avian (PB2, PA) lineages (triple reassortant; Zhou et al., 1999). By the end of 1999, viruses antigenically and genetically related to the triple reassortant lineage were widespread in the U.S. swine population (Webby et al., 2000), whereas the double reassortant virus did not spread efficiently among swine. The double and triple reassortant H3N2 viruses contained similar HA genes with identical residues in critical receptor binding regions, suggesting that their different successes were due to factors not associated with the HA and receptor binding. The
SIV: A North American Perspective
133
A
H3N2
H3N2
1930 1918 Spanish flu pandemic
H1N1 evolves
H4N6
1997–98 Double reassortant H3N2
cH1N1
Triple reassortant H3N2
H3N1
H2N3
2003–05
2006
Current
hu-H1N1 hu-H1N2
rH2N3
H3N2 H1N2 rH1N1 cH1N1 huH1N1 huH1N2
B HA NA PB1 NS NP M PB2 PA
HA NA PB1 NS NP M PB2 PA
H3H2
HA NA PB1 NS NP M PB2 PA
H3N2
H1H1
HA NA PB1 NS NP M PB2 PA
HA NA PB1 NS NP M PB2 PA
rH1H1
H1H2 HA NA PB1 NS NP M PB2 PA
huH1N1
HA NA PB1 NS NP M PB2 PA
HA NA PB1 NS NP M PB2 PA
huH1N2
huH1N1
FIGURE 3 Epidemiology of SIVs in North America since 1918. Swine virus lineage is color coded pink, avian lineage is coded green, human lineage is coded blue or purple. (A) Chronology of transmission events leading to reassortant viruses with genes from
134
Amy L. Vincent et al.
major difference between the two viruses was the acquisition of two avian polymerase genes (PA, PB2) in the triple reassortant H3N2 (Fig. 3B). Once established in the swine population, the H3N2 viruses evolved through genetic mutation and reassortment with cH1N1 swine viruses. Currently, there are a number of reassortant viruses that have been identified, including further H3N2 genotypes (Richt et al., 2003; Webby et al., 2000, 2004), H1N2 (Choi et al., 2002; Karasin et al., 2002), reassortant H1N1 (rH1N1; Webby et al., 2004), and H3N1 viruses (Lekcharoensuk et al., 2006; Ma et al., 2006; Fig. 3B). The H3N2, rH1N1, and H1N2 viruses have become endemic and co-circulate in most major swine producing regions of both the U.S. and Canada. More recently, introductions of human-like H1 viruses that are genetically and antigenically distinct from the classical swine H1 lineage were identified in pigs in Canada (Karasin et al., 2006). All of the successful SIV reassortants that have become endemic in the U.S. pig population that have been characterized to date contain a similar triple reassortant internal gene (TRIG) cassette including the PA and PB2 genes of avian lineage, NS, NP, and M genes of classical swine lineage, and the PB1 gene of human lineage (Fig. 3B). This would suggest that the TRIG cassette can accept multiple HA and NA types and may endow a selective advantage to swine viruses possessing this gene constellation. With the acquisition of the avian PA and PB2 genes and the human PB1 gene, the current swine viruses appear to have increased the rate of antigenic drift and reassortment, and thereby, the ability to evade established herd immunity. This was not seen with the classical swine H1N1, which remained relatively stable antigenically for nearly 70 years (Luoh et al., 1992; Noble et al., 1993; Olsen et al., 1993; Sheerar et al., 1989). Classical swine H1N1 isolated as recently as 1999 maintained moderate to good cross-reactivity with viruses isolated decades earlier (Vincent et al., 2006).
swine, human and avian influenza virus (AIV) lineages. The ‘‘Spanish flu’’ virus was transmitted from avian/human origin to pigs and evolved into the cH1N1, as indicated by the transition in color of pigs from blue to purple to red to pink. The human and avian images above the horizontal timeline represent the species origin of viral gene segments donated to give rise to the swine influenza virus (SIV) reassortants listed below the horizontal timeline. Note, timeline not drawn to scale. (B) Diagram of viruses with their eight gene segment constellations in currently circulating SIVs. The triple reassortant H3N2 reassorted with the cH1N1 to produce rH1N1 and H1N2 subtype viruses with the triple reassortant internal gene (TRIG) cassette. Subsequent reassortment events with human H1 subtype viruses led to the human-like H1N2 and human-like H1N1 SIVs. The TRIG cassette is highlighted by the gray box.
SIV: A North American Perspective
135
H3N2 SIV isolated since 1998 have been evaluated at the genetic and antigenic level (Richt et al., 2003; Webby et al., 2004) and were demonstrated to have arisen from at least three introductions of human H3-subtype viruses, leading to phylogenetic clusters I, II, and III. There was variable antigenic cross-reactivity between the clusters. The cluster III viruses have become dominant in North America (Gramer, 2007) and have continued to evolve into cluster III variants, also known as cluster IV (Olsen et al., 2006). We have evaluated and compared the pathogenesis of 10 H1 SIV isolates dating from 1930 to more recent isolates (Vincent et al., 2006). In addition, the HA and NA genes of each isolate were sequenced for genetic comparison, and serological cross-reactivity was evaluated using sera and virus combinations in HI assays. Differences in pathogenicity were detected between H1 isolates, with recent isolates tending to produce more severe disease, increased nasal shedding, and higher virus titers in the lung. Serologically, the historical classical viruses tended to have better cross-reaction between historical sera and antigens, with moderate to good cross-reactivity with modern viral antigens. However, the modern sera were less reactive to historical viruses and tended to be less consistent in cross-reactivity within the modern group. There appeared to be an increase in genetic and antigenic diversity coincident with the emergence of the swine triple reassortant H3N2 in 1998 and the acquisition of the TRIG cassette. Many of the recent isolates had accumulated amino acid changes in the predicted antigenic and receptor binding sites on the HA protein. Existence of antigenic diversity in H1N1 and H1N2 SIVs is similar to the observations made in the diversity of the triple reassortant H3N2 SIV (Richt et al., 2003; Vincent et al., 2006). Since 2005, H1N1 and H1N2 viruses with the HA gene derived from human viruses have spread across the U.S. in swine herds (Gramer, 2007). The HA from the human-like swine H1 (hu-H1) viruses are genetically and antigenically distinct from swine H1 viruses. However, the six internal genes appear to be similar to those found in the TRIG cassette of contemporary swine triple reassortant viruses (Vincent, unpublished results). The NAs from these newly emerged viruses also are primarily human lineage N1 or N2. The hu-H1 SIVs have become one of the major types of SIV isolated and characterized from swine respiratory disease outbreaks (Gramer, personal communication). We evaluated one hu-H1N1 isolate in our experimental infection and transmission model and demonstrated that it was pathogenic and transmissible in 4-week-old pigs (Vincent, unpublished results). The hu-H1N1 isolate evaluated in our studies demonstrated differences in kinetics of lung lesion development, viral load in the lung, and nasal shedding when compared to a virulent rH1N1 SIV. These studies suggest this emerging virus genotype may not be fully adapted to the swine host since virus replication in the lung and virus shedding from the nose were reduced compared to the contemporary rH1N1 SIV. Nonetheless, the hu-H1 viruses have become established in the U.S. pig population.
136
Amy L. Vincent et al.
III. CROSS-SPECIES TRANSMISSION OF INFLUENZA A VIRUSES AND NOVEL SUBTYPES IN NORTH AMERICAN SWINE Influenza A viruses of all 16 HA and 9 NA subtypes have been recovered from wild waterfowl and seabirds (Fouchier et al., 2005; Webster et al., 1992). From these studies it was concluded that waterfowl provide a vast global reservoir of influenza viruses in nature from which novel viruses can emerge and infect mammalian species (Webby and Webster, 2001). Prominent examples of cross-species transmission of influenza viruses from avian to mammalian species or vice versa are the recent infections of humans, cats, and martens with the highly pathogenic avian H5N1 viruses (Klopfleisch et al., 2007; Tiensin et al., 2005; Webster et al., 2006) and the transmission of triple reassortant H3N2 SIVs to turkeys (Yassine et al., 2007). The outbreak of severe respiratory disease in racing greyhounds due to infection with an H3N8 influenza virus closely related to an equine influenza virus (Crawford et al., 2005) represents an intra-mammalian crossspecies transmission of influenza viruses. Cross-species spill-over of influenza viruses occur rather frequently; however, they tend to be self-limiting and the viruses are rarely maintained in the new host species (Webster et al., 1992; Webster, 2002). As discussed previously, the segmented nature of the influenza virus genome is a key feature for influenza virus evolution and cross-species transmissibility. However, specific subtypes differ in their ability to cross species barriers (Brown, 2000). Viral and host factors obviously play a role in cross-species transmissions and experimental evidence suggest that all eight gene segments, not only the surface proteins HA and NA, as well as specific gene combinations are involved in influenza virus species specificity (Horimoto and Kawaoka, 2001; Neumann and Kawaoka, 2006; Scholtissek et al., 1985). Given the plasticity of the virus genome, influenza fulfills the prerequisites of a virus with emerging disease potential (Webster et al., 1993). It is highly likely that sometime in the near future a ‘‘new’’ influenza A virus, for example, one of the H5N1 viruses currently circulating in the wild bird population in large parts of Asia or a different virus, will be able to emerge from its animal reservoir to cause widespread disease in mammalian species.
A. SI Infections of Humans In a recent review by Myers and colleagues (Myers et al., 2007) the entire literature on cases of SI in humans was reviewed. These authors reported that 50 cases of zoonotic SIV infections, 37 civilian cases and 13 military personnel cases, are described in the literature. The majority belonged to the H1N1 subtype, a few to the H3N2 subtype. The case-fatality rate of all reported cases was 14% (7/50). Civilian cases were described in the U.S.
SIV: A North American Perspective
137
(19 cases), Czechoslovakia (6 cases), The Netherlands (4 cases), Russia and Switzerland (3 cases each), and Canada and Hong Kong (1 case each). The median age of the patients was 24.5 years and the majority of the patients (61%) reported a recent exposure to pigs. A well publicized outbreak of SI due to an H1N1 virus resulted in 1 death and respiratory illness in 12 soldiers at Fort Dix, NJ, in early 1976 (Gaydos et al., 1977). Interestingly, no evidence of exposure to pigs was ever found. It has since been shown, however, that persons who work with swine are at increased risk of zoonotic influenza virus infection (Myers et al., 2006). Farmers, meat processing workers, and veterinarians were studied and all three exposed study groups demonstrated elevated serologic titers and higher odds for exposure to H1N1 and/or H1N2 SIV isolates, compared with control subjects. This indicates that occupational exposure to pigs greatly increases workers’ risk of SIV infection. Recently, an H1N1 SIV infection of pigs and people at an Ohio county fair was reported (Swenson, 2008). Pigs and people in close contact with them became clinically affected with an acute influenza-like illness, and virus was isolated from several pigs and at least two people (parent and child). The viruses isolated from the humans were 100% identical to the viruses isolated from the pigs, indicating that the virus was shared between pigs and people at the fair, again emphasizing the zoonotic risk for SIV.
B. Novel SI Isolates in North America A number of novel subtypes were isolated from swine in the past decade. Most of these novel SI subtypes were not able to establish themselves in the swine population. However, the following examples indicate that there is an ever-present chance of a new influenza subtype being established within the swine population which could have dire consequences for human health. The species barrier for the transmission of avian influenza viruses to pigs may be less stringent, since pigs contain receptors for both avian and mammalian influenza viruses in their respiratory tract (Ito et al., 1998). It is therefore, not surprising that pigs can be experimentally infected at least transiently with a wide variety of subtypes of avian influenza viruses (Kida et al., 1994). In addition, co-infection of pigs with a swine virus and with an avian virus unable to replicate in pigs generated reassortant viruses that could be passaged in pigs, indicating that even avian viruses that do not replicate in pigs can contribute genes to generate reassortant viruses (Kida et al., 1994).
1. The H4 Experience An example of infection of pigs with an avian influenza virus (AIV) occurred on a swine farm in Canada in October 1999. Genetic and antigenic analyses demonstrated that viruses isolated from pigs during an
138
Amy L. Vincent et al.
outbreak of respiratory disease were wholly avian H4N6 viruses of the North American lineage (Karasin et al., 2000). It was found that the farm of origin is located near a lake on which large numbers of waterfowl congregate each fall and from which the farm drew water. Therefore, the source of this virus was most likely ducks on the adjacent lake. It is well known that ducks shed high level of virus which can be isolated from unconcentrated lake water (Laver et al., 2000). The H4N6 virus spread to additional units of the original farm, suggesting that it has the ability to spread from pig-to-pig (Olsen, 2002). Fortunately, it has not been detected outside the original farm system. Interestingly, the HA of this virus contained amino acids in the receptor binding pocket that have been associated with mammalian receptor binding (Karasin et al., 2000).
2. The H2 Experience Unique H2N3 influenza viruses were recently isolated from clinically affected pigs from two farms in the central U.S. (Ma et al., 2007). Sequencing demonstrated they were H2N3 influenza A viruses with 99.3–99.9% homology between the isolates. The HA segment was similar to an AIV H2N3 isolated from mallards and the NA sequence was similar to an AIV H4N3 isolated from blue-winged teal. The PA segment had high homology to an AIV H6N5 isolated from mallards and the remaining genes were similar to influenza virus gene segments found in the contemporary TRIG cassette (human-like PB1, swine M, NP and NS, avian-like PB2) in U.S. SIVs. In addition to half of the gene segments being avian-like, the avian-like H2 HA has an amino acid sequence constellation in the receptor binding area indicating a preferential binding to the mammalian influenza receptor. This HA mutation is identical to the initial reassortant human influenza isolates found at the beginning of the 1957 H2N2 pandemic. In vivo studies in mice, swine and in ferrets, surrogate model for human influenza infection, were conducted. Experimentally-infected pigs developed lung lesions following challenge and virus was shed to contact control pigs that became infected and seroconverted. Similarly, in ferrets, virus was transmitted to contact ferrets. In addition, mortality was induced in young mice. The only recognized common thread between the two pig farms were geographic location and the use of pond water for both drinking and cleaning in the pig barns. The ability of the H2N3 viruses with avian origin surface glycoproteins to infect and replicate in three mammalian hosts without serial passage for adaptation in each species suggests this virus is already adapted to the mammalian host and may have potential risk to the human population. Although viruses of each of the 16 influenza A HA subtypes are potential human pathogens, only viruses of the H1, H2, and H3 subtype are known to have been successfully established in humans (Hilleman, 2002). H2 influenza viruses have been absent from human circulation since 1968. As such they pose a substantial human pandemic risk because of lack of population immunity.
SIV: A North American Perspective
139
IV. VACCINATION OF PIGS AGAINST SI Vaccinating pigs against influenza A virus has become a common practice in the U.S. swine industry over the last 10 years. Inactivated influenza vaccines became commercially available in 1994. In 1995, influenza vaccine usage was not reported in the National Animal Health Monitoring System survey of the U.S. swine operations (USDA, 1995). However, by 2000, over 40% of large producers reported that they vaccinated breeding females and approximately 20% vaccinated weaned pigs (USDA, 2003). In the survey conducted in 2006, the number of large producers vaccinating breeding females increased to 70%, whereas vaccinating weaned pigs remained relatively unchanged (USDA, 2007). Importantly, of those farms that vaccinated breeding females in 2006, approximately 20% reported using autogenous SIV vaccines rather than commercial vaccines. Autogenous vaccines prepared from virus cultures that have been inactivated may be used only in the herd of origin under the direction of a veterinarian. Autogenous vaccine usage against influenza virus has increased due to the diversity of viruses circulating in the North American pig population and the inability of the animal biologics industry to change the vaccine composition as rapidly as the viruses are changing. In contrast to human influenza virus epidemiology, SIV is no longer seasonal and there are too many circulating variants in North America to include a representative few in a bivalent or trivalent killed vaccine. There are three major problems with the control and prevention of SI in the U.S.: (a) SIV is changing faster than traditional vaccines can be developed, (b) There is a need for vaccines that can induce better crossprotection among SIV isolates, and (c) Passively acquired immunity is believed to block vaccine efficacy in pigs. The first line of defense against influenza virus infection is the innate immune system. Host cells have molecular sensors that recognize specific motifs from prokaryotic, protozoan, and viral pathogens. Some of the known sensors for single stranded RNA viruses like influenza viruses include the RNA helicases RIG-I and MDA-5 and the RNA binding and signaling proteins TLR3 and TLR 7 (reviewed in Garcia-Sastre, 2006). Many of these sensors have pathways that converge to upregulate the type 1 interferons (IFN a/b). IFN a/b, in turn, sound the alarm to other nearby cells and activate them through the production of cytokines and chemokines. In addition, the presence of type 1 interferons upregulates the production of host antiviral proteins such as Mx, PKR, and OAS, impairing or destroying the invading virus. Many pathogens, including influenza virus, have evolved to interfere with the IFN a/b signaling cascade as part of their survival mechanisms. The NS1 protein of influenza virus contributes to virulence by interacting with the IFN a/b
140
Amy L. Vincent et al.
antiviral response. The carboxy-terminus of NS1 is reported to contain the effector domain responsible for antagonizing the type 1 IFN pathway (Wang et al., 2002). The amino terminus is reported to contain the RNA binding domain (Wang et al., 2000), which may allow the NS1 protein to sequester viral RNA and therefore avoid detection by the host cell’s virus sensor. Using reverse genetics approaches, we have produced H3N2 SIVs with deletions in the 30 end of the NS1 gene; The NS1-truncated mutants are highly attenuated in vitro and in pigs (Solorzano et al., 2005), demonstrating that the NS1 is a virulence factor of SIV in pigs. The attenuation is due, at least in a major part, to a loss in the ability of the mutated virus to block antiviral defense mechanisms, with subsequent host cell upregulation of type 1 interferons (Solorzano et al., 2005) and downstream effector molecules, such as Mx and PKR (Wang et al., 2002). Protective immunity against infection with influenza involves both the humoral and cell mediated (CMI) arms of the adaptive immune system. The responses of the humoral and CMI systems are interwoven and both are necessary for protective immunity. Antibodies play a significant role in attenuating and preventing swine influenza as shown by the protective capacity of colostrum (Renshaw, 1975) and inactivated vaccines (Bikour et al., 1996). Clinical protection against challenge virus appears to be directly correlated with the hemagglutination inhibition (HI) titer in the serum of an individual animal, that is, a high HI titer provides better protection against challenge than a low HI titer. This information has led to the suggestion that the presence and magnitude of an HI titer could be a predictor of protection. Unfortunately, this seems only true when the priming HA antigen inducing the HI titer is antigenically closely related to the HA of the challenge virus. Other studies have demonstrated the protective qualities of antibodies at the mucosal level. Pigs immunized with virulent, live SIV, and then challenged with the same virus 42 days later did have a detectable anamnestic antibody response at the mucosal level but not in the serum (Larsen et al., 2000). Specifically, a rise in IgA and IgG was detected in the nasal cavity, the site of challenge. This data supports the hypothesis that antibody mediated immune reactions at the mucosal level and not the systemic level are important for protecting the respiratory tract from SIV. Extensive studies investigating the immune response of mice to influenza virus infection indicate they can develop homosubtypic (same subtype) and heterosubtypic (different subtype) immunity (Het-I). Homotypic immunity tends to exert a more complete protection, whereas Het-I may fail to prevent an initial infection, but is successful in reduction of virus shedding and a more rapid recovery from infection, (reviewed in Tamura et al., 2005). Collectively, studies on hetero- and homosubtypic immunity in mice demonstrate that virus elimination and protection from disease are dependent on virus-specific neutralizing antibodies and T
SIV: A North American Perspective
141
cells, as well as the virus-specific mucosal immune response. In mice, cross-reactive IgA induced by natural infection was shown to be strongly correlated to protection from challenge with a homosubtypic virus belonging to a different, heterologous genotype (Liew et al., 1984). Serum HI antibody titer and the presence of cross-reactive cytotoxic T-lymphocytes did not correlate with protection, but might be crucial for recovery. IgA was shown to be more cross-reactive than IgG against heterologous influenza viruses and passive transfer of IgA to non-immune mice conferred protection (Tamura et al., 1991). Although cytotoxic T-lymphocyte activity has been shown to be stimulated in heterosubtypic primed mice (Nguyen et al., 1999), protection against heterosubtypic challenge in mice was largely dependent on the presence of B-cells and CD4þ T-helper cells, specifically those with a Th1 phenotype (Moran et al., 1999; Nguyen et al., 1999, 2001). CD4þ T cells primed against conserved internal influenza proteins may be responsible for the rapid development of cross-reacting antibodies following a heterosubtypic challenge (Scherle and Gerhard, 1986). These cross-reacting antibodies appeared to provide at least partial protection and a more rapid recovery after heterosubtypic challenge. In contrast, an inactivated virus challenge in mice stimulated a Th2 response and no heterosubtypic immunity (Moran et al., 1999). However, heterosubtypic immunity could be induced when mice were immunized with inactivated virus and, in addition, received an injection of interleukin (IL) 12 and antibodies against IL 4 (Moran et al., 1999). Although impractical for swine vaccination, these results suggest that improved adjuvants may enhance the protective immunity of killed vaccines. The continual consolidation of the swine industry into larger swine herds housed in swine-dense regions and the emergence of novel SIV subtypes ensures that future SIV control will be heavily dependent upon vaccination protocols. When swine are infected with a virulent influenza virus, complete protective immunity typically develops against rechallenge with the homologous virus, that is, there is little or no detectable virus replication following secondary challenge and there are no lung lesions associated with challenge (Larsen et al., 2000). Exposure to live H1N1 and H3N2 viruses also conferred complete protection against an H1N2 with an unrelated HA protein (Van Reeth et al., 2003), however vaccination with commercial killed vaccines containing H1N1 and H3N2 did not protect against H1N2 challenge (Reeth et al., 2004). In studies using inactivated whole virus vaccines only partial protection was found following homologous challenge (Bikour et al., 1996; Macklin et al., 1998). These studies indicate that inactivated vaccines have limited ability to cross-protect against heterologous homosubtypic, or heterosubtypic viruses. Good protection can only be achieved when challenge and vaccine strain show cross-reactivity. The development of attenuated modified live-virus vaccine (MLV) or vector-based subunit vaccines for swine
142
Amy L. Vincent et al.
that induce an immune response based on both humoral and cell mediated mechanisms are likely to improve homosubtypic and heterosubtypic protection. To evaluate the clinical relevance of in vitro serum cross-reactivity, we studied two H1 isolates, IA30 (H1N1) and MN03 (H1N2), with substantial genetic divergence in the HA gene and failure to cross-react in the HI assay (Vincent et al., 2008) in more detail. Inactivated vaccines were prepared from both isolates and used to immunize two groups of conventional pigs. In addition, two groups of pigs were primed with live, virulent virus. The vaccinated pigs (either live or inactivated vaccine) were then challenged with the homologous and heterologous viruses. Both inactivated vaccines provided excellent protection against homologous challenge. However, the inactivated IA30 vaccine failed to protect against the heterologous MN03 challenge, whereas the MN03 vaccine was partially protective against the heterologous IA30 challenge. Surprisingly, 3 of the 9 pigs in the MN03-challenged, IA30-immunized group had substantially greater percentages of lung lesions compared to non-vaccinated MN03 challenge controls. This suggests that the IA30 inactivated vaccine may have potentiated the level of pneumonia when challenged with the heterologous MN03 virus. This was not true when MN03 vaccinated pigs were challenged with the IA30 virus. The potentiation of lung lesions may have been immune-mediated due to the induction of lower levels of IgA in conjunction with higher levels of IgG antibodies in the lungs of the three IA30-immunized pigs. The inactivated and live vaccines induced an isolate-specific serum HI response against homologous virus, but there was no cross-reactivity with heterologous viruses. We concluded from this study that divergent H1 viruses that do not cross-react serologically may not provide complete cross-protection when used as an inactivated vaccine. Although mild lung lesions consistent with SIV were seen in pigs primed with live IA30 or MN03 and challenged with MN03 or IA30, respectively, the live vaccination prevented virus shedding from the nose and no virus was isolated from the lungs in our experimental pig model (Vincent et al., 2008). In summary, these results suggest that the use of live virus or a mucosal route for immunization may enhance the efficacy of vaccines and prevent virus shedding when used in the face of antigenically heterologous viruses of the same subtype. Reverse genetics or the de novo synthesis of negative sense RNA viruses from cloned cDNA, has become a reliable laboratory method that provides a powerful tool for studying various aspects of the viral life cycle, the role of viral proteins in pathogenicity and the interplay of viral proteins with components of the host’s immune system. It also opens the way to develop live attenuated virus vaccines and vaccine vectors. A reverse genetics system that allows the generation of influenza A viruses entirely from cloned cDNAs has been established (Fodor et al.,
SIV: A North American Perspective
143
1999; Hoffmann et al., 2000; Neumann et al., 1999). This technology allowed the generation of mouse-adapted viruses with mutations in the NS1 gene which exhibited an attenuated phenotype in cell culture, mice, and embryonated eggs (Talon et al., 2000). The attenuation in these models was believed to be due to a loss of function of the viral NS1 protein, a type 1 interferon antagonist. We have investigated the role of the NS1 protein in the virulence of a SIV isolate in the natural host, the pig, producing various mutants encoding carboxy-truncated NS1 proteins. Similar to the other model systems, we found that these NS1 truncations decreased the ability of SIVs to prevent IFN-a/b synthesis in pig cells and conveyed attenuation in pigs (Solorzano et al., 2005). We proposed NS1mutated SIVs might have a great potential as live attenuated vaccine candidates against SIV infections of pigs (Solorzano et al., 2005). The development of attenuated MLV or vectored subunit vaccines for swine that induce a balanced immune response including humoral and cell mediated mechanisms are likely to improve homosubtypic and heterosubtypic protection. A cold-adapted live attenuated intranasal (IN) influenza vaccine has been approved in the U.S. for use in humans with results from clinical and field trials showing good efficacy (Belshe, 2004). A similar vaccine is available for horses as well (Townsend et al., 2001). A prototype H3N2 SIV (Sw/A/TX/98) virus with a carboxy-terminal truncation of the NS1 gene starting at amino acid 126 (D126) generated by reverse genetics has been shown to be highly attenuated in pigs, was not shed from the nose but was capable of stimulating an immune response (Solorzano et al., 2005). The potential of this NS1 mutant, called TX98 NS1D126, for use as a MLV vaccine in pigs has been recently evaluated. To evaluate the TX98 NS1D126 as an MLV vaccine, 4-week-old pigs were vaccinated and boosted with the TX98 NS1D126 MLV via the intratracheal route (Richt et al., 2006). Pigs were challenged with wild type homologous H3N2 or heterosubtypic classical H1N1 SIVs and necropsied 5 days later. The MLV was highly attenuated and completely protected against challenge with the homologous virus. Vaccinated pigs challenged with the heterosubtypic cH1N1 virus demonstrated pathologic lung changes similar to the nonvaccinated H1N1 control pigs. However, vaccinated pigs challenged with cH1N1 had significantly reduced virus shedding from the respiratory tract when compared to nonvaccinated, cH1N1 challenged pigs. All vaccinated pigs developed a significant level of HI titer, serum IgG, and mucosal IgG and IgA antibodies against parental H3N2 SIV antigens (Richt et al., 2006). A separate study evaluated the efficacy of the TX98 NS1D126 MLV when used via the IN or intramuscular (IM) route and challenged with homologous virus (Fig. 4). Furthermore, pigs vaccinated via the IN route were also challenged with a homosubtypic, but genetically and
144
Amy L. Vincent et al.
Challenge Control of isolate disease Hom (1 X MLV) Hom (2 X MLV)
Mucosal lg response
Control of replication
Control of pneumonia
+++
++
+++
−
++++
++++
+++
++
Hom
++++
++++
++++
Mucosal lg response ++++
Hom-het
+++
+++
++++
+++
Het
+
+
−
+
Challenge isolate
Control of disease
Control of Control of replication pneumonia
IM
IN
FIGURE 4 Influence of route and dose on the efficacy of the TX98 NS1D126 MLV against SIV. Challenge isolate: Hom ¼ homologous wild type H3N2; Hom-het ¼ homosubtypic heterologous H3N2; and Het ¼ heterosubtypic H1N1. Scale of 0–4þ, with the greater number of þ being most protective. Control of disease is based on clinical signs and rectal temperatures. Control of replication is based on virus titers from nasal swabs and bronchioalveolar lavage fluid. Control of pneumonia is based on percentage of macroscopic pneumonia and microscopic evaluation. Mucosal immunoglobulin (Ig) response is based on levels of IgG and IgA in the lung. IM ¼ intramuscular route; vaccination results are summarized in the black box with one dose (1) or two doses (2). IN ¼ intranasal route; vaccination results are summarized in the gray box.
antigenically heterologous H3N2 (CO99) and a rH1N1 (IA04) SIV that contained the TRIG cassette similar to the triple reassortant H3N2 viruses (Vincent et al., 2007). A single dose of MLV administered intranasally conferred complete protection against homologous virus and nearly complete protection against the heterologous H3N2 CO99 virus challenge (Fig. 4). When challenged with the rH1N1 IA04 virus, MLV vaccinated animals displayed reduced fever and virus titers despite minimal reduction in lung lesions (Fig. 4). In vaccinated pigs, there was no serologic cross-reactivity by HI assays with the heterologous or heterosubtypic viruses. However, there appeared to be substantial cross-reactivity in antibodies at the mucosal level with the CO99 virus in MLV vaccinated pigs (Vincent et al., 2007). It is apparent that a complex host response involving CMI and humoral mechanisms contribute to the immunity established via the TX98 NS1D126 MLV SIV vaccine and the immune response to MLV seems to be superior to that induced by inactivated influenza vaccines. One of the primary reasons for vaccinating breeding sows with inactivated vaccines is to stimulate passive antibody transfer to the suckling pig. The level of protection is dependent on the level of maternal derived antibody (MDA). However, several studies have demonstrated that
SIV: A North American Perspective
145
MDA rarely prevents infection with influenza virus and only provides partial protection (Loeffen et al., 2003; Renshaw, 1975). In addition, the presence of MDA interferes with a primary immune response to SIV, either by infection or vaccination (Kitikoon et al., 2006; Loeffen et al., 2003; Renshaw, 1975). We recently evaluated the ability of IN applied TX98 NS1D126 MLV to overcome maternal antibody interference when challenged with homologous (TX98 wild type) or heterologous (CO99) H3N2 virus. The MDA present in the vaccinated pigs was shown to interfere with the serologic SIV-antibody response to either an inactivated TX98 vaccine or the TX98 NS1D126 MLV; however, protection from challenge with homologous virus was demonstrated for both vaccines (Vincent, unpublished results). MDA reduced the efficacy of one-dose IN application of the MLV when compared to pigs vaccinated in the absence of MDA, although the virus levels in the respiratory tract were significantly reduced compared to nonvaccinated controls. The most remarkable finding from this study was the observation of a dramatic enhancement of disease and pneumonia in pigs which were vaccinated with an inactivated TX98-based vaccine in the presence of MDA, followed by challenge with heterologous CO99 virus. This was not seen in MDA positive pigs vaccinated with the MLV nor in MDA negative pigs given inactivated or MLV vaccine and challenged with heterologous H3N2 CO99 virus. Enhancement of pneumonia by inactivated vaccine used in the face of MDA with an H1N1 challenge was reported previously by Kitikoon et al. (2006), which supports our findings with H3N2 viruses. These results indicate a much more insidious role for MDA rather than simple interference with primary immune responses when using inactivated vaccines in young pigs. Recombinant human adenoviruses have been demonstrated to be effective vectors for insertion of antigens from infectious agents for use as vaccine candidates in many species, including those of veterinary importance (Casimiro et al., 2003; Elahi et al., 1999; Eloit et al., 1990; Mayr et al., 2001; Pacheco et al., 2005). Several of these vaccine candidates, specifically those created from human adenovirus serotype 5 (HAd5), have been shown to provide excellent protection from challenge with foot-and-mouth disease virus and SIV (Mayr et al., 2001; Wesley et al., 2004). Vaccination with HAd vectors has been shown to induce both humoral and cell-mediated immunity, making them potentially more effective than inactivated vaccines and more similar to the response elicited from MLV, reviewed in Gamvrellis et al. (2004). In addition, HAd vectored vaccines given by a mucosal route have been shown to provide superior, long lasting mucosal immunity (Baca-Estrada et al., 1995). Replication-defective adenovirus recombinants were developed as potential vaccines against H3N2 influenza viruses (Wesley et al., 2004). Pigs in the groups given the recombinant adenovirus expressing
146
Amy L. Vincent et al.
HA protein developed high levels of virus-specific HI antibody by 4 weeks postvaccination. Pigs in the group vaccinated with recombinant viruses expressing both the HA and NP in a mixture were completely protected against homologous challenge, shown by the lack of nasal shedding of virus following challenge and by the lack of lung lesions at 1 week following the challenge infection. In addition, the efficacy of the HAd5 vaccine for protecting weaned pigs against SIV subtype H3N2 infection were evaluated when administered via two injection methods, either with a needle-free injection device or by traditional IM injection (Wesley and Lager, 2005). Traditional IM-administered vaccination induced consistently higher HI responses than vaccination via needlefree injection, but the differences were not significant. Likewise, traditional IM administration was superior at reducing nasal virus shedding except at the highest dose, at which both methods blocked virus replication. The severity of lung lesions was reduced in a dose-dependent manner by both vaccination methods. The replication-defective vaccine HAd5 virus was not transmitted to sentinel pigs (Wesley and Lager, 2005). In addition to the success in naı¨ve pigs, recombinant HAd5 vectored SIV vaccines were demonstrated to prime the immune system in the presence of MDA. Piglets with H3N2-specific MDA were sham-inoculated with a nonexpressing HAd5 vector or given a primary vaccination with replication-defective HAd5 expressing the HA and the NP of an H3N2 SIV subtype virus (Wesley and Lager, 2006). The HI titer of the shaminoculated group showed continued antibody decay whereas piglets vaccinated with HAd5-SIV developed an active immune response by the second week postvaccination. At 4-weeks of age, when the HI titer of the sham-inoculated group had decayed, the sham-inoculated group and half of the HAd5 SIV vaccinated pigs were boosted with a commercial inactivated SIV vaccine. The boosted pigs that had been primed with the HAd5 expressing SIV genes in the presence of MDA had a strong anamnestic response while sham-inoculated pigs did not respond to the commercial vaccine. Two weeks after the booster vaccination the pigs were challenged with a heterologous virulent H3N2 SIV. The pigs primed with the HAd5-SIV vaccine and boosted with inactivated vaccine showed a reduction of clinical signs, reduced virus levels in the respiratory tract, and the absence lung lesions (Wesley and Lager, 2006). In contrast, MDA positive pigs not primed with the HAd5-SIV vaccine and only vaccinated with the inactivated vaccine demonstrated a vaccine failure (Wesley and Lager, 2006). It is evident from the increasing number of novel subtypes and genetic variants isolated from pigs that controlling swine flu will only continue to be difficult. New strategies of vaccine development must be considered to keep up with the ever-evolving influenza viruses and to overcome the problem of maternal antibody interference with inactivated vaccines.
SIV: A North American Perspective
147
The demonstrated safety and efficacy of cold-adapted modified live virus vaccines in human and equine medicine has paved the way for investigating modified live vaccines in swine medicine. The few studies using different SIV vaccination concepts in pigs have shown that strain, route of administration, and use of vaccine additives can play a role in enhancing heterologous protection. Future studies are needed to address each of these areas. The use of reverse genetics to genetically engineer viruses with vaccine potential (live or inactivated) and to identify virulence genes will certainly help in this pursuit as will vectored vaccines. In order to gain a better understanding of homosubtypic and heterosubtypic vaccine efficacy, the CMI and humoral immune responses at the systemic and mucosal levels need to be included in future pig studies.
V. CONCLUSIONS AND OUTLOOK The impact of influenza A in humans and animals, whether measured by morbidity, mortality, or economic losses, is significant. It is, therefore, essential to understand the mechanisms that allow these viruses to jump species barriers and establish themselves in new animal populations. The emergence of new subtypes of SIVs (hu-H1, H3N2, H4N6, and H2N3) in North American pigs has implications not only for pigs but also for the people who care for them. These newly emerging viruses are capable of epidemics at the herd or U.S. swine industry level since they are antigenically distinct from previously circulating and/or currently used commercial vaccine strains, are virulent in the pig and can infect and transmit from pig to pig. The potential for human infection as North American SIVs continue to drift, shift, and adapt to a mammalian host is unclear, but definitely remains a risk. It is increasingly evident that improved vaccination strategies with novel vaccine platforms are needed. Therefore, novel vaccine approaches using genetically engineered MLV and vectored vaccines are discussed in this review. The influenza epidemiology in North American pigs clearly indicates that the potential for pandemic influenza virus emergence exists not only in the traditionally considered ‘‘influenza hotbeds’’ of Southeast Asia (Shortridge and Stuart-Harris, 1982) but also in North America. This review underscores the need for vigilance in examining influenza A viruses from swine (and other species) for human pathogenic potential in addition to the major focus currently placed on AIVs. Although there does not appear to be a simple solution to the SIV problem in North America, some lessons can be learned from our and others experiences over the last two decades. People to pig, pig to people, pig to avian, avian to human, and avian to pig transmissions occur. Strategies that may reduce the risk for these transmission events should be employed in the swine industry, such as vaccination of workers
148
Amy L. Vincent et al.
with occupational exposure to swine (Gray et al., 2007); sick policies for workers, and the use of disposable respirators; bird-proofing swine facilities; and using only treated water for barn cleaning and consumption. The swine industry should be aware that the use of untreated pond or lake water can be a threat to animal health. Pig to people transmission must be emphasized as well. Education and caution for workers with occupational exposure as well as for those in the human health care system is critical for reducing and/or monitoring these transmission events. Our strength in monitoring and reacting to newly emerging or re-emerging subtypes in the swine or human population will be much greater as the veterinary and public health communities collaborate and engage together in these efforts.
ACKNOWLEDGMENTS The authors thank the team of technical and animal care staff that has significantly contributed to the many studies described in this review. We thank Mike Marti for illustrations and Drs A. Garcia-Sastre, M. R. Gramer, and R.J. Webby for their continued support. Projects described in this review were funded by the USDA-Agricultural Research Service and in part with Federal funds from the National Institute Allergy and Infectious Diseases, National Institutes of Health, Department of Health and Human Services, under Contract No. HHSN266200700005C and from the Center for Disease Control and Prevention, grant No. U01 CI000357-01.
REFERENCES Alexander, D. J., and Brown, I. H. (2000). Recent zoonoses caused by influenza A viruses. Rev. Sci. Tech. 19(1):197–225. Baca-Estrada, M. E., Liang, X., Babiuk, L. A., and Yoo, D. (1995). Induction of mucosal immunity in cotton rats to haemagglutinin-esterase glycoprotein of bovine coronavirus by recombinant adenovirus. Immunology 86(1):134–140. Belshe, R. B. (2004). Current status of live attenuated influenza virus vaccine in the US. Virus Res. 103(1–2):177–185. Bikour, M. H., Cornaglia, E., and Elazhary, Y. (1996). Evaluation of a protective immunity induced by an inactivated influenza H3N2 vaccine after an intratracheal challenge of pigs. Can. J. Vet. Res. 60(4):312–314. Brown, E. G. (2000). Influenza virus genetics. Biomed. Pharmacother. 54(4):196–209. Buonagurio, D. A., Nakada, S., Parvin, J. D., Krystal, M., Palese, P., and Fitch, W. M. (1986). Evolution of human influenza A viruses over 50 years: Rapid, uniform rate of change in NS gene. Science 232(4753):980–982. Casimiro, D. R., Tang, A., Chen, L., Fu, T. M., Evans, R. K., Davies, M. E., Freed, D. C., Hurni, W., Aste-Amezaga, J. M., Guan, L., Long, R., Huang, L., et al. (2003). Vaccineinduced immunity in baboons by using DNA and replication-incompetent adenovirus type 5 vectors expressing a human immunodeficiency virus type 1 gag gene. J. Virol. 77 (13):7663–7668. Chambers, T. M., Hinshaw, V. S., Kawaoka, Y., Easterday, B. C., and Webster, R. G. (1991). Influenza viral infection of swine in the United States 1988–1989. Arch. Virol. 116 (1–4):261–265.
SIV: A North American Perspective
149
Choi, Y. K., Goyal, S. M., Farnham, M. W., and Joo, H. S. (2002). Phylogenetic analysis of H1N2 isolates of influenza A virus from pigs in the United States. Virus Res. 87 (2):173–179. Compans, R. W., Content, J., and Duesberg, P. H. (1972). Structure of the ribonucleoprotein of influenza virus. J. Virol. 10(4):795–800. Conenello, G. M., and Palese, P. (2007). Influenza A virus PB1-F2: A small protein with a big punch. Cell Host Microbe 2(4):207–209. Crawford, P. C., Dubovi, E. J., Castleman, W. L., Stephenson, I., Gibbs, E. P., Chen, L., Smith, C., Hill, R. C., Ferro, P., Pompey, J., Bright, R. A., Medina, M. J., et al. (2005). Transmission of equine influenza virus to dogs. Science 310(5747):482–485. Easterday, B. C., and van Reeth, K. (1999). Swine Influenza. In ‘‘Diseases of Swine’’ (B. E. Straw, S. D’Allaire, W. L. Mengeling, and D. J. Taylor, Eds.), 8th Ed. pp. 277–290. Iowa State University Press, Ames. Elahi, S. M., Shen, S. H., Talbot, B. G., Massie, B., Harpin, S., and Elazhary, Y. (1999). Induction of humoral and cellular immune responses against the nucleocapsid of bovine viral diarrhea virus by an adenovirus vector with an inducible promoter. Virology 261(1):1–7. Ellis, J., Clark, E., Haines, D., West, K., Krakowka, S., Kennedy, S., and Allan, G. M. (2004). Porcine circovirus-2 and concurrent infections in the field. Vet. Microbiol. 98(2):159–163. Eloit, M., Gilardi-Hebenstreit, P., Toma, B., and Perricaudet, M. (1990). Construction of a defective adenovirus vector expressing the pseudorabies virus glycoprotein gp50 and its use as a live vaccine. J. Gen. Virol. 71(Pt. 10):2425–2431. Fodor, E., Devenish, L., Engelhardt, O. G., Palese, P., Brownlee, G. G., and Garcia-Sastre, A. (1999). Rescue of influenza A virus from recombinant DNA. J. Virol. 73(11):9679–9682. Fouchier, R. A., Munster, V., Wallensten, A., Bestebroer, T. M., Herfst, S., Smith, D., Rimmelzwaan, G. F., Olsen, B., and Osterhaus, A. D. (2005). Characterization of a novel influenza A virus hemagglutinin subtype (H16) obtained from black-headed gulls. J. Virol. 79(5):2814–2822. Gamvrellis, A., Leong, D., Hanley, J. C., Xiang, S. D., Mottram, P., and Plebanski, M. (2004). Vaccines that facilitate antigen entry into dendritic cells. Immunol. Cell. Biol. 82(5):506–516. Garcia-Sastre, A. (2006). Antiviral response in pandemic influenza viruses. Emerg. Infect. Dis. 12(1):44–47. Gaydos, J. C., Hodder, R. A., Top, F. H., Jr., Soden, V. J., Allen, R. G., Bartley, J. D., Zabkar, J. H., Nowosiwsky, T., and Russell, P. K. (1977). Swine influenza A at Fort Dix, New Jersey (January–February 1976). I. Case finding and clinical study of cases. J. Infect. Dis. 136(Suppl.):S356–S362. Gottschalk, A. (1957). Neuraminidase: The specific enzyme of influenza virus and Vibrio cholerae. Biochim. Biophys. Acta 23(3):645–646. Gramer, M. R. (2007). 38th Annual Meeting of the American Association of Swine Veterinarians, Orlando, FL. Gray, G. C., Trampel, D. W., and Roth, J. A. (2007). Pandemic influenza planning: Shouldn’t swine and poultry workers be included? Vaccine 25(22):4376–4381. Hilleman, M. R. (2002). Realities and enigmas of human viral influenza: Pathogenesis, epidemiology and control. Vaccine 20(25–26):3068–3087. Hoffmann, E., Neumann, G., Kawaoka, Y., Hobom, G., and Webster, R. G. (2000). A DNA transfection system for generation of influenza A virus from eight plasmids. Proc. Natl. Acad. Sci. USA 97(11):6108–6113. Horimoto, T., and Kawaoka, Y. (2001). Pandemic threat posed by avian influenza A viruses. Clin. Microbiol. Rev. 14(1):129–149. Ito, T. (2000). Interspecies transmission and receptor recognition of influenza A viruses. Microbiol. Immunol. 44(6):423–430. Ito, T., Couceiro, J. N., Kelm, S., Baum, L. G., Krauss, S., Castrucci, M. R., Donatelli, I., Kida, H., Paulson, J. C., Webster, R. G., and Kawaoka, Y. (1998). Molecular basis for the
150
Amy L. Vincent et al.
generation in pigs of influenza A viruses with pandemic potential. J. Virol. 72 (9):7367–7373. Karasin, A. I., Brown, I. H., Carman, S., and Olsen, C. W. (2000). Isolation and characterization of H4N6 avian influenza viruses from pigs with pneumonia in Canada. J. Virol. 74 (19):9322–9327. Karasin, A. I., Landgraf, J., Swenson, S., Erickson, G., Goyal, S., Woodruff, M., Scherba, G., Anderson, G., and Olsen, C. W. (2002). Genetic characterization of H1N2 influenza A viruses isolated from pigs throughout the United States. J. Clin. Microbiol. 40 (3):1073–1079. Karasin, A. I., Carman, S., and Olsen, C. W. (2006). Identification of human H1N2 and human-swine reassortant H1N2 and H1N1 influenza A viruses among pigs in Ontario, Canada (2003 to 2005). J. Clin. Microbiol. 44(3):1123–1126. Kida, H., Ito, T., Yasuda, J., Shimizu, Y., Itakura, C., Shortridge, K. F., Kawaoka, Y., and Webster, R. G. (1994). Potential for transmission of avian influenza viruses to pigs. J. Gen. Virol. 75(Pt. 9):2183–2188. Kitikoon, P., Nilubol, D., Erickson, B. J., Janke, B. H., Hoover, T., Sornsen, S., and Thacker, E. L. (2006). The immune response and maternal antibody interference to a heterologous H1N1 swine influenza virus infection following vaccination. Vet. Immunol. Immunopathol. 112(3–4):117–128. Klopfleisch, R., Wolf, P. U., Wolf, C., Harder, T., Starick, E., Niebuhr, M., Mettenleiter, T. C., and Teifke, J. P. (2007). Encephalitis in a stone marten (Martes foina) after natural infection with highly pathogenic avian influenza virus subtype H5N1. J. Comp. Pathol. 137 (2–3):155–159. Klumpp, K., Ruigrok, R. W., and Baudin, F. (1997). Roles of the influenza virus polymerase and nucleoprotein in forming a functional RNP structure. EMBO J. 16(6):1248–1257. Knipe, D., Howley, P. M., Griffin, D. E., Lamb, R. A., and Martin, M. A., Eds. (2007). Fields Virology. 5th Ed. Lippincott Williams & Wilkins. Koen, J. S. (1919). A practical method for field diagnosis of swine diseases. Am. J. Vet. Med. 14:468–470. Larsen, D. L., Karasin, A., Zuckermann, F., and Olsen, C. W. (2000). Systemic and mucosal immune responses to H1N1 influenza virus infection in pigs. Vet. Microbiol. 74 (1–2):117–131. Laver, W. G., Bischofberger, N., and Webster, R. G. (2000). The origin and control of pandemic influenza. Perspect. Biol. Med. 43(2):173–192. Lekcharoensuk, P., Lager, K. M., Vemulapalli, R., Woodruff, M., Vincent, A. L., and Richt, J. A. (2006). Novel swine influenza virus subtype H3N1, United States. Emerg. Infect. Dis. 12(5):787–794. Liew, F. Y., Russell, S. M., Appleyard, G., Brand, C. M., and Beale, J. (1984). Cross-protection in mice infected with influenza A virus by the respiratory route is correlated with local IgA antibody rather than serum antibody or cytotoxic T cell reactivity. Eur. J. Immunol. 14 (4):350–356. Loeffen, W. L., Heinen, P. P., Bianchi, A. T., Hunneman, W. A., and Verheijden, J. H. (2003). Effect of maternally derived antibodies on the clinical signs and immune response in pigs after primary and secondary infection with an influenza H1N1 virus. Vet. Immunol. Immunopathol. 92(1–2):23–35. Luoh, S. M., McGregor, M. W., and Hinshaw, V. S. (1992). Hemagglutinin mutations related to antigenic variation in H1 swine influenza viruses. J. Virol. 66(2):1066–1073. Ma, W., Gramer, M., Rossow, K., and Yoon, K. J. (2006). Isolation and genetic characterization of new reassortant H3N1 swine influenza virus from pigs in the midwestern United States. J. Virol. 80(10):5092–5096. Ma, W., Vincent, A. L., Gramer, M. R., Brockwell, C. B., Lager, K. M., Janke, B. H., Gauger, P. C., Patnayak, D. P., Webby, R. J., and Richt, J. A. (2007). Identification of
SIV: A North American Perspective
151
H2N3 influenza A viruses from swine in the United States. Proc. Natl. Acad. Sci. USA 104 (52):20949–20954. Macklin, M. D., McCabe, D., McGregor, M. W., Neumann, V., Meyer, T., Callan, R., Hinshaw, V. S., and Swain, W. F. (1998). Immunization of pigs with a particle-mediated DNA vaccine to influenza A virus protects against challenge with homologous virus. J. Virol. 72(2):1491–1496. Matrosovich, M. N., Matrosovich, T. Y., Gray, T., Roberts, N. A., and Klenk, H. D. (2004). Neuraminidase is important for the initiation of influenza virus infection in human airway epithelium. J. Virol. 78(22):12665–12667. Mayr, G. A., O’Donnell, V., Chinsangaram, J., Mason, P. W., and Grubman, M. J. (2001). Immune responses and protection against foot-and-mouth disease virus (FMDV) challenge in swine vaccinated with adenovirus-FMDV constructs. Vaccine 19(15– 16):2152–2162. McQueen, J. L., Steele, J. H., and Robinson, R. Q. (1968). Influenza in animals. Adv. Vet. Sci. 12:285–336. Moran, T. M., Park, H., Fernandez-Sesma, A., and Schulman, J. L. (1999). Th2 responses to inactivated influenza virus can be converted to Th1 responses and facilitate recovery from heterosubtypic virus infection. J. Infect. Dis. 180(3):579–585. Myers, K. P., Olsen, C. W., Setterquist, S. F., Capuano, A. W., Donham, K. J., Thacker, E. L., Merchant, J. A., and Gray, G. C. (2006). Are swine workers in the United States at increased risk of infection with zoonotic influenza virus? Clin. Infect. Dis. 42(1):14–20. Myers, K. P., Olsen, C. W., and Gray, G. C. (2007). Cases of swine influenza in humans: A review of the literature. Clin. Infect. Dis. 44(8):1084–1088. Neumann, G., and Kawaoka, Y. (2006). Host range restriction and pathogenicity in the context of influenza pandemic. Emerg. Infect. Dis. 12(6):881–886. Neumann, G., Watanabe, T., Ito, H., Watanabe, S., Goto, H., Gao, P., Hughes, M., Perez, D. R., Donis, R., Hoffmann, E., Hobom, G., and Kawaoka, Y. (1999). Generation of influenza A viruses entirely from cloned cDNAs. Proc. Natl. Acad. Sci. USA 96(16):9345–9350. Nguyen, H. H., Moldoveanu, Z., Novak, M. J., van Ginkel, F. W., Ban, E., Kiyono, H., McGhee, J. R., and Mestecky, J. (1999). Heterosubtypic immunity to lethal influenza A virus infection is associated with virus-specific CD8(þ) cytotoxic T lymphocyte responses induced in mucosa-associated tissues. Virology 254(1):50–60. Nguyen, H. H., van Ginkel, F. W., Vu, H. L., McGhee, J. R., and Mestecky, J. (2001). Heterosubtypic immunity to influenza A virus infection requires B cells but not CD8þ cytotoxic T lymphocytes. J. Infect. Dis. 183(3):368–376. Noble, S., McGregor, M. S., Wentworth, D. E., and Hinshaw, V. S. (1993). Antigenic and genetic conservation of the haemagglutinin in H1N1 swine influenza viruses. J. Gen. Virol. 74(Pt. 6):1197–1200. Ohwada, K., Kitame, F., Sugawara, K., Nishimura, H., Homma, M., and Nakamura, K. (1987). Distribution of the antibody to influenza C virus in dogs and pigs in Yamagata Prefecture, Japan. Microbiol. Immunol. 31(12):1173–1180. Olsen, C. W. (2002). The emergence of novel swine influenza viruses in North America. Virus Res. 85(2):199–210. Olsen, C. W., McGregor, M. W., Cooley, A. J., Schantz, B., Hotze, B., and Hinshaw, V. S. (1993). Antigenic and genetic analysis of a recently isolated H1N1 swine influenza virus. Am. J. Vet. Res. 54(10):1630–1636. Olsen, C. W., Karasin, A. I., Carman, S., Li, Y., Bastien, N., Ojkic, D., Alves, D., Charbonneau, G., Henning, B. M., Low, D. E., Burton, L., and Broukhanski, G. (2006). Triple reassortant H3N2 influenza A viruses, Canada, 2005. Emerg. Infect. Dis. 12 (7):1132–1135. Pacheco, J. M., Brum, M. C., Moraes, M. P., Golde, W. T., and Grubman, M. J. (2005). Rapid protection of cattle from direct challenge with foot-and-mouth disease virus (FMDV) by a
152
Amy L. Vincent et al.
single inoculation with an adenovirus-vectored FMDV subunit vaccine. Virology 337 (2):205–209. Palese, P., Schulman, J. L., Bodo, G., and Meindl, P. (1974). Inhibition of influenza and parainfluenza virus replication in tissue culture by 2-deoxy-2,3-dehydro-N-trifluoroacetylneuraminic acid (FANA). Virology 59(2):490–498. Palese, P., Muster, T., Zheng, H., O’Neill, R., and Garcia-Sastre, A. (1999). Learning from our foes: A novel vaccine concept for influenza virus. Arch. Virol. Suppl. 15:131–138. Reeth, K. V., Brown, I., Essen, S., and Pensaert, M. (2004). Genetic relationships, serological cross-reaction and cross-protection between H1N2 and other influenza A virus subtypes endemic in European pigs. Virus Res. 103(1–2):115–124. Renshaw, H. W. (1975). Influence of antibody-mediated immune suppression on clinical, viral, and immune responses to swine influenza infection. Am. J. Vet. Res. 36(1):5–13. Richardson, J. C., and Akkina, R. K. (1991). NS2 protein of influenza virus is found in purified virus and phosphorylated in infected cells. Arch. Virol. 116(1–4):69–80. Richt, J. A., Lager, K. M., Janke, B. H., Woods, R. D., Webster, R. G., and Webby, R. J. (2003). Pathogenic and antigenic properties of phylogenetically distinct reassortant H3N2 swine influenza viruses cocirculating in the United States. J. Clin. Microbiol. 41(7):3198–3205. Richt, J. A., Lekcharoensuk, P., Lager, K. M., Vincent, A. L., Loiacono, C. M., Janke, B. H., Wu, W. H., Yoon, K. J., Webby, R. J., Solorzano, A., and Garcia-Sastre, A. (2006). Vaccination of pigs against swine influenza viruses using a ns1-truncated modified live virus vaccine. J. Virol. 80(22):11009–11018. Rogers, G. N., and Paulson, J. C. (1983). Receptor determinants of human and animal influenza virus isolates: Differences in receptor specificity of the H3 hemagglutinin based on species of origin. Virology 127(2):361–373. Scherle, P. A., and Gerhard, W. (1986). Functional analysis of influenza-specific helper T cell clones in vivo. T cells specific for internal viral proteins provide cognate help for B cell responses to hemagglutinin. J. Exp. Med. 164(4):1114–1128. Scholtissek, C. (1995). Molecular evolution of influenza viruses. Virus Genes 11(2–3):209–215. Scholtissek, C., Burger, H., Kistner, O., and Shortridge, K. F. (1985). The nucleoprotein as a possible major factor in determining host specificity of influenza H3N2 viruses. Virology 147(2):287–294. Sheerar, M. G., Easterday, B. C., and Hinshaw, V. S. (1989). Antigenic conservation of H1N1 swine influenza viruses. J. Gen. Virol. 70(Pt. 12):3297–3303. Shope, R. E. (1931). Swine influenza. Filtration experiments and etiology. J. Exp. Med. 54:373–385. Shortridge, K. F., and Stuart-Harris, C. H. (1982). An influenza epicentre? Lancet 2 (8302):812–813. Skehel, J. J., and Wiley, D. C. (2000). Receptor binding and membrane fusion in virus entry: The influenza hemagglutinin. Annu. Rev. Biochem. 69:531–569. Solorzano, A., Webby, R. J., Lager, K. M., Janke, B. H., Garcia-Sastre, A., and Richt, J. A. (2005). Mutations in the NS1 protein of swine influenza virus impair anti-interferon activity and confer attenuation in pigs. J. Virol. 79(12):7535–7543. Swenson, S. L., Landgraf, J. G., Killian, M. L., Shu, B., Lindstrom, S., Yu, X., Klimov, A., Vincent, A. L., Zhang, Y., and Bowman, A. S. (2008). Swine influenza virus infection of pigs and people at a county fair. AASV(Abstract). Talon, J., Salvatore, M., O’Neill, R. E., Nakaya, Y., Zheng, H., Muster, T., Garcia-Sastre, A., and Palese, P. (2000). Influenza A and B viruses expressing altered NS1 proteins: A vaccine approach. Proc. Natl. Acad. Sci. USA 97(8):4309–4314. Tamura, S., Funato, H., Hirabayashi, Y., Suzuki, Y., Nagamine, T., Aizawa, C., and Kurata, T. (1991). Cross-protection against influenza A virus infection by passively transferred respiratory tract IgA antibodies to different hemagglutinin molecules. Eur. J. Immunol. 21(6):1337–1344.
SIV: A North American Perspective
153
Tamura, S., Tanimoto, T., and Kurata, T. (2005). Mechanisms of broad cross-protection provided by influenza virus infection and their application to vaccines. Jpn. J. Infect. Dis. 58(4):195–207. Thacker, E. L., Thacker, B. J., and Janke, B. H. (2001). Interaction between Mycoplasma hyopneumoniae and swine influenza virus. J. Clin. Microbiol. 39(7):2525–2530. Tiensin, T., Chaitaweesub, P., Songserm, T., Chaisingh, A., Hoonsuwan, W., Buranathai, C., Parakamawongsa, T., Premashthira, S., Amonsin, A., Gilbert, M., Nielen, M., and Stegeman, A. (2005). Highly pathogenic avian influenza H5N1, Thailand, 2004. Emerg. Infect. Dis. 11(11):1664–1672. Townsend, H. G., Penner, S. J., Watts, T. C., Cook, A., Bogdan, J., Haines, D. M., Griffin, S., Chambers, T., Holland, R. E., Whitaker-Dowling, P., Youngner, J. S., and Sebring, R. W. (2001). Efficacy of a cold-adapted, intranasal, equine influenza vaccine: Challenge trials. Equine Vet. J. 33(7):637–643. USDA. (1995). Trends in vaccination practices on U.S. swine operations, (C. USDA:APHIS: VS, ed.) Fort Collins, CO. USDA. (2003). Trends in vaccination practices for respiratory diseases on U.S. swine operations—2000, (C. USDA:APHIS:VS, ed.) Fort Collins, CO. USDA. (2007). Swine 2006, Part II: Reference of swine health and health management practices in the United States, 2006, (C. USDA:APHIS:VS, ed.) Fort Collins, CO. Van Reeth, K., Gregory, V., Hay, A., and Pensaert, M. (2003). Protection against a European H1N2 swine influenza virus in pigs previously infected with H1N1 and/or H3N2 subtypes. Vaccine 21(13–14):1375–1381. Vincent, A. L., Lager, K. M., Ma, W., Lekcharoensuk, P., Gramer, M. R., Loiacono, C., and Richt, J. A. (2006). Evaluation of hemagglutinin subtype 1 swine influenza viruses from the United States. Vet. Microbiol. 118(3–4):212–222. Vincent, A. L., Ma, W., Lager, K. M., Janke, B. H., Webby, R. J., Garcia-Sastre, A., and Richt, J. A. (2007). Efficacy of intranasal administration of a truncated NS1 modified live influenza virus vaccine in swine. Vaccine 25(47):7999–8009. Vincent, A. L., Lager, K. M., Janke, B. H., Gramer, M. R., and Richt, J. A. (2008). Failure of protection and enhanced pneumonia with a US H1N2 swine influenza virus in pigs vaccinated with an inactivated classical swine H1N1 vaccine. Vet. Microbiol. 126 (4):310–323. Wang, C., Takeuchi, K., Pinto, L. H., and Lamb, R. A. (1993). Ion channel activity of influenza A virus M2 protein: Characterization of the amantadine block. J. Virol. 67(9):5585–5594. Wang, X., Li, M., Zheng, H., Muster, T., Palese, P., Beg, A. A., and Garcia-Sastre, A. (2000). Influenza A virus NS1 protein prevents activation of NF-kappaB and induction of alpha/ beta interferon. J. Virol. 74(24):11566–11573. Wang, X., Basler, C. F., Williams, B. R., Silverman, R. H., Palese, P., and Garcia-Sastre, A. (2002). Functional replacement of the carboxy-terminal two-thirds of the influenza A virus NS1 protein with short heterologous dimerization domains. J. Virol. 76 (24):12951–12962. Webby, R. J., and Webster, R. G. (2001). Emergence of influenza A viruses. Philos. Trans. R. Soc. Lond. B., Biol. Sci. 356(1416):1817–1828. Webby, R. J., Swenson, S. L., Krauss, S. L., Gerrish, P. J., Goyal, S. M., and Webster, R. G. (2000). Evolution of swine H3N2 influenza viruses in the United States. J. Virol. 74 (18):8243–8251. Webby, R. J., Rossow, K., Erickson, G., Sims, Y., and Webster, R. (2004). Multiple lineages of antigenically and genetically diverse influenza A virus co-circulate in the United States swine population. Virus Res. 103(1–2):67–73. Webster, R. G. (2002). The importance of animal influenza for human disease. Vaccine 20 (Suppl. 2):S16–S20.
154
Amy L. Vincent et al.
Webster, R. G., Bean, W. J., Gorman, O. T., Chambers, T. M., and Kawaoka, Y. (1992). Evolution and ecology of influenza A viruses. Microbiol. Rev. 56(1):152–179. Webster, R. G., Wright, S. M., Castrucci, M. R., Bean, W. J., and Kawaoka, Y. (1993). Influenza—A model of an emerging virus disease. Intervirology 35(1–4):16–25. Webster, R. G., Peiris, M., Chen, H., and Guan, Y. (2006). H5N1 outbreaks and enzootic influenza. Emerg. Infect. Dis. 12(1):3–8. Wesley, R. D., and Lager, K. M. (2005). Evaluation of a recombinant human adenovirus-5 vaccine administered via needle-free device and intramuscular injection for vaccination of pigs against swine influenza virus. Am. J. Vet. Res. 66(11):1943–1947. Wesley, R. D., and Lager, K. M. (2006). Overcoming maternal antibody interference by vaccination with human adenovirus 5 recombinant viruses expressing the hemagglutinin and the nucleoprotein of swine influenza virus. Vet. Microbiol. 118(1–2):67–75. Wesley, R. D., Tang, M., and Lager, K. M. (2004). Protection of weaned pigs by vaccination with human adenovirus 5 recombinant viruses expressing the hemagglutinin and the nucleoprotein of H3N2 swine influenza virus. Vaccine 22(25–26):3427–3434. Yassine, H. M., Al-Natour, M. Q., Lee, C. W., and Saif, Y. M. (2007). Interspecies and intraspecies transmission of triple reassortant H3N2 influenza A viruses. Virol. J. 4:129. Zhou, N. N., Senne, D. A., Landgraf, J. S., Swenson, S. L., Erickson, G., Rossow, K., Liu, L., Yoon, K., Krauss, S., and Webster, R. G. (1999). Genetic reassortment of avian, swine, and human influenza A viruses in American pigs. J. Virol. 73(10):8851–8856.
CHAPTER
4 Replication and Partitioning of Papillomavirus Genomes Alison A. McBride
Contents
I. Background A. Papillomavirus associated disease B. Life cycle C. Genome organization and expression D. Function of viral proteins E. Modes of viral DNA replication II. Replication Initiation A. The E1 initiator protein B. The E2 loading factor C. The replication origin D. Regulation of replication initiation III. Maintenance Replication A. Role of the E1 protein in maintenance replication B. The E2 tethering protein C. Genome partitioning in different papillomaviruses D. Other viral tethering proteins E. The role of cis elements in genome partitioning F. Papillomavirus chromosomal tethering targets G. Replication licensing H. Regulation of genome copy number and partitioning IV. Vegetative Replication V. Other Aspects of Papillomavirus Replication A. Establishment of a replication competent environment
156 156 156 159 160 161 161 162 164 168 169 172 173 173 175 176 178 179 183 183 184 186 186
Laboratory of Viral Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, Maryland Advances in Virus Research, Volume 72 ISSN 0065-3527, DOI: 10.1016/S0065-3527(08)00404-1
155
156
Alison A. McBride
B. Differences in replication strategies of different papillomaviruses C. Comparison of papillomavirus DNA replication with cellular DNA replication and replication of other viruses D. Papillomavirus replication in Saccharomyces cerevisiae E. Anti-viral replication therapies F. Papillomavirus-based vectors Acknowledgments References
Abstract
187
188 189 189 190 190 190
Papillomaviruses establish persistent infection in the dividing, basal epithelial cells of the host. The viral genome is maintained as a circular, double-stranded DNA, extrachromosomal element within these cells. Viral genome amplification occurs only when the epithelial cells differentiate and viral particles are shed in squames that are sloughed from the surface of the epithelium. There are three modes of replication in the papillomavirus life cycle. Upon entry, in the establishment phase, the viral genome is amplified to a low copy number. In the second maintenance phase, the genome replicates in dividing cells at a constant copy number, in synchrony with the cellular DNA. And finally, in the vegetative or productive phase, the viral DNA is amplified to a high copy number in differentiated cells and is destined to be packaged in viral capsids. This review discusses the cis elements and protein factors required for each stage of papillomavirus replication.
I. BACKGROUND A. Papillomavirus associated disease Papillomaviruses are ubiquitous, epitheliotrophic viruses that cause warts or papillomas. There are hundreds of different human and animal papillomavirus types that have coevolved with their hosts over millions of years. Each virus is species specific and replicates persistently in a specific type of cutaneous or mucosal epithelium. Papillomaviruses cause a spectrum of disease ranging from clinically inapparent infections, through a variety of benign lesions such as common warts, anogenital warts, and laryngeal papillomatosis (Lacey, 2005) to malignant carcinomas. In humans, persistent infection with high-oncogenic risk HPVs is responsible for virtually all cervical cancer (Parkin et al., 2005; Walboomers et al., 1999) and is associated with a subset of head and neck cancers (Gillison and Lowy, 2004).
Replication and Partitioning of Papillomavirus Genomes
157
B. Life cycle
Differentiation
All papillomaviruses have similar life cycles that are tightly linked to differentiation of the host epithelium (see Fig. 1). In a normal epithelium, only cells in the lower basal layer are mitotically active. After cell division, one daughter cell is pushed up to replenish the overlying differentiated layers, which are eventually shed from the surface of the epithelium. To initiate infection, papillomaviruses must access the dividing basal cells and it is thought that they do so through microabrasions or wounds.
Vegetative replication
Maintenance replication and genome partitioning
Initial infection and limited amplification of extrachromosomal viral DNA
FIGURE 1 Modes of Replication in the Life cycle of Papillomaviruses. The diagram shows a stratified epithelium. The lowermost basal layer provides the germinal cells necessary for renewal of the epithelium. Papillomaviruses infect these cells through microabrasions and the viral DNA undergoes a limited amplification after delivery to the nucleus. The viral genome is maintained and partitioned in a regulated manner in these dividing cells. They provide a reservoir of infected cells and, as they divide, individual infected daughter cells are pushed upwards and begin the differentiation process. These differentiated cells support vegetative replication resulting in a great amplification of the viral genome. Eventually, the cells at the outermost layer, containing virion particles, are sloughed from the epithelium.
158
Alison A. McBride
Certain cells in the epithelium, such as those in the bulge of the hair follicle or the transformation zone of the cervix or epithelial stem cells, may be more susceptible to, or more likely to support, long term persistent infection. Papillomaviruses must target a site of epithelial wounding and it has been proposed that laminin 5, secreted by migrating keratinocytes, acts as a transient receptor to trap the virions (Culp et al., 2006). Viral particles also adsorb to membrane-associated heparan sulfate proteoglycans (Giroglou et al., 2001; Joyce et al., 1999; Shafti-Keramat et al., 2003) before transfer to more specific receptors such as the candidate receptor, a6 integrin (Evander et al., 1997; McMillan et al., 1999). Viral particles enter the cell by either calveolae or clathrin-mediated endocytic pathways (Bousarghin et al., 2003; Day et al., 2003; Hindmarsh and Laimins, 2007; Smith et al., 2007). After exiting the endosomes, the L2 minor capsids protein transports the viral DNA into the nucleus to the subnuclear promyelocytic leukemia protein (PML) bodies (Day et al., 2004). Upon infection, many DNA viruses localize to, initiate transcription at, and often disrupt, these bodies, leading to the hypothesis that they are involved in antiviral defense (Negorev and Maul, 2001). In contrast, the presence of the PML protein enhances early transcription of papillomaviruses (Day et al., 2004). After transportation to the nucleus, the viral genome is amplified to a copy number that may range from 10 to 50 copies per cell, and the infected cell is driven to proliferate to provide a pool of infected basal cells to form the basis of the papilloma. The viral genomes are maintained in the nucleus of the dividing basal cells as extrachromosomal replicating elements that replicate in synchrony with the host cellular DNA in a cell cycle dependent manner. Papillomavirus infections are usually long-lived and persistent, and the continually dividing basal cells serve as a reservoir of infected cells for the continual progressive vertical differentiation that occurs during the maturation of the epithelium. Therefore, papillomaviruses need a robust mechanism to retain their extrachromosomal genomes within the nucleus of continually dividing cells to ensure that the infection is sustainable. Viral genome amplification, late capsid protein synthesis and virion assembly occur in the upper, terminally differentiated cells of the epithelium. Virions are assembled in the superficial differentiated layers and are found throughout the nuclei, frequently organized into paracrystalline arrays. These virus containing cells are destined to be sloughed from the epidermis. The strategy of restricting viral DNA amplification and synthesis of large amounts of viral antigens to the superficial layers of an epithelium ensures long-term persistent infection and is important for immune evasion by the virus. However, this strategy necessitates that the virus amplifies its DNA in cells that would normally have withdrawn
Replication and Partitioning of Papillomavirus Genomes
159
from the cell cycle and are undergoing differentiation. To achieve this, the papillomaviruses must disrupt the normal differentiation process and sustain cells in an S-phase like state so that the virus has the available enzymes to replicate its own DNA. An unintended result of cell cycle disregulation by a subset of ‘‘high risk’’ viruses is the absence of crucial cell cycle checkpoints, which can lead to genetic instability and malignant progression.
C. Genome organization and expression Papillomaviruses have circular, double-stranded DNA genomes of approximately 7000–8000 bp (see Fig. 2). The coding region is divided into early and late regions. The early region contains open reading frames designated E1 through E8, which, for the most part, are expressed in the lower, less differentiated layers of a papilloma. The late region encodes the capsid antigens, L1 and L2, which are expressed in the more superficial, differentiated cells. All viral RNA species are transcribed from one strand and undergo extensive alternative splicing. A region of approximately 500–1000 bp, located upstream from the coding region, is called the Long Control Region (LCR) or Upstream Regulatory Region (URR). This region contains the replication origin, sequences required for genome maintenance and transcriptional enhancers and promoters. MME
Origin E6 E7 E8
L1
E1
E2 L2 E4 E5
FIGURE 2 Papillomavirus genome. The early and late open reading frames, are indicated as E1–E8 and L1–L2, respectively. The replication elements MME (minichromosome maintenance element) and origin are shown in the LCR region. The E1 and E2 replication proteins are highlighted in green and purple, respectively, with the corresponding binding sites shown in the origin region.
160
Alison A. McBride
D. Function of viral proteins Papillomaviruses appear to be simple viruses that encode only a few genes, but each viral protein interacts with and regulates a multitude of cellular proteins. Overall, the functions of the viral proteins contribute to initiating an efficient, persistent infection, to evading the host immune system, and to producing large amounts of progeny virus. Basal cells act as a reservoir of cells for the continual renewal of the epithelium and their division is normally regulated by growth factors. Immediately after virus infection, the newly infected cell is driven to proliferate independently from these cellular factors in order to establish a pool of infected basal cells. The E5 protein is a membrane protein that promotes proliferation of infected basal cells by inducing constitutive, ligand-independent activation of growth factor receptors. E5 also interferes with Golgi acidification, resulting in increased cycling and activation of growth factor receptors (reviewed in (DiMaio and Mattoon, 2001). E5 down-regulates surface expression of major histocompatibility complex (MHC) class I molecules, helping the virus to evade the host immune system (Ashrafi et al., 2006; Zhang et al., 2003). The E6 and E7 proteins also induce cellular proliferation and delay differentiation of the host cells to sustain them in an S-phase like state to ensure the availability of host DNA synthesis proteins for viral DNA replication. The E7 proteins of many ‘‘high risk’’ HPVs accomplish this by binding and inactivating the cellular pRB protein, resulting in expression of S-phase genes. However, this aberrant state induces proteins, such as p53, to arrest the growth of abnormal cells. In turn, the E6 protein binds to the p53 protein and targets it for degradation, thus enabling the cells to remain in a sustained S-phase-like state. The continual division of cells without checkpoints leads to genetic instability and, eventually, a malignant phenotype (reviewed in Wise-Draper and Wells, 2008). It is not clear how the ‘‘low risk’’ papillomaviruses fulfill this same requirement as they do not inactivate pRb and p53 functions; they most likely use alternative interactions to accomplish the same goal. For example, the ‘‘low risk’’ HPV E7 proteins bind a unique pRb and calmodulin-binding scaffolding protein called p600, (DeMasi et al., 2005; Huh et al., 2005) and both ‘‘lowand high-risk’’ HPVs are able to target the pRB family member p130 for degradation (Zhang et al., 2006). ‘‘Low risk’’ E6 proteins disrupt the actin cytoskeleton and bind a number of cellular proteins, including ERC-55, the focal adhesion protein paxillin, the E3 ubiquitin ligase E6AP, and the clathrin adaptor complex AP-1 (reviewed in Wise-Draper and Wells, 2008). E5, E6, and E7 are also important for immune evasion and they act by down regulating interferon responsive genes and MHC Class I surface expression (reviewed in Woodworth, 2002).
Replication and Partitioning of Papillomavirus Genomes
161
E1 is the primary replication initiator protein. It binds and unwinds the viral replication origin in concert with the E2 protein. The E2 protein is the major transcriptional regulator of the virus and both activates and represses transcription from viral promoters. Additionally, the E2 protein maintains and partitions the replicating extrachromosomal genome. This review will focus on the replication associated functions of these proteins. The E4 protein, usually expressed from a spliced mRNA species as E1^E4, is synthesized at very high levels in the differentiated layers of the papilloma. E4 interferes with the cell cycle and may also be important for sustaining an S-phase like state in cells that are amplifying viral DNA (Davy et al., 2002). E4 also increases the egress of virions by weakening the virus laden cornified envelopes (Brown et al., 2006) and by disrupting keratin filaments (Doorbar et al., 1991). Papillomavirus virions form non-enveloped icosahedral structures of 55–60 nm diameter containing the genome packaged in a nucleohistone complex (Favre et al., 1977). L1 is the major capsid antigen and can selfassemble into virus like particles (Kirnbauer et al., 1992; Zhou et al., 1991). L2, the minor capsid protein, is important late in infection for packaging the viral genome into the capsid (Day et al., 1998; Zhao et al., 1998) and at early stages of infection to transport the viral DNA to the nucleus to establish a permissive infection (Day et al., 2004).
E. Modes of viral DNA replication The papillomavirus life cycle requires three different modes of viral DNA replication which can be defined as ‘‘Establishment’’, ‘‘Maintenance’’, and ‘‘Amplification’’. Upon initial infection, the incoming viral genome is transported to the nucleus where it undergoes a limited amplification to a low copy number to establish the infection. In the second stage of replication, these genomes are replicated and maintained at a constant copy number in mitotically active basal cells. And finally, as the infected cells differentiate and mature, the genome is vegetatively amplified to a large copy number to provide progeny genomes to be encapsidated in virion particles. Some of these steps are very well understood, but others still remain elusive.
II. REPLICATION INITIATION Initiation of papillomavirus DNA replication requires the replication origin, which minimally contains an E1 and E2 binding site (Mohr et al., 1990; Ustav and Stenlund, 1991; Ustav et al., 1991), and the viral E1 and E2 proteins. E1 is the primary replication protein; it is an ATP-dependent
162
Alison A. McBride
helicase that specifically binds, melts, and unwinds the viral replication origin to allow access of the cellular replication proteins. Initially, E1 cooperatively binds to the origin with the E2 protein but E1 then assembles into double hexamers that encircle the DNA and bidirectionally unwind the origin.
A. The E1 initiator protein The papillomavirus E1 proteins are 70-kDa ATP-dependent helicases that bind specifically to the origin to initiate replication (Yang et al., 1993). They consist of four domains; an N-terminal domain, a sequence specific DNA binding domain, an oligomerization domain, and a helicase domain (see Fig. 3A). The replication origin contains binding sites for E1 and the loading protein, E2 (Ustav et al., 1991). As shown in Fig. 4, a dimer of E1 and a dimer of E2 cooperatively bind to their adjacent sites with the N-terminal domain of E2 forming an interaction with the helicase domain of E1 (Sarafi and McBride, 1995; Sedman et al., 1997). After loading, E2 is dissociated from the complex and E1 coverts into a double hexameric ring helicase that encircles the DNA (Sanders and Stenlund, 1998; Schuck and
A
S235P
S301P S298P
BPV-1 E2 DNA binding Dimerization
Transactivation B BPV-1
S283P S305P
S109P S48P T102P
K514SUMO S584P
NLS
E1 HPV-11
NES S89P Cyclin S93P S107P binding
T468P
motif
N
DNA binding
Oligomerization
Helicase
FIGURE 3 Domain structure of the E1 and E2 proteins. (A) The full-length E2 proteins of all papillomaviruses consist of two conserved domains linked by a less conserved hinge region. The N-terminal domain is important for transcriptional regulation and interaction with the E1 protein. The C-terminal domain is a specific DNA binding and dimerization domain. Phosphorylation sites mapped in BPV-1 E2 are indicated (Lehman and Botchan, 1998; McBride et al., 1989). (B) The E1 proteins consist of four domains. The N-terminal domain is important for intracellular localization and contains both nuclear localization signals (NLS) and nuclear export signals (NES). The next domain is an origin specific DNA binding domain, followed by an oligomerization domain. The C-terminal domain contains the helicase function. Phosphorylation sites mapped in BPV-1 are indicated above and those mapped in HPV11 are indicated below. See text for references.
Replication and Partitioning of Papillomavirus Genomes
TA
TA H
H
N
N O
163
DBD
DBD
O
DBD
DBD
N H
DBD O
O
H
DBD N
FIGURE 4 Initiation of DNA replication. Initially, dimers of the E1 and E2 proteins co-operate together to bind to their specific binding sites on the replication origin. The transactivation domain of E2 interacts with the helicase domain of E1. This complex converts to a double E1 hexamer that encircles the origin. In the absence of E2, the helicase domain can make additional, non-specific interactions with DNA surrounding the origin.
Stenlund, 2005; Sedman and Stenlund, 1998; Titolo et al., 2000). E1 has the ability to bind DNA both specifically and non-specifically via two different DNA binding regions. Site-specific DNA binding is provided by the DNA binding domain (DBD) (Chen and Stenlund, 1998; Sarafi and McBride, 1995), which recognizes and binds to two pairs of E1 binding sites (E1 BS) in the replication origin. The helicase domain also binds DNA, but with low sequence specificity. This activity is required for the ability of the E1 helicase domain to contact the DNA sequences flanking the E1 BS, including a region that has been termed the A/T-rich region (Schuck and Stenlund, 2006). In the initial binding complex the interaction of the E2 protein with the helicase domain masks this non-specific DNA binding function, but when E2 is disassociated the non-specific DNA binding region is available to interact with DNA (Stenlund, 2003). An
164
Alison A. McBride
X-ray crystal structure of the BPV-1 E1 helicase, bound to DNA, has shown that single-stranded DNA is threaded through the hexamer channel and so the replication origin must be melted as E1 converts from a dimer to a double hexamer (Enemark and Joshua-Tor, 2006). Compared to most known cellular helicases, virally encoded helicase proteins such as E1 and SV40 T antigen are unique in their ability to cause melting, as well as unwinding, of the DNA helix. For E1, a beta hairpin (also found in T antigen) pries apart the DNA strands to melt the helix (Castella et al., 2006; Liu et al., 2007). When the double hexamer has formed, each subunit of the helicase domain projects its beta hairpin into the channel of the helicase in a right-handed staircase pattern. Each hairpin contacts adjacent nucleotides of single stranded DNA in a sequential pattern and ATP hydrolysis and ADP release causes the helicase to move along these nucleotides in what has been termed a coordinated escort mechanism (Enemark and Joshua-Tor, 2006). Each of the hexamers translocates bidirectionally with a 30 to 50 polarity to unwind the origin. The oligomerization domain provides a rigid collar around the DNA to help stabilize the interactions between the helicase domains (Enemark and Joshua-Tor, 2006). The structure of the E1 DNA binding domain and the hexameric helicase are shown in Fig. 5D and E, respectively. The N-terminal domain of E1 is only required for replication in vivo (Ferran and McBride, 1998; Sun et al., 1998) and likely plays a role in regulating intracellular localization. The E1 protein shuttles from the nucleus to the cytoplasm and the N-terminal domain contains both nuclear import and export signals that are regulated by cyclin A/ECdk2 (Deng et al., 2004; Hsu et al., 2007; Lentz et al., 1993; Ma et al., 1999). A conserved caspase cleavage site located between residues 46 and 49 in this domain is also important for vegetative DNA replication (Moody et al., 2007) (see below). In addition to origin recognition, melting and unwinding, E1 recruits and interacts with many cellular replication factors. Viral DNA synthesis requires replication protein A (RPA) to stabilize single stranded DNA, topoisomerase I to relieve tortional stress, and the host DNA polymerase a primase to prime replication (Clower et al., 2006; Conger et al., 1999; Han et al., 1999; Kuo et al., 1994; Loo and Melendy, 2004; Masterson et al., 1998; Melendy et al., 1995; Muller et al., 1994.). Also recruited are Proliferating Cell Nuclear Antigen (PCNA), Replication Factor C (RFC), and Polymerase d (Kuo et al., 1994).
B. The E2 loading factor The papillomavirus E2 proteins regulate transcription, as well as being required for viral DNA replication. The E2 gene encodes multiple proteins that are the result of expression from different promoters and/or
Replication and Partitioning of Papillomavirus Genomes
165
FIGURE 5 Structures of Papillomavirus E1 and E2 protein domains and complexes. (A) The transactivation domain (residues 1–201) of the HPV-16 E2 protein from the pdb file 1DTO (Antson et al., 2000). Residues R37 and I73 are shown in green and residue D39 is shown in blue. (B) A dimer of the DNA binding/dimerization domain of BPV-1 E2 bound to DNA (residues 326–410) from the pdb file 2BOP (Hegde et al., 1992). The DNA is shown in gray. (C) The transactivation domain of HPV-18 E2 (residues 1–215; red) in complex with a c-terminal domain of HPV18 E1 (residues 428–631; green) from the pdb file 1TUE (Abbate et al., 2004). E2 residue D43 (equivalent to BPV-1 D39 in panel A)
166
Alison A. McBride
alternative RNA splicing. The full-length E2 protein is a polypeptide of between 350 and 500 amino acids in length, and is often referred to as E2-TA or E2 transactivator. E2-TA regulates the activity of viral promoters by binding to multiple 12bp palindromic sequences in E2-specific enhancers within the LCR. As shown in Fig. 3A, E2 consists of two conserved domains; a 200 amino acid N-terminal domain, which is important for transcriptional activation and repression, interaction with the E1 protein, as well as many cellular factors, and a C-terminal domain of approximately 100 amino acids that has sequence-specific DNA binding and dimerization properties (reviewed in (McBride and Myers, 1997). The region between these domains varies in sequence and in length and is not well conserved among different genera of viruses. However, within each papillomavirus genus, the E2 hinge region contains auxiliary elements that are only conserved among related viruses. For example, there are elements important for regulated E2 degradation in BPV-1 E2 (Penrose and McBride, 2000), for nuclear localization in HPV-11 E2 (Lai et al., 1999; Zou et al., 2000), and for mitotic chromosome binding (Poddar et al., 2008) and transcriptional regulation (Steger et al., 2002) in HPV-8 E2. The DNA binding domain of the E2 proteins forms a dimeric betabarrel structure that positions two alpha recognition helices to contact the consensus binding site (Hegde et al., 1992). A crystal structure of the BPV1 E2 DNA binding domain is shown in Fig. 5B. The consensus binding site motif with the highest affinity has the sequence ACCGN4CGGT, with a preference for AT nucleotides in the center for many HPVs (Androphy et al., 1987; Bedrosian and Bastia, 1990; Li et al., 1989). The E2 protein also binds to the motif ACCN6GGT and binding to sites AACN6GGT, ACCN6GTT, and ACAN5CGGT has also been observed (Li et al., 1989; Newhouse and Silverstein, 2001). Some viruses contain few consensus E2 sites (the genital HPVs have four conserved sites) and others contain many more (the BPV-1 genome has 17 binding sites). These sites are important for viral transcriptional regulation, initiation of replication, and for viral genome partitioning and maintenance. Figure 6 shows the position of E2 binding sites in the LCR of several different papillomaviruses. The E2-TA protein recruits many basic cellular transcription factors and coactivators to activate viral transcription. E2 also represses
is shown in blue. (D) A dimer of the BPV-1 E1 DNA binding domain (residues 159–303) from the pdb file 1FO8 (Enemark et al., 2000). Residues shown in blue are important for DNA contact. (E) Double hexameric ring structure of the BPV-1 E1 helicase domain (residues 306–577) from the pdb file 2GXA (Enemark and Joshua-Tor, 2006). Each E1 monomer is shown in a different color.
167
Replication and Partitioning of Papillomavirus Genomes
HPV-16
E6 Alpha
L1
HPV-8
HPV-50
BPV-1
L1
E6
L1
E6
Beta
Gamma
E6 Delta
L1
CRPV
L1
E6
Kappa
FIGURE 6 Map of the E1 and E2 binding sites in the long control regions in a range of papillomaviruses. The LCRs of papillomaviruses from five different genera are shown. The end of the L1 and beginning of the E6 open reading frames are indicated. The E1 binding site is indicated as a green box. E2 binding sites that match the consensus ACCN6GGT are shown in purple. E2 sites that deviate from this consensus are shown in hatched purple.
transcription from viral promoters when the E2 binding sites overlap essential promoter elements, such as the TATA box or Sp1 binding sites (Bernard et al., 1989; Cripe et al., 1987; Thierry and Yaniv, 1987). However, repression also requires a function of the transactivation domain in addition to DNA binding (Dowhanick et al., 1995; Goodwin et al., 1998; Soeda et al., 2006; Thierry and Yaniv, 1987). The transactivation domain of the E2 proteins is also well characterized and forms a cashew shaped structure consisting of a bundle of three alpha helices in the N-terminal half linked to a beta sheet region in the C-terminal half by a region that has been termed the fulcrum (Antson et al., 2000). The structure of the E2 transactivation domain in complex with the helicase domain of E1 has also been solved (Abbate et al., 2004). Representative structures are shown in Fig. 5. The regions of the transactivation domain required for transcriptional regulation and replication are well defined and separated (Abroi et al., 1996; Brokaw et al., 1996; Sakai et al., 1996). For example, E2 residues arginine 37 and isoleucine 73, located on the face of two adjacent helices in the N-terminal portion of the transactivation domain, are important for transcriptional regulation (Antson et al., 2000) whereas glutamate at position 39 on the opposite side of the domain is key for interaction with arginine 454 in the helicase domain of E1 (Abbate et al., 2004). The E2 transactivation domain has also been implicated in selfinteraction and looping of DNA containing E2 binding sites and looped
168
Alison A. McBride
E2-DNA structures have been observed by electron microscopy for BPV-1, HPV-16, and HPV-11 E2 proteins (Antson et al., 2000; Hernandez-Ramon et al., 2008; Knight et al., 1991; Sim et al., 2008). However, the precise interaction between N-terminal domains of different E2 proteins, at least in the observed crystal structures, is not consistent. In the X-ray crystal structure of the HPV16 E2 transactivation domain, intermolecular interactions were observed between residues arginine 37 and isoleucine 73, which are crucial for the transcriptional activation function (Antson et al., 2000). In contrast, a crystal structure of the BPV-1 N-terminal domain revealed a disulphide bond between cysteine residues 57, which interfered with the E1 interaction face of the domain (Sanders et al., 2007). A different monomer-monomer interface was also observed in the protein crystal lattice obtained for the HPV-11 E2 protein (Wang et al., 2004). Thus, selfinteraction of the transactivation domains may be important for E2 function, but the interactions observed in X-ray structures could also be the result of non-specific crystal packaging interfaces ( Janin et al., 2007). Several additional E2 functions are important for initiation of viral DNA replication. As described above, E2 helps load the E1 helicase by cooperatively binding to adjacent binding motifs in the origin and by increasing the specificity of E1 DNA binding by masking its non-specific binding activity (Stenlund, 2003). E2 also enhances replication by alleviating repression by nucleosomes to allow the E1/E2 complex to bind to the origin (Li and Botchan, 1994). Furthermore, E2 interacts with RPA, a single-stranded binding protein required for DNA replication (Li and Botchan, 1993) and with Replication Factor C, RF/C (Wu et al., 2006).
C. The replication origin The papillomavirus replication origin contains an E1 binding site, several E2 binding sites, and an A/T rich region (Mohr et al., 1990; Ustav et al., 1991, 1993). The E1 binding sites consist of an 18bp palindrome that contains multiple overlapping recognition sequences for the E1 protein (Chen and Stenlund, 2001; Holt et al., 1994; Titolo et al., 2003). E1 is a monomer in solution but binds to the origin as a dimer, along with a dimer of the E2 protein. The E1 binding sites consist of overlapping hexanucleotide sequences with a consensus ATTGTT, which are separated by three nucleotides from the adjacent half of the palindrome (Auster and Joshua-Tor, 2004). The sequence is not highly conserved but, in most cases, the thymidines at position 2 and 5 are invariant. The sequences of a series of E1 binding sites are shown in Fig. 7. For most papillomaviruses, at least one E2 binding site is required for transient replication and additional sites greatly stimulate replication (Akgul et al., 2003; Chiang et al., 1992; Lee et al., 1997; McShan and Wilson, 1997b; Remm et al., 1992; Russell and Botchan, 1995;
Replication and Partitioning of Papillomavirus Genomes
HPV-16 HPV-11 HPV-8 BPV-1 OvPV-1 HPV-1 HPV-4 CRPV
ATTGTAGTT ATTGTTATT GTTATTGCC ATTGTTGTT ATTGTTGTT ATGGTAGTT ATCATAGTT GTTGTTGCT
TAGTATTAT AACTATAAG AACAACCAT AACAATAAT AACAATAAT AACAACAAC GGCAACAAT AACAATAAT
169
]Alpha Beta
]Delta
mu Gamma Kappa
FIGURE 7 E1 binding origin sequences. The palindromic E1 binding origin sequence from a range of papillomaviruses is shown. The genus to which each virus belongs is shown to the right.
Sun et al., 1996). In some cases, the E1 binding site is dispensable and two E2 binding sites can suffice (Lu et al., 1995; Sverdrup and Khan, 1995). Presumably, in the latter case the E2 protein can load the E1 helicase efficiently enough in the absence of specific E1 binding. It has also been shown that, at high levels, the E1 protein can support replication in the absence of the E2 protein (Gopalakrishnan and Khan, 1995). Again, presumably at higher concentrations the E1 protein is less dependent on E2 for loading. Although these studies provide insight into the mechanisms of viral replication, it is almost certain that in the actual viral life cycle initiation of DNA replication in the initial amplification stage requires both the E1 and E2 proteins, and their cognate binding sites. HPV-11 E2 can be observed as a large disk/ring-shaped protein particle bound to the three E2 sites adjacent to the HPV-11 origin. Inclusion of the fourth, upstream E2 site results in DNA molecules containing loops from this site to the promoter proximal sites (Sim et al., 2008).
D. Regulation of replication initiation There are many ways in which papillomavirus replication can be regulated. The mechanisms described in this section could regulate initiation of DNA synthesis in the establishment phase but may also affect maintenance replication and vegetative viral DNA amplification. Additional regulatory mechanisms that are more likely to be specific for the latter modes are described later in the article. After initial infection, the papillomavirus DNA is delivered to the nucleus in complex with the L2 minor capsid protein (Day et al., 2004) and, similar to many other viruses, localizes adjacent to the PML nuclear domains. L2 also recruits E2 to these domains (Day et al., 1998) and presumably this is a privileged site for viral DNA replication. Transient replication studies with HPV-11, show that the E1 and E2 proteins accumulate in replication foci that are often associated with PML nuclear
170
Alison A. McBride
domains (Swindle et al., 1999). E1 localization is also regulated by sumoylation (Rangasamy and Wilson, 2000), which may regulate intranuclear localization (Rangasamy et al., 2000) and E1 access to the nuclear replication domains. Although not well understood, it is likely that localization to the subnuclear sites of viral replication is tightly regulated. The E1 proteins are extensively regulated by posttranslational modifications and at least ten phosphorylation sites have been identified in the BPV-1 E1 protein (Lentz et al., 2006). It appears that many of these phosphorylation events regulate nucleocytoplasmic shuttling of the E1 proteins. The N-terminal domain of the E1 protein contains both a bipartite nuclear localization signal (NLS) (Lentz et al., 1993) and a nuclear export signal (NES) (Rosas-Acosta and Wilson, 2008) and localization appears to be regulated by phosphorylation (Bian et al., 2007; Deng et al., 2004; Hsu et al., 2007; Yu et al., 2007a). The cell cycle kinases that modify the cellular MCM helicase complex, cyclin E/cdk2, and cyclin A/cdk2, also phosphorylate the E1 proteins from several papillomaviruses (Cueille et al., 1998; Lin et al., 2000; Ma et al., 1999). The E1 proteins are also modified by CK2 (Lentz, 2002; McShan and Wilson, 1997a), mitogenactivated protein kinases (Yu et al., 2007a), protein kinases A and C (Zanardi et al., 1997), and p34cdc2 (Lentz et al., 1993). Initiation of replication in the establishment phase is dependent on the levels of the E1 and E2 proteins. Levels of both proteins can be regulated at the transcriptional and translational levels, but there is strong evidence that turnover of both proteins is tightly regulated. The BPV-1 E1 protein is quite short-lived and is a target of the ubiquitin ligase APC (Mechali et al., 2004). However, when it is bound to cyclin E-cyclin-dependent kinase 2 (Cdk2) before the start of DNA synthesis, E1 becomes resistant to ubiquitin-mediated degradation. Thus, APC controls the levels of E1, and this might be important to limit the level of amplificational replication and/or to maintain a constant low copy number of the viral genome during the maintenance phase of replication. The half-life of the E2 protein is also regulated by CK2 phosphorylation (Penrose et al., 2004), which in turn regulates the levels of replication and genome copy number. The mechanism will be described in more detail below. Both the E1 and E2 open reading frames encode truncated products that can potentially negatively regulate replication. For example, some papillomaviruses encode a truncated E1 protein, designated E1-M or E1 modulator, which consists of the N-terminal domain of the E1 protein (Lusky and Botchan, 1986; Rotenberg et al., 1989). However, although earlier studies indicated that the BPV-1 E1-M protein might modulate maintenance replication (Berg et al., 1986), subsequent studies showed that it was non-essential (Hubert and Lambert, 1993). Most papillomaviruses also encode truncated E2 protein products. For example, BPV-1 encodes two E2 ‘‘repressor’’ proteins. One, E2-TR, is expressed from an
Replication and Partitioning of Papillomavirus Genomes
171
internal promoter and initiation codon (Lambert et al., 1987) and the other is encoded by a spliced message that fuses 11 amino acids from the E8 open reading frame to the C-terminal domain of E2 (Choe et al., 1989; Lambert et al., 1990). All E2 proteins contain the DNA binding and dimerization domain and so the shorter forms of the protein can antagonize the activities of the E2-TA protein, either by competing for binding to the DNA sites or by forming heterodimers (Barsoum et al., 1992; Lambert et al., 1987; Lim et al., 1998). Similar truncated E2 proteins that can regulate both transient amplification replication and stable episomal maintenance are encoded by the HPVs. For example, an E1M^E2C protein encoded by HPV-11 (Chiang et al., 1992), an HPV-31 E8^E2C protein (Stubenrauch et al., 2000), and a CRPV E9^E2C protein (Jeckel et al., 2003) all regulate replication of their respective viruses. The HPV-31 E8^E2 protein has been studied in some detail and it seems that the E8 residues contribute specifically to this function and it is not simply due to direct interference with E2 DNA binding domain functions (Zobel et al., 2003). Of note is the fact that the BPV-1 E2 binding sites adjacent to the origin have relatively weak affinity for the E2 protein compared to other sites (Li et al., 1989) and binding is only efficient in the presence of the E1 protein (Sarafi and McBride, 1995). Since the E2 transactivation domain is required for efficient co-operative binding with E1 (Winokur and McBride, 1996), this would preclude binding of the shorter E2-TR proteins to these sites. This mechanism could allow the E2 repressor proteins to regulate E2-TA binding at other sites (e.g. for transcription) without interfering with replication. Papillomavirus replication is also regulated by the p53 protein. p53 inhibits transient amplificational replication of BPV-1 and HPV-11 but does not affect stable maintenance replication (Ilves et al., 2003). An intact DNA-binding and oligomerization domain of p53 is necessary for repression, while the N-terminal transcription activation and C-terminal regulatory domains are dispensible (Ilves et al., 2003). Replication inhibition can be mediated by direct binding of p53 to the HPV16 E2 protein (Brown et al., 2008) or, as in the case of HPV-8, by competitively binding to a p53 site adjacent to the origin which overlaps an E2 site (Akgul et al., 2003). This inhibition may be important to limit the amount of replication in the newly infected basal cells. The papillomavirus E6 proteins bind, and in some cases degrade, the p53 protein. In turn, the E2 protein modulates transcriptional expression of the E6/E7 genes. These components could form a regulatory loop to balance and limit replication (Ilves et al., 2003). Papillomavirus replication is also regulated by cellular factors such as the TATA-binding protein (TBP), Yin-yang 1 (YY1), and the CCAAT displacement protein (CDP). TBP prevents formation of the HPV 11 E1-E2 complex at the origin of replication by antagonizing E2 binding to the origin (Hartley and Alexander, 2002) and YY1 can inhibit replication by
172
Alison A. McBride
sequestering the E2 protein (Lee et al., 1998; O’Connor et al., 1996). CDP represses transcription and replication (Narahari et al., 2006; O’Connor et al., 2000; Pattison et al., 1997) by binding to a site that overlaps the E1 binding site in the replication origin of many papillomaviruses. Binding of the E2 proteins to the origin region helps to relieve repression by displacing CDP (Narahari et al., 2006). Since CDP is expressed in undifferentiated but not in differentiated cells (Ai et al., 1999), keratinocyte differentiation would alleviate repression and permit vegetative genome amplification. Clearly there is much interplay between regulation of transcription and replication in the early stages of the papillomavirus life cycle. Cellular proteins inhibit replication both directly and indirectly by modulating expression of the E1 and E2 replication proteins. In many cases, the viral E6 and E7 proteins are equipped to counteract the effects of these cellular repressors and the E2 protein itself can alleviate repression. At a certain level, the E2 protein downregulates E6 and E7 expression and so all of these processes can reach equilibrium.
III. MAINTENANCE REPLICATION After the initial amplification step, the viral genome must be maintained in the dividing basal cells to sustain a persistent infection. The genomes can be maintained indefinitely at a constant copy number as extrachromosomal elements. The genomes are very difficult to detect by in situ techniques in the basal cells of a papilloma and thus there are estimated to be less than 20 copies per cell (Evans et al., 2003). A few cell lines that have been isolated from clinical lesions also maintain the viral genome extrachromosomally (Bedell et al., 1991; Stanley et al., 1989), as do keratinocytes transfected with the viral genome (Flores et al., 1999; Frattini et al., 1996; Meyers et al., 1997). The genome replicates in these cells in culture at about 50 to 200 copies per cell. As described above, initiation of viral DNA replication requires the E1 and E2 proteins and the origin of replication. However, plasmids containing the minimal origin will only replicate transiently and the replicated DNA is lost with time. Long-term, stable replication of BPV-1 requires multiple E2 DNA binding sites in cis to the origin (Piirsoo et al., 1996). This observation was pivotal in identifying that the E2 protein had a separate function in viral genome maintenance. It was subsequently shown that E2 partitions and maintains the viral genomes by attaching them to mitotic chromosomes (Ilves et al., 1999; Skiadopoulos and McBride, 1998). A model of E2-mediated viral genome partitioning is shown in Fig. 8A and B. It can be argued that a specific viral genome partitioning mechanism is not required when there are sufficient copies of the viral genome. In this
Replication and Partitioning of Papillomavirus Genomes
173
scenario, each daughter cell is likely to randomly contain at least one genome, which can be amplified to a level established by a copy number control mechanism. However, compared to cell lines, the viral genome copy number in the basal cells of papillomas is virtually undetectable (Evans et al., 2003). Furthermore, it has been shown that the E2 protein tethers the viral genomes to mitotic chromosomes. This ensures that the viral genomes are partitioned to daughter cells in approximately equal numbers and guarantees that they are retained in the nucleus after cell division. Furthermore, this strategy of faithfully maintaining and partitioning viral genomes by tethering them to host mitotic chromosomes using a viral DNA binding protein is common to many persistent DNA viruses (McBride et al., 2006).
A. Role of the E1 protein in maintenance replication In general, it has been assumed that initiation of replication of the viral DNA in the maintenance phase occurs in an E1 and E2-dependent manner, similar to that described for the establishment phase of replication. However, it is also feasible that viral DNA synthesis could be initiated by cellular proteins in the maintenance phase and that viral functions are only required for retention and partitioning of the viral genomes. Evidence that E1 might not be required for maintenance replication comes from a study with a BPV-1 genome encoding a temperature sensitive E1 protein (Kim and Lambert, 2002). It was observed that while the E1 function was required for establishment of the genomes, they could be maintained as extrachromosomal elements at the non-permissive temperature, thus, indicating that E1 was not absolutely required. Other evidence that the modes of replication initiation might be different comes from the observation that p53 can inhibit amplificational replication but does not seem to repress stable episomal maintenance (Ilves et al., 2003). In general, the requirements for initiation of replication may be less stringent than previously presumed as recent studies have revealed that cellular origins may be a passive result of local chromatin structure rather than well defined sequence elements (Gilbert, 2004).
B. The E2 tethering protein The first clue to the precise role of E2 in genome partitioning was the observation that both the BPV-1 E2 protein and the viral genomes are observed as small speckles over the arms of all mitotic chromosomes (Skiadopoulos and McBride, 1998; Zheng et al., 2005). Further analysis showed that E2 binds mitotic chromosomes through protein-protein interactions mediated by the transactivation domain. The DNA binding domain binds to multiple E2 binding sites in the viral genome
174
Alison A. McBride
FIGURE 8 Mechanism of papillomavirus genome partitioning. (A). Model for partitioning of papillomavirus genomes. Cellular chromosomes (blue crescents) are partitioned by attachment to the spindle in mitotis. The viral E2 protein (purple) binds to the viral genome and tethers it to mitotic chromosomes, thus hitchhiking on the cellular chromosomes. (B). Model for chromosomal tethering by the BPV-1 E2 protein. The E2 DNA binding domain binds to sites in the viral genome while the transactivation domain interacts with proteins, such as Brd4, on mitotic chromosomes. (C) Binding patterns of three different E2 proteins, as detected by immunofluorescence, on
Replication and Partitioning of Papillomavirus Genomes
175
(in the MME) and tethers it to the condensed chromosomes (Bastien and McBride, 2000; Ilves et al., 1999). In cells expressing E2, plasmids containing at least eight E2 binding sites are maintained and segregated and are observed bound to mitotic chromosomes (Ilves et al., 1999; Piirsoo et al., 1996). Examples of the pattern of mitotic chromosome binding of different E2 proteins are shown in Fig. 8.
C. Genome partitioning in different papillomaviruses It was initially assumed that all papillomaviruses would maintain their genomes using a conserved mechanism similar to BPV-1. However, characterization of E2 tethering in other papillomaviruses has revealed that, although most E2 proteins associate with mitotic chromosomes, the E2 chromosomal target varies for different papillomaviruses. A study of the mitotic chromosomal binding pattern of E2 proteins from 13 different papillomaviruses showed that most E2 proteins, from very different genera and host species, were found bound to cellular mitotic chromosomes. However, the binding pattern varied among different E2 proteins (Oliveira et al., 2006). For example, BPV-1 E2 is detected in many small speckles over the arms of all chromosomes but HPV-8 E2 binds primarily to the peri-centromeric region of a subset of chromosomes. The binding target of the alpha-papillomaviruses, which infect the mucosal epithelia and include the cancer associated ‘‘high-risk’’ HPVs, has been more controversial. Using conditions similar to those used for the BPV-1 studies, the alpha papillomaviruses E2 proteins are only found in association with mitotic chromosomes in early (prophase) and late (telophase) stages of mitosis (Donaldson et al., 2007; Gammoh et al., 2006; Oliveira et al., 2006). In theory, the association of the E2 protein/viral genome complex with telophase chromatin would be sufficient to ensure that the genomes are partitioned and maintained in the nucleus. However, under certain fixation conditions, the alpha E2 proteins are observed in a pericentromeric pattern on metaphase chromosomes (Oliveira et al., 2006) and so may have a target similar to that of the beta-papillomaviruses (as exemplified by HPV-8 E2). The E2 proteins of the alpha-papillomaviruses may need a specialized cellular environment or additional cellular or viral factors to stabilize their association with mitotic chromosomes. Notably, stable extrachromosomal genome replication of these viruses
mitotic chromosomes. BPV-1 E2 forms small speckles on the arms of all chromosomes. HPV-8 E2 is observed in fewer, larger speckles that bind adjacent to the centromere of acrocentric chromosomes. HPV-11 E2 is also observed binding to the pericentromeric region of chromosomes, but only in certain fixation conditions (Oliveira et al., 2006). The E2 proteins are shown in green, cellular DNA in blue and the mitotic spindle in red.
176
Alison A. McBride
also requires E6 and E7 gene functions (Park and Androphy, 2002; Thomas et al., 1999). Unfortunately, the E2 proteins are very difficult to detect when expressed from replicating viral genomes and so it is still not clear whether the localization observed reflects the mechanism of genome partitioning used by these viruses. A different study of the alpha-papillomavirus HPV11 E2 protein demonstrated that both the N-terminal and C-terminal E2 domains, when fused to GFP, could independently associate with the mitotic spindle rather than the chromosomes (Van Tine et al., 2004). This localization was found to be dynamic, similar to many cellular chromosomal passenger proteins, and by late stages of mitosis the C-terminal domain of E2 was observed associated with the central spindle microtubules and by cytokinesis was located in the midbody (Dao et al., 2006). A similar study reports the colocalization of a small percentage of the BPV-1 E2 protein with the central spindle microtubules, in complex with the mitotic kinesin-like protein, MKlp (Yu et al., 2007b).Whether this microtubule localization of E2 is involved in genome partitioning remains to be determined as genomes tethered to E2 through this interaction would ultimately be stranded in the cytoplasm of cells.
D. Other viral tethering proteins Remarkably, the strategy of maintaining and partitioning extrachromosomal viral genomes by tethering them to cellular chromosomes is a common strategy among diverse DNA viruses that cause persistent infection. This has been shown for the gamma herpesviruses EpsteinBarr virus (EBV), Kaposi’s sarcoma associated Herpesvirus (KSHV), Herpesvirus saimiri (HVS) and murine gamma herpesvirus-68 (MHV68). Each virus encodes a DNA binding protein that binds specifically to repeated sites in the viral DNA and tethers the genome to the cellular mitotic chromosomes. The best studied of these herpesvirus tethering proteins are EBNA-1 and LANA, from EBV and KHSV, respectively. It has been proposed that EBNA-1 interacts with mitotic chromosomes in two different ways. The nucleolar proliferation antigen, p40, otherwise known as EBP2, is the major chromosomal protein target for EBNA-1 (Wu et al., 2000). However, the regions of EBNA-1 that bind mitotic chromosomes have also been proposed to be AT hooks that directly interact with the cellular DNA (Sears et al., 2004). LANA interacts with a number of different mitotic chromosome binding proteins such as histone H1, DEK, Methyl CpG binding protein, Brd2, Brd4 and a H2A/H2B histone dimer (Barbera et al., 2006; Cotter and Robertson, 1999; Krithivas et al., 2002; Platt et al., 1999; You et al., 2006). These targets are summarized in Table I.
TABLE I Chromosomal targets of viral tethering proteins Virus
Tethering Protein
Putative Target or Accessory factor
References
Papillomaviruses BPV-1 HPV-8 HPV-11 BPV-1, HPV-16
E2 E2 E2 E2
Brd4 rDNA Mitotic spindle ChLR1
(Baxter et al., 2005; You et al., 2004) (Poddar et al., 2008) (Van Tine et al., 2004) (Parish et al., 2006)
Herpesviruses Epstein-Barr Virus, EBV
EBNA-1
hEBP2 (p40) A-T hook
(Kapoor and Frappier, 2003) (Sears et al., 2004)
LANA
Brd2
ORF 73
Brd4 Histone H1 Histones H2A/B DEK, MeCBP NuMA MeCP2
(Platt et al., 1999; Viejo-Borbolla et al., 2005) (You et al., 2006) (Cotter and Robertson, 1999) (Barbera et al., 2006) (Krithivas et al., 2002) (Si et al., 2008) (Calderwood et al., 2004; Griffiths and Whitehouse, 2007)
Kaposi’s sarcoma associated herpesvirus (HHV-8)
Herpesvirus saimiri
178
Alison A. McBride
The EBNA-1 and LANA tethering proteins have many features in common with E2. In addition to genome tethering, each protein regulates viral transcription and is involved in initiation of viral DNA replication (although the herpesviruses have no E1 counterpart and use cellular proteins to initiate replication). Despite having no sequence homology, the DNA binding domains of E2 and EBNA (Bochkarev et al., 1995) and the putative structure of LANA (Grundhoff and Ganem, 2003) form similar dimeric anti-parallel beta-barrel DNA binding structures.
E. The role of cis elements in genome partitioning The LCR of BPV-1 contains 11 E2 binding sites (Li et al., 1989); at least eight of which are required for genome maintenance and have been mapped to an element designated the Minichromosome Maintenance Element or MME (Piirsoo et al., 1996). Four E2 sites are sufficient for E2mediated plasmid maintenance in yeast (Brannon et al., 2005). However, yeast undergoes a ‘‘closed mitosis’’ in which the nuclear membrane does not break down and this might result in less stringent requirements for plasmid maintenance. The herpesvirus tethering proteins, EBNA-1 and LANA, also bind to large numbers of tandem binding sites in their respective viral genomes to link them to the host chromosomes. Most other papillomaviruses contain only four consensus E2 binding sites and it is not yet clear whether this would be sufficient for stable genome maintenance using a mechanism similar to BPV-1. Fewer sites may suffice if these viruses use different chromosomal targets and mechanisms to partition their genomes. But these genomes may also contain additional elements that bind cellular or viral proteins and stabilize the tethering complex. It is likely that for many of the papillomaviruses, the E2 binding sites will not be sufficient for stable genome maintenance. One potential candidate cis elements is a Matrix attachment region (MAR). These DNA sequences anchor chromatin fibers to the nuclear matrix and generate transcriptionally active domains. MARs have been mapped in papillomavirus genomes in vitro (Tan et al., 1998). There is also precedent that these elements could be important for viral genome maintenance; plasmids containing cellular MAR elements are stably maintained in cells over many cell generations by attachment to cellular mitotic chromosomes (Baiker et al., 2000) via an interaction with the scaffold attachment protein SAF-A ( Jenke et al., 2002). Several studies have shown that sub-genomic fragments of HPV DNA can replicate autonomously in yeast in an E1 and E2 independent manner (see Section V.D. below). A recent study has shown that similar multiple sub-genomic fragments of HPV16 DNA can also replicate and/or persist in mammalian cells, in the absence of E1 and E2, further implicating the role of cellular factors in binding to viral cis elements (Pittayakhajortwut and
Replication and Partitioning of Papillomavirus Genomes
179
Angeletti, 2008). Sequences such as MAR elements, HMG, Topoisomerase II, Telomere-repeat binding factor and CENP-B binding sites have been identified in the HPV16 genome and could be potential candidates for binding to cellular genome maintenance factors (Pittayakhajortwut and Angeletti, 2008). Another potential mechanism of regulation is methylation of the E2 binding sites. E2 is unable to bind to its consensus sequence when it contains a methylated CpG dinucleotide (Kim et al., 2003; Thain et al., 1996). In replicating cells maintaining the HPV16 genome as an episome, the E2 binding sites, especially binding site 2, were often methylated. In contrast, in differentiated cells the genomes become hypomethylated (Kim et al., 2003). This may also indicate that the E2 binding sites are not crucial for genome maintenance. One of the difficulties in designing a papillomavirus segregation assay is the multifunctional nature of the E2 protein. Because E2 is also important for replication initiation and transcriptional activation, it is necessary to separate these functions to study the role of E2 in genome maintenance. Analysis of the roles of individual E2 binding sites on long-term replication of the viral genome is difficult because the sites are also required for replication initiation and for transcriptional regulation of the virus, including regulation of expression of the E1 and E2 proteins themselves. In HPV31, mutation of each E2 site, except BS2, resulted in integration of the viral genome (Stubenrauch et al., 1998b). To date, the only successful partitioning assay is for BPV1 (Abroi et al., 2004).
F. Papillomavirus chromosomal tethering targets Disruption of the chromosomal interaction of the papillomavirus E2/ genome tethering complex could have great therapeutic potential and so an important goal has been to identify and characterize the cellular chromosomal targets that mediate viral genome segregation. Table I lists chromosomal targets for a range of papillomavirus E2 proteins as well as proposed targets for other viral tethering proteins.
1. Brd4 The best characterized chromosomal target, to date, is the cellular protein, Brd4. Brd4 is a major component of the tethering complex of BPV-1, and probably a subset of other papillomaviruses (Baxter et al., 2005; McPhillips et al., 2006; You et al., 2004). Brd4 is a double bromodomain protein that binds to acetylated histone tails of histones H3 and H4 and, in many cell types, remains bound to chromatin through mitosis (Dey et al., 2003). Brd4 is a member of the BET family of double bromodomain proteins (Florence and Faller, 2001) that ‘‘read’’ the histone code by
180
Alison A. McBride
binding to acetylated residues. Brd4 has been isolated in complex with the replication factor, RFC (Maruyama et al., 2002) and as part of the mouse transcriptional Mediator complex (Jiang et al., 1998). It is also a component of P-TEFb (Jang et al., 2005) and stimulates RNA polymerase II-dependent transcriptional elongation. In many cells, Brd4 is detected as a diffuse coat around mitotic chromosomes and it has been hypothesized to play a role in epigenetic memory (Dey et al., 2003). In the presence of the BPV-1 E2 protein, both E2 and Brd4 colocalize on mitotic chromosomes in punctuate spots (McPhillips et al., 2005) and analysis of mutated E2 proteins shows that the mitotic chromosome binding activity of E2 correlates strongly with Brd4 interaction (Baxter et al., 2005). Two highly conserved residues in the transactivation domain, R37 and I73, previously shown to be crucial for transcriptional activation, are critical for Brd4 binding in all E2 proteins tested (Baxter et al., 2005; McPhillips et al., 2006; Senechal et al., 2007). In fact, interaction of E2 with Brd4 is also crucial for E2-mediated transcriptional regulation in all E2 proteins analyzed (Ilves et al., 2006; McPhillips et al., 2006; Schweiger et al., 2006; Wu et al., 2006). The dimerization of E2 through the E2 C-terminal DNA binding domain promotes efficient binding of E2 to Brd4 and to mitotic chromosomes (Cardenas-Mora et al., 2008). Furthermore, novel E2 ‘‘single chain heterodimer’’ proteins containing one transactivation domain linked to a dimeric DNA binding domain binds Brd4 much less efficiently than the wildtype E2 protein (Kurg et al., 2006). In support of this, an E2-Brd4 heterotetramer, with two Brd4 peptides sandwiched between two E2 N-terminal domains, was recently observed in a crystallographic structure of the HPV16 E2 transactivation domain in complex with a C-terminal 20 amino acid peptide from Brd4 (Abbate et al., 2006). Dimerization of the E2 protein through the C-terminal domain would greatly stabilize such a complex. Cooperative binding of E2 and Brd4 into higher order complexes could explain why the Brd4-E2 complex is observed as punctuate speckles on mitotic chromosomes (McPhillips et al., 2005), whereas in the absence of E2 it forms a diffuse coating (Dey et al., 2000). When exogenously expressed, the C-terminal E2 binding region of Brd4 interferes with the E2-Brd4 interaction, the association of E2 and viral genomes with mitotic chromosomes, BPV-1-induced cellular transformation and viral genome maintenance (You et al., 2004, 2005). E2 is unable to maintain plasmids containing E2 binding sites in Saccharomyces cerevisiae but this function can be reconstituted with Brd4 expression (Brannon et al., 2005). Thus, the E2-Brd4 association is pivotal to BPV-1 maintenance replication. However, while Brd4 is clearly an important component of the tethering complex for several papillomaviruses, it is not clear that it is the key link between the E2/genome complex and the
Replication and Partitioning of Papillomavirus Genomes
181
cellular chromosomes. Brd4 is not a stable tether, interacting only with histones that are acetylated (Dey et al., 2003) and most of which become deacetylated during mitosis (Kruhlak et al., 2001). Perhaps because of this, the interaction of E2 and Brd4 is not passive and E2 actively stabilizes the interaction of Brd4 with chromatin and redistributes it into punctate dots on the mitotic chromosomes (McPhillips et al., 2005). Thus, the E2/genome complex does not passively hitchhike on mitotic chromosomes but instead modifies and stabilizes the Brd4-chromatin interaction to ensure the stable transmission of the viral genome. It has also not been conclusively shown that disruption of the E2-Brd4 interaction directly impacts viral genome maintenance in mammalian cells. Inhibition of the E2-Brd4 interaction by either downregulation of Brd4 by siRNA, or by expression of the dominant negative CTD of Brd4 results in loss of viral genomes, but it is not clear whether this is due to inhibition of the transcriptional function of E2, which also requires Brd4, or whether it is due to a direct inhibition of chromosomal tethering (Schweiger et al., 2006; You et al., 2004, 2005). A peptide from Brd4, corresponding to the E2 interacting region, was also able to displace HPV16 genomes from mitotic chromosomes (Abbate et al., 2006), but it has yet to be shown that alpha HPV16 E2 protein binds to mitotic chromosomes in a similar pattern. Brd4 is clearly an important part of the BPV-1 E2 tethering complex but it does not seem to be involved in genome tethering of all papillomaviruses (McPhillips et al., 2006). The E2 proteins of some viruses (EEPV, ROPV, HPV-1a) show complete colocalization of E2 and Brd4 on mitotic chromosomes as is seen for BPV1, others some show partial colocalization and others show none (McPhillips et al., 2006). Furthermore, mutations in E2 that abrogate Brd4 binding (in R37 and I73 residues) do not affect mitotic chromosomal localization for E2 proteins from viruses such as HPV8 and HPV31 (McPhillips et al., 2006). Consistent with these findings, an HPV31 genome containing a mutation in E2 residue 73 (I73L), which disrupts the interaction of E2 and Brd4 (Senechal et al., 2007), is maintained as a stable episome and can be amplified in differentiated keratinocytes (Stubenrauch et al., 1998a). In CRPV, however, E2 mutations that abrogate the Brd4 interaction have a detrimental effect on the induction of tumors (a domestic rabbit model resulting in abortive infection) suggesting that in CRPV the E2-Brd4 interaction is important for some aspects of pathogenesis (Jeckel et al., 2002). Notably, the CRPV E2 protein interacts with Brd4 much more efficiently that the alpha papillomavirus E2 proteins and so may have a more significant role in the CRPV life cycle (McPhillips et al., 2006). In a different study, HPV11 E2 was observed bound to mitotic microtubules but also did not colocalize with Brd4 (Dao et al., 2006). Nevertheless, all E2 proteins tested interact with Brd4, albeit with different binding efficiencies. This is because Brd4 is required
182
Alison A. McBride
for E2-mediated transcription, but not genome partitioning, of all papillomaviruses.
2. rDNA Loci
Oliveira et al., observed that certain E2 proteins, as exemplified by HPV-8, associate with the pericentromeric region of mitotic chromosomes rather than being distributed over the chromosome arms (Oliveira et al., 2006). Further investigation has shown the peri-centromeric binding target to be the ribosomal RNA gene loci on the short arms of acrocentric chromosomes (Poddar et al., 2008). This interaction does not require the Brd4 protein and is an attractive target for chromosomal tethering for several reasons. The tandemly-arranged repeating rDNA units (400 per diploid cell) greatly increase the local concentration of the E2 binding target. Also, the rDNA loci have specialized mechanisms for replication, cohesion, condensation and segregation (D’Amours et al., 2004; Freeman et al., 2000; Kobayashi et al., 1998; Sullivan et al., 2004; Torres-Rosell et al., 2005) and RNA polymerase I transcription factors remain bound throughout mitosis (McStay, 2006). The resulting, unique chromatin structure could provide unique targets on the host chromosomes. Furthermore, the rDNA genes are sequestered in the nucleoli in interphase, which would leave the E2 protein free to participate in viral transcription and replication until the nucleolar envelope breaks down in early mitosis. Notably, the cellular target of the Epstein-Barr virus tethering protein, EBNA-1 is a protein termed p40 nucleolar proliferation antigen, or hEBP2, and is involved in pre-RNA processing (Shire et al., 1999). hEBP2 is nucleolar in interphase (Chatterjee et al., 1987) but is distributed along condensed chromosomes in mitosis (Wu et al., 2000). The C-terminal domain of the LANA tethering protein of HHV-8 has also been shown to bind to pericentromeric and telomeric regions of certain metaphase chromosomes (Kelley-Clarke et al., 2007), some of which the authors speculate might be acrocentric chromosomes. In certain conditions, the E2 proteins from the alpha-genus, are also observed binding to pericentromeric regions of chromosomes (Oliveira et al., 2006). Thus, the rDNA loci might be an attractive tethering target for several papillomaviruses.
3. ChLR1 Another cellular protein that is involved in papillomavirus genome partitioning is ChLR1, a DNA helicase involved in sister chromatid cohesion (Parish et al., 2006). E2 interacts with and colocalizes with ChLR1, but only at early stages of mitosis. Furthermore, BPV-1 genomes that encode an E2 protein which is unable to interact with ChLR1 are not maintained extrachromosomally and downregulation of hChLR1 results in viral genome loss. Therefore, hCHLR1 may be important for loading, but not maintaining, the E2/genome complex on mitotic chromosomes.
Replication and Partitioning of Papillomavirus Genomes
183
4. Mitotic spindle In a different study, several of the alpha-papillomavirus E2 proteins have been observed to associate with the mitotic spindle (Van Tine et al., 2004). Both the N-terminal and C-terminal domains of HPV-11 E2 can associate with the microtubules and a short sequence in the DNA binding domain important for this interaction has been mapped (Dao et al., 2006). Notably, this region is not well conserved in BPV-1 E2. The E2 proteins have also been observed to bind to the centrosome (Donaldson et al., 2007; Van Tine et al., 2004). Whether this localization reflects the tethering function of E2, or the site of its proteasomal degradation, (Fabunmi et al., 2000) remains to be determined.
G. Replication licensing Maintenance papillomavirus DNA replication is coupled to that of the host cell and, on average, each genome is replicated once per cell cycle to give an overall constant copy number. This copy number could be maintained if replication depends on the availability of a limiting factor; genomes would replicate randomly and replication would cease when this factor is exhausted. An alternative scenario is that each genome is licensed and is only replicated once per cell cycle. Early studies indicated that BPV-1 was subject to replication licensing in mouse cells (Botchan et al., 1986), but further studies showed that when cells were isolated after S-phase a portion of the genomes had undergone multiple rounds of replication indicating a random choice mechanism (Gilbert and Cohen, 1987; Ravnan et al., 1992). A more recent study analyzed the mode of replication of the alpha papillomaviruses, HPV-16 and HPV-31. However, the results were mixed in that the different genomes seemed to have different modes of replication and this could change in different cell types (Hoffmann et al., 2006). One hypothesis is that the mode of replication is actually determined by the host cell and that lines derived from infection of transit amplifying cells give rise to random choice viral replication whereas infection of keratinocyte stem cells results in genomes that undergo replication licensing (Hoffmann et al., 2006). The genomes of Epstein-Barr virus (EBV) are also maintained as episomes and each genome is replicated in an ordered once per cell cycle fashion (Yates and Guan, 1991). However, latent EBV replication is initiated by the cellular ORC and MCM proteins, which are normally licensed in each cell cycle. Since papillomaviruses encode their own initiator protein, it is less likely that E1-dependent replication would be subject to licensing.
H. Regulation of genome copy number and partitioning It is not well understood how papillomavirus genome copy number is maintained and regulated. For BPV-1, E2 protein levels correlate with genome copy number and greatly increased amounts of E2 protein and
184
Alison A. McBride
viral genomes are observed tethered to the host chromosomes when the half-life of the E2 protein is increased (Penrose and McBride, 2000; Skiadopoulos and McBride, 1998). Mutation of a CK2 phosphorylation site in the BPV-1 E2 protein greatly reduces protein degradation by the proteasome and results in a protein with a much longer half-life and a virus with greatly increased genome copy number (McBride and Howley, 1991). However, the E2 CK2 phosphorylation site is conserved only among the delta papillomaviruses, which cause fibropapillomas. Nevertheless, the levels of all E2 proteins are observed to dramatically decrease in mitosis (Bellanger et al., 2001). Since the E2-genome complex must be tethered to the host chromosomes throughout the length of mitosis, it is very likely that these processes are tightly regulated during persistent infection. Expression of the E1 protein of BPV-1 interferes with E2 proteinmediated tethering of the viral DNA to mitotic chromosomes, introducing yet another level of regulation to papillomavirus genome partitioning (Voitenleitner and Botchan, 2002). The truncated E2 repressor proteins can also regulate genome copy number and may be important to limit runaway replication. BPV-1 containing mutations that eliminate expression of the E2-TR protein replicate at greatly increased copy number (Lambert et al., 1990; Riese et al., 1990). In HPV31, the E8^E2 protein also regulates genome copy number (Zobel et al., 2003). The E2 repressor proteins contain the DNA binding and dimerization domain of E2 and could potentially regulate replication either by directly forming heterodimers with the full-length E2 protein or by competing for binding to sites in the viral genome (Lim et al., 1998).
IV. VEGETATIVE REPLICATION The third stage of viral replication is vegetative DNA replication, where viral genomes are amplified to a high copy number destined to be packaged in the capsids of progeny virions. Vegetative DNA replication occurs only in the differentiating layers of a papilloma and less is known about this mode of replication. Vegetative DNA replication will take place in organotypic raft cultures and in xenografts of mice (McBride et al., 2000; Meyers et al., 1992), but usually requires that genomes are maintained extrachromosomally at earlier stages of the infection. This makes it difficult to separate the requirements for the three modes of replication. By studying replicative intermediates, some studies have reported that there is a switch from bidirectional theta replication in the maintenance stage to a rolling circle mode in the vegetative stage (Dasgupta et al., 1992; Flores and Lambert, 1997). The rolling circle mechanism is an efficient
Replication and Partitioning of Papillomavirus Genomes
185
method for generating large amounts of viral DNA; for example, bacteriophage lambda replicates in a bidirectional theta mode early in infection, and then switches to the sigma rolling circle mode at late times. This leads to long concatamers of DNA that are cut into unit genome lengths for packaging (reviewed by (Narajczyk et al., 2007)). Similarly, herpesviruses such as Epstein Barr virus have two modes of replication and initiation switches from the latent ‘‘plasmid’’ origin (oriP) to the lytic origin (oriLyt) for vegetative replication (Hammerschmidt and Sugden, 1988). Vegetative replication of EBV yields intermediates of large head to tail concatemeric molecules, which are subsequently cleaved and packaged as described for lambda, and it is assumed that they are generated by a rolling circle mechanism (reviewed in (Tsurumi et al., 2005). However, initiator proteins involved in rolling circle replication usually have an associated nuclease activity and rolling circle replication (RCR) motifs (Ilyina and Koonin, 1992). Notably, the Rep initiator protein of Adeno-Associated virus (AAV), which initiates rolling circle replication, has a DNA binding domain structure very related to the SV40 T antigen and the papillomavirus E1 protein (Hickman et al., 2004). However, while the RCR motif regions of the latter two proteins are structurally homologous to the analogous region in Rep, the catalytic RCR residues have not been conserved. Therefore, it remains unclear whether papillomaviruses switch to a different replication mechanism for vegetative replication. Vegetative replication is triggered in differentiated cells and this may be due to an increase in the levels of the E1 and E2 replication proteins. Certainly in BPV-1 infected papillomas, cells undergoing genome amplification contain greatly increased levels of the E2 protein (Burnett et al., 1990; Penrose and McBride, 2000) and BPV-1 containing cells which spontaneously amplify the viral genome also contain high levels of E2 (Burnett et al., 1990). An example of this colocalization is shown in Fig. 9. To date, immunological detection of the E1 protein has not been possible but mRNA species that are predicted to express both the E1 and E2 proteins are induced at this stage of infection in HPV31 (Ozbun and Meyers, 1998). E1 transcripts are expressed from both early and late promoters, but only the early promoter is repressed by the E2 protein. Therefore, a switch of transcription to the late promoter would allow E1 to be expressed to high levels that were not possible from genomes being maintained in the basal cells. Moody et al., recently demonstrated that HPV infection could specifically activate caspases upon differentiation and, in turn, the caspases cleaved the E1 protein in the N-terminal domain at a highly conserved DxxD motif located between residues 46 and 49 (Moody et al., 2007). Mutation of this motif inhibited genome amplification indicating that E1 cleavage might also be important for regulating vegetative replication.
186
Alison A. McBride
Cellular DNA
E2 protein
Merge
Cellular DNA
Viral DNA
Merge
FIGURE 9 Vegetative DNA amplification in a BPV-1 Papilloma. E2-specific immunofluorescence and BPV DNA-specific fluorescent in situ hybridization were performed on serial sections of bovine wart tissue. E2 protein expression is shown in green in the top panels and viral DNA is shown in green in the bottom panels. In both cases cellular DNA is counterstained in red. Cells containing high levels of both E2 protein and viral DNA are indicate by arrows.
V. OTHER ASPECTS OF PAPILLOMAVIRUS REPLICATION A. Establishment of a replication competent environment For all modes of replication, the papillomaviruses rely on the host replication machinery. Initial genome amplification and maintenance takes place in the cycling basal cells and so the virus has available the host DNA replication factors it requires. Nevertheless, the regulatory activities of the E5, E6 and E7 proteins are likely to enhance replication, both directly and indirectly. Both the E6 and E7 proteins of HPV-16 and HPV-31 are required for long term maintenance of the viral genome (Park and Androphy, 2002; Thomas et al., 1999). E6 and E7 could indirectly enhance replication by promoting cellular proliferation and division but they could also interfere with inhibitory factors such as TBP, CDP, YY1 and p53 (Hartley and Alexander, 2002; Ilves et al., 2003; Narahari et al., 2006).
Replication and Partitioning of Papillomavirus Genomes
187
The cell cycle regulatory functions of the E6 and E7 proteins are absolutely required for vegetative amplification in differentiated cells as these cells would normally have withdrawn from the cell cycle and the E6 and E7 proteins are required to maintain or induce an pseudo S-phase like state so that the cellular replication machinery is available. In addition, the expression of the E1^E4 protein can cause a G2 arrest and so expression of both proteins in the same cell results in an pseudo S-phase like environment conducive to viral genome amplification. At high expression levels, which is also observed in cells amplifying the genome, E2 can also result in G2 arrest (Frattini et al., 1997). This cell cycle arrest could allow sustained synthesis of viral DNA and be important for vegetative replication.
B. Differences in replication strategies of different papillomaviruses Papillomaviruses have coevolved with their hosts over millions of years and individual HPV types have existed since the evolutionary origin of humans (Bernard et al., 2006). Each virus is epitheliotropic but infects either mucosal or cutaneous epithelia, usually at a specific site of the body. Although each virus has a very similar genomic organization and the viral proteins are relatively well conserved, infection by different papillomaviruses results in a wide spectrum of pathologies. These can range from clinically inapparent infections, through a diverse variety of benign papillomas and warts, to a subset of infections that can progress to malignant carcinomas. These pathological differences are most likely due to different viral types, different epithelial host cells and the immune response of the host. However, in each case the virus has a similar strategy in that it must establish a persistent infection in the dividing cells of the epithelium and restrict vegetative growth to the more differentiated cells that are destined to be sloughed from the epithelium. The timing of productive replication, however, differs among the papillomaviruses. Some, such as the cutaneous mu viruses, trigger vegetative viral DNA amplification as soon as the host cells leave the basal layer while others, such as the mucosal alpha viruses, postpone productive replication until later stages of differentiation (Peh et al., 2002). Similarly, it is becoming apparent that different papillomaviruses use different chromosomal targets for partitioning the viral genomes and the chromosome binding pattern of E2 seems to be characteristic of each evolutionary group. Notably, all E2 proteins studied to date interact with the Brd4 protein to regulate viral transcription (McPhillips et al., 2006; Schweiger et al., 2006), but only a subset of viruses also use Brd4 as a chromosomal tether in mitosis (McPhillips et al., 2006). For regulation of transcription and initiation of DNA replication, the E2 proteins must
188
Alison A. McBride
interact precisely with many cellular protein complexes that tightly regulate these processes. For genome partitioning, the E2 protein must simply tether the viral genome to a stable chromosomal target. This could be a cellular protein such as Brd4, which is also used for other E2 functions, or could be a completely different target used only for mitotic partitioning.
C. Comparison of papillomavirus DNA replication with cellular DNA replication and replication of other viruses The replicon model of Jacob and Brenner first proposed that replication was triggered by an initiator protein binding to a genetic element designated the replicator ( Jacob and Brenner, 1963). In higher eukaryotes, the initiator is actually a complex of proteins that is assembled on the replicator in a regulated manner. The replicator element is recognized by ORC, the origin recognition complex, followed by Cdt1 and CDC6, which function to recruit and load the helicase, MCM. The assembled MCM complex must be activated by cyclin dependent kinases (reviewed in (Masai et al., 2005). While many viruses utilize the cellular replication machinery, they also often encode their own initator proteins. For papillomaviruses, the specific DNA binding domain of E1 is analogous to ORC and the helicase function is analogous to the cellular helicase MCM. The loading functions of Cdt1 and CDC6 are carried out by the E2 protein. Other viruses also encode initiator proteins. The large T antigen of SV40 initiates replication in a very similar manner to E1. Both proteins have limited sequence homology (Mansky et al., 1997) but extensive structural homology in the helicase and DNA binding domains (Enemark and Joshua-Tor, 2006; Enemark et al., 2000; Li et al., 2003; Luo et al., 1996; Meinke et al., 2006, 2007). However, SV40 has no counterpart to E2 and the double hexameric helicase of T antigen assembles onto the replication origin in the absence of a cellular or viral loading factor (Borowiec et al., 1990). In permissive cells, SV40 is a lytic virus that undergoes runaway replication and does not establish its genome as a persistent episome. Therefore, the role of E2 in papillomavirus regulation may be to regulate runaway replication as well as to maintain and segregate the viral genome. Another virus, Epstein-Barr virus (EBV), establishes long-term persistent infections similar to the papillomaviruses. EBV replicates in synchrony with the cellular DNA and does not encode a viral helicase initiator protein (Kirchmaier and Sugden, 1998; Yates and Guan, 1991). Instead, replication is regulated by assembly of ORC, MCM and other cellular replication factors (Chaudhuri et al., 2001; Schepers et al., 2001). EBV encodes a single protein, EBNA-1, that is required for maintenance replication and segregation of the viral genome. EBNA-1, binds to the latent replication origin, oriP (Rawlins et al., 1985; Yates et al., 1985), which contains a family of repeats (FR) and a dyad symmetry (DS) element.
Replication and Partitioning of Papillomavirus Genomes
189
The FR element is essential for stable episomal maintenance and contains multiple EBNA-1 binding sites. It functions in concert with the DS element, which contains phased EBNA1 sites adjacent to telomere repeat factor (TRF) binding sites, and together they function as an efficient origin of DNA replication initiation (Deng et al., 2002, 2003).Thus the function of EBNA is analogous to that of the E2 protein.
D. Papillomavirus replication in Saccharomyces cerevisiae There are many genetic systems and screens in yeast that enable the study of plasmid maintenance in a much more practical and efficient manner than in mammalian cells. Several studies have shown that papillomavirus genomes will replicate stably in yeast (Angeletti et al., 2002; Chattopadhyay et al., 2005; Kim et al., 2005). Papillomavirus genomes, and sub-genomic fragments, are replicated and maintained through sequences that can substitute for autonomously replicating sequence (ARS) and centromere (CEN) elements, even in the absence of the E1 and E2 proteins. Although this does not directly parallel the requirements for papillomavirus replication in mammalian cells, the elements identified in these studies may be important for viral genome replication in addition to E1 and E2-dependent elements. In a different type of study, yeast plasmids containing an ARS element and E2 binding sites could be maintained in yeast in the presence of the BPV-1 E2 protein and the mammalian Brd4 protein (Brannon et al., 2005). Notably, S. cerevisiae do express a Brd4 family member, Bdf1, but this is not sufficient for E2-dependent maintenance. Bdf1 does not contain a region homologous to the C-terminal region of Brd4 that interacts with E2, but a fusion protein of Bdf1-Brd4 can support E2-plasmid maintenance.
E. Anti-viral replication therapies Many antiviral therapies for HPV-associated disease have focused on the replication proteins (reviewed in Fradet-Turcotte and Archambault, 2007). Small molecule indandione inhibitors, designed to inhibit interaction between the N-terminal domain of E2 and the helicase domain of E1, inhibit HPV-11 replication (White et al., 2003). Polyamides have been designed to inhibit specific E1 and E2 DNA binding function (Schaal et al., 2003). Another attractive target is the enzymatic activities of the E1 ATPase/helicase and several pharmaceutical companies have candidates for these inhibitors (reviewed in Fradet-Turcotte and Archambault, 2007). There is also great interest in designing therapeutics that would disrupt the tethering of the viral genome to mitotic chromosomes, which is predicted to cure persistent papillomavirus infection.
190
Alison A. McBride
F. Papillomavirus-based vectors Gene therapy vectors that replicate extrachromosomally are advantageous compared to those that integrate randomly into the host genome as they will not result in insertional mutagenesis and are not susceptible to positional effects, yet will persist long term in dividing cells. One of the first descriptions of the concept of viral based vectors was described by Rogers in 1966 in a paper entitled ‘‘Shope Papillomavirus: a passenger in man and its significance to the potential control of the host genome’’ (Friedmann, 2001; Rogers, 1966). However, the actual use of papillomavirus-based vectors was not realized until the 1980s (Sarver et al., 1981). The first vectors consisted of the early region of BPV-1 linked to an expression cassette for the foreign gene. However, these early vectors were not widely adopted because they transformed cells, had a limited host range and transcription of the foreign gene often caused the plasmid to integrate (Waldenstrom et al., 1992). This was likely due to interference between the viral regulatory elements and the insertion of a heterologous enhancer and promoter sequences in the compact papillomavirus genomes. More recently vectors have been developed based on the modular elements required for replication. For example, newer BPV-1 based vectors contain only the LCR and the E1 and E2 genes (Mannik et al., 2002; Ohe et al., 1995) and are maintained in transgenic mice for several generations (Mannik et al., 2003). Human papillomavirus-based vectors have also been explored and shown to be able to replicate, at least transiently, in human cell lines and in the lungs of mice (Gadi et al., 1999; Sverdrup et al., 1999). However, at this point these studies have shown mainly ‘‘proof of principle’’ of papillomavirus based vectors and they have not yet been widely or practically used to express foreign genes. More detailed knowledge of the mechanisms of human papillomavirus replication and genome maintenance should enable the development of more successful vectors.
ACKNOWLEDGMENTS The author’s research is supported by the Intramural Research Program of the NIH, NIAID.
REFERENCES Abbate, E. A., Berger, J. M., and Botchan, M. R. (2004). The X-ray structure of the papillomavirus helicase in complex with its molecular matchmaker E2. Genes Dev. 18:1981–1996. Abbate, E. A., Voitenleitner, C., and Botchan, M. R. (2006). Structure of the papillomavirus DNA-tethering complex E2:Brd4 and a peptide that ablates HPV chromosomal association. Mol. Cell 24:877–889. Abroi, A., Kurg, R., and Ustav, M. (1996). Transcriptional and replicational activation functions in the BPV1 E2 protein are encoded by different structural determinants. J. Virol. 70:6169–6179.
Replication and Partitioning of Papillomavirus Genomes
191
Abroi, A., Ilves, I., Kivi, S., and Ustav, M. (2004). Analysis of chromatin attachment and partitioning functions of bovine papillomavirus type 1 E2 protein. J. Virol. 78:2100–2113. Ai, W., Toussaint, E., and Roman, A. (1999). CCAAT displacement protein binds to and negatively regulates human papillomavirus type 6 E6, E7, and E1 promoters. J. Virol. 73:4220–4229. Akgul, B., Karle, P., Adam, M., Fuchs, P. G., and Pfister, H. J. (2003). Dual role of tumor suppressor p53 in regulation of DNA replication and oncogene E6-promoter activity of epidermodysplasia verruciformis-associated human papillomavirus type 8. Virology 308:279–290. Androphy, E. J., Lowy, D. R., and Schiller, J. T. (1987). Bovine papillomavirus E2 trans-activating gene product binds to specific sites in papillomavirus DNA. Nature 325:70–73. Angeletti, P. C., Kim, K., Fernandes, F. J., and Lambert, P. F. (2002). Stable replication of papillomavirus genomes in Saccharomyces cerevisiae. J. Virol. 76:3350–3358. Antson, A. A., Burns, J. E., Moroz, O. V., Scott, D. J., Sanders, C. M., Bronstein, I. B., Dodson, G. G., Wilson, K. S., and Maitland, N. J. (2000). Structure of the intact transactivation domain of the human papillomavirus E2 protein. Nature 403:805–809. Ashrafi, G. H., Brown, D. R., Fife, K. H., and Campo, M. S. (2006). Down-regulation of MHC class I is a property common to papillomavirus E5 proteins. Virus Res. 120:208–211. Auster, A. S., and Joshua-Tor, L. (2004). The DNA-binding domain of human papillomavirus type 18 E1. Crystal structure, dimerization, and DNA binding. J. Biol. Chem. 279:3733–3742. Baiker, A., Maercker, C., Piechaczek, C., Schmidt, S. B., Bode, J., Benham, C., and Lipps, H. J. (2000). Mitotic stability of an episomal vector containing a human scaffold/matrixattached region is provided by association with nuclear matrix. Nat. Cell Biol. 2:182–184. Barbera, A. J., Chodaparambil, J. V., Kelley-Clarke, B., Joukov, V., Walter, J. C., Luger, K., and Kaye, K. M. (2006). The nucleosomal surface as a docking station for Kaposi’s sarcoma herpesvirus LANA. Science 311:856–861. Barsoum, J., Prakash, S. S., Han, P., and Androphy, E. J. (1992). Mechanism of action of the papillomavirus E2 repressor: Repression in the absence of DNA binding. J. Virol. 66:3941–3945. Bastien, N., and McBride, A. A. (2000). Interaction of the papillomavirus E2 with mitotic chromosomes. Virology 270:124–134. Baxter, M. K., McPhillips, M. G., Ozato, K., and McBride, A. A. (2005). The mitotic chromosome binding activity of the papillomavirus E2 protein correlates with interaction with the cellular chromosomal protein, Brd4. J. Virol. 79:4806–4818. Bedell, M. A., Hudson, J. B., Golub, T. R., Turyk, M. E., Hosken, M., Wilbanks, G. D., and Laimins, L. A. (1991). Amplification of human papillomavirus genomes in vitro is dependent on epithelial differentiation. J. Virol. 65:2254–2260. Bedrosian, C. L., and Bastia, D. (1990). The DNA-binding domain of HPV-16 E2 protein interaction with the viral enhancer: Protein-induced DNA bending and role of the nonconserved core sequence in binding site affinity. Virology 174:557–575. Bellanger, S., Demeret, C., Goyat, S., and Thierry, F. (2001). Stability of the human papillomavirus type 18 E2 protein is regulated by a proteasome degradation pathway through its amino-terminal transactivation domain. J. Virol. 75:7244–7251. Berg, L., Lusky, M., Stenlund, A., and Botchan, M. R. (1986). Repression of bovine papilloma virus replication is mediated by a virally encoded trans-acting factor. Cell 46:753–762. Bernard, B. A., Bailly, C., Lenoir, M. C., Darmon, M., Thierry, F., and Yaniv, M. (1989). The human papillomavirus type 18 (HPV18) E2 gene product is a repressor of the HPV18 regulatory region in human keratinocytes. J. Virol. 63:4317–4324.
192
Alison A. McBride
Bernard, H. U., Calleja-Macias, I. E., and Dunn, S. T. (2006). Genome variation of human papillomavirus types: Phylogenetic and medical implications. Int. J. Cancer 118:1071–1076. Bian, X. L., Rosas-Acosta, G., Wu, Y. C., and Wilson, V. G. (2007). Nuclear import of bovine papillomavirus type 1 E1 protein is mediated by multiple alpha importins and is negatively regulated by phosphorylation near a nuclear localization signal. J. Virol. 81:2899–2908. Bochkarev, A., Barwell, J. A., Pfuetzner, R. A., Furey, W. J., Edwards, A. M., and Frappier, L. (1995). Crystal structure of the DNA-binding domain of the Epstein-Barr virus originbinding protein EBNA 1. Cell 83:39–46. Borowiec, J. A., Dean, F. B., Bullock, P. A., and Hurwitz, J. (1990). Binding and unwinding– how T antigen engages the SV40 origin of DNA replication. Cell 60:181–184. Botchan, M., Berg, L., Reynolds, J., and Lusky, M. (1986). The bovine papillomavirus replicon. Ciba Found. Symp. 120:53–67. Bousarghin, L., Touze, A., Sizaret, P. Y., and Coursaget, P. (2003). Human papillomavirus types 16, 31, and 58 use different endocytosis pathways to enter cells. J. Virol. 77:3846–3850. Brannon, A. R., Maresca, J. A., Boeke, J. D., Basrai, M. A., and McBride, A. A. (2005). Reconstitution of papillomavirus E2-mediated plasmid maintenance in Saccharomyces cerevisiae by the Brd4 bromodomain protein. Proc. Natl. Acad. Sci. USA 102:2998–3003. Brokaw, J. L., Blanco, M., and McBride, A. A. (1996). Amino acids critical for the functions of the bovine papillomavirus type 1 E2 transactivator. J. Virol. 70:23–29. Brown, C., Kowalczyk, A. M., Taylor, E. R., Morgan, I. M., and Gaston, K. (2008). p53 represses human papillomavirus type 16 DNA replication via the viral E2 protein. Virol. J. 5:5. Brown, D. R., Kitchin, D., Qadadri, B., Neptune, N., Batteiger, T., and Ermel, A. (2006). The human papillomavirus type 11 E1–E4 protein is a transglutaminase 3 substrate and induces abnormalities of the cornified cell envelope. Virology 345:290–298. Burnett, S., Strom, A. C., Jareborg, N., Alderborn, A., Dillner, J., Moreno-Lopez, J., Pettersson, U., and Kiessling, U. (1990). Induction of bovine papillomavirus E2 gene expression and early region transcription by cell growth arrest: Correlation with viral DNA amplification and evidence for differential promoter induction. J. Virol. 64:5529–5541. Calderwood, M. A., Hall, K. T., Matthews, D. A., and Whitehouse, A. (2004). The herpesvirus saimiri ORF73 gene product interacts with host-cell mitotic chromosomes and selfassociates via its C terminus. J. Gen. Virol. 85:147–153. Cardenas-Mora, J., Spindler, J. E., Jang, M. K., and McBride, A. A. (2008). Dimerization of the Papillomavirus E2 Protein is Required for Efficient Mitotic Chromosome Association and Brd4 Binding. J. Virol. 82:7298–7305. Castella, S., Bingham, G., and Sanders, C. M. (2006). Common determinants in DNA melting and helicase-catalysed DNA unwinding by papillomavirus replication protein E1. Nucleic Acids Res. 34:3008–3019. Chatterjee, A., Freeman, J. W., and Busch, H. (1987). Identification and partial characterization of a Mr 40,000 nucleolar antigen associated with cell proliferation. Cancer Res. 47:1123–1129. Chattopadhyay, A., Schmidt, M. C., and Khan, S. A. (2005). Identification of a 450-bp region of human papillomavirus type 1 that promotes episomal replication in Saccharomyces cerevisiae. Virology 340:133–142. Chaudhuri, B., Xu, H., Todorov, I., Dutta, A., and Yates, J. L. (2001). Human DNA replication initiation factors, ORC and MCM, associate with oriP of Epstein-Barr virus. Proc. Natl. Acad. Sci. USA 98:10085–10089.
Replication and Partitioning of Papillomavirus Genomes
193
Chen, G., and Stenlund, A. (1998). Characterization of the DNA-binding domain of the bovine papillomavirus replication initiator E1. J. Virol. 72:2567–2576. Chen, G., and Stenlund, A. (2001). The E1 initiator recognizes multiple overlapping sites in the papillomavirus origin of DNA replication. J. Virol. 75:292–302. Chiang, C. M., Dong, G., Broker, T. R., and Chow, L. T. (1992). Control of human papillomavirus type 11 origin of replication by the E2 family of transcription regulatory proteins. J. Virol. 66:5224–5231. Choe, J., Vaillancourt, P., Stenlund, A., and Botchan, M. (1989). Bovine papillomavirus type 1 encodes two forms of a transcriptional repressor: Structural and functional analysis of new viral cDNAs. J. Virol. 63:1743–1755. Clower, R. V., Fisk, J. C., and Melendy, T. (2006). Papillomavirus E1 protein binds to and stimulates human topoisomerase I. J. Virol. 80:1584–1587. Conger, K. L., Liu, J. S., Kuo, S. R., Chow, L. T., and Wang, T. S. (1999). Human papillomavirus DNA replication. Interactions between the viral E1 protein and two subunits of human dna polymerase alpha/primase. J. Biol. Chem. 274:2696–2705. Cotter, M. A., and Robertson, E. S. (1999). The latency-associated nuclear antigen tethers the Kaposi’s sarcoma- associated herpesvirus genome to host chromosomes in body cavitybased lymphoma cells. Virology 264:254–264. Cripe, T. P., Haugen, T. H., Turk, J. P., Tabatabai, F., Schmid, P. G., 3d., Du¨rst, M., Gissmann, L., Roman, A., and Turek, L. P. (1987). Transcriptional regulation of the human papillomavirus- 16 E6-E7 promoter by a keratinocyte-dependent enhancer, and by viral E2 trans-activator and repressor gene products: Implications for cervical carcinogenesis. EMBO J 6:3745–3753. Cueille, N., Nougarede, R., Mechali, F., Philippe, M., and Bonne-Andrea, C. (1998). Functional interaction between the bovine papillomavirus virus type 1 replicative helicase E1 and cyclin E-Cdk2. J. Virol. 72:7255–7262. Culp, T. D., Budgeon, L. R., Marinkovich, M. P., Meneguzzi, G., and Christensen, N. D. (2006). Keratinocyte-secreted laminin 5 can function as a transient receptor for human papillomaviruses by binding virions and transferring them to adjacent cells. J. Virol. 80:8940–8950. D’Amours, D., Stegmeier, F., and Amon, A. (2004). Cdc14 and condensin control the dissolution of cohesin-independent chromosome linkages at repeated DNA. Cell 117:455–469. Dao, L. D., Duffy, A., Van Tine, B. A., Wu, S. Y., Chiang, C. M., Broker, T. R., and Chow, L. T. (2006). Dynamic localization of the human papillomavirus type 11 origin binding protein E2 through mitosis while in association with the spindle apparatus. J. Virol. 80:4792–4800. Dasgupta, S., Zabielski, J., Simonsson, M., and Burnett, S. (1992). Rolling-circle replication of a high-copy BPV-1 plasmid. J. Mol. Biol. 228:1–6. Davy, C. E., Jackson, D. J., Wang, Q., Raj, K., Masterson, P. J., Fenner, N. F., Southern, S., Cuthill, S., Millar, J. B., and Doorbar, J. (2002). Identification of a G(2) arrest domain in the E1 wedge E4 protein of human papillomavirus type 16. J. Virol. 76:9806–9818. Day, P. M., Baker, C. C., Lowy, D. R., and Schiller, J. T. (2004). Establishment of papillomavirus infection is enhanced by promyelocytic leukemia protein (PML) expression. Proc. Natl. Acad. Sci. USA 101:14252–14257. Day, P. M., Lowy, D. R., and Schiller, J. T. (2003). Papillomaviruses infect cells via a clathrindependent pathway. Virology 307:1–11. Day, P. M., Roden, R. B., Lowy, D. R., and Schiller, J. T. (1998). The papillomavirus minor capsid protein, L2, induces localization of the major capsid protein, L1, and the viral transcription/replication protein, E2, to PML oncogenic domains. J. Virol. 72:142–150.
194
Alison A. McBride
DeMasi, J., Huh, K. W., Nakatani, Y., Munger, K., and Howley, P. M. (2005). Bovine papillomavirus E7 transformation function correlates with cellular p600 protein binding. Proc. Natl. Acad. Sci. USA 102:11486–11491. Deng, W., Lin, B. Y., Jin, G., Wheeler, C. G., Ma, T., Harper, J. W., Broker, T. R., and Chow, L. T. (2004). Cyclin/CDK regulates the nucleocytoplasmic localization of the human papillomavirus E1 DNA helicase. J. Virol. 78:13954–13965. Deng, Z., Atanasiu, C., Burg, J. S., Broccoli, D., and Lieberman, P. M. (2003). Telomere repeat binding factors TRF1, TRF2, and hRAP1 modulate replication of Epstein-Barr virus OriP. J. Virol. 77:11992–12001. Deng, Z., Lezina, L., Chen, C. J., Shtivelband, S., So, W., and Lieberman, P. M. (2002). Telomeric proteins regulate episomal maintenance of Epstein-Barr virus origin of plasmid replication. Mol. Cell 9:493–503. Dey, A., Chitsaz, F., Abbasi, A., Misteli, T., and Ozato, K. (2003). The double bromodomain protein Brd4 binds to acetylated chromatin during interphase and mitosis. Proc. Natl. Acad. Sci. USA 100:8758–8763. Dey, A., Ellenberg, J., Farina, A., Coleman, A. E., Maruyama, T., Sciortino, S., LippincottSchwartz, J., and Ozato, K. (2000). A bromodomain protein, MCAP, associates with mitotic chromosomes and affects G(2)-to-M transition. Mol. Cell Biol. 20:6537–6549. DiMaio, D., and Mattoon, D. (2001). Mechanisms of cell transformation by papillomavirus E5 proteins. Oncogene 20:7866–7873. Donaldson, M. M., Boner, W., and Morgan, I. M. (2007). TopBP1 regulates human papillomavirus type 16 E2 interaction with chromatin. J. Virol. 81:4338–4342. Doorbar, J., Ely, S., Sterling, J., McLean, C., and Crawford, L. (1991). Specific interaction between HPV-16 E1-E4 and cytokeratins results in collapse of the epithelial cell intermediate filament network. Nature 352:824–827. Dowhanick, J. J., McBride, A. A., and Howley, P. M. (1995). Suppression of cellular proliferation by the papillomavirus E2 protein. J. Virol. 69:7791–7799. Enemark, E. J., Chen, G., Vaughn, D. E., Stenlund, A., and Joshua-Tor, L. (2000). Crystal structure of the DNA binding domain of the replication initiation protein E1 from papillomavirus. Mol. Cell 6:149–158. Enemark, E. J., and Joshua-Tor, L. (2006). Mechanism of DNA translocation in a replicative hexameric helicase. Nature 442:270–275. Evander, M., Frazer, I. H., Payne, E., Qi, Y. M., Hengst, K., and McMillan, N. A. (1997). Identification of the alpha6 integrin as a candidate receptor for papillomaviruses. J. Virol. 71:2449–2456. Evans, M. F., Aliesky, H. A., and Cooper, K. (2003). Optimization of biotinyl-tyramide-based in situ hybridization for sensitive background-free applications on formalin-fixed, paraffin-embedded tissue specimens. BMC.Clin. Pathol. 3:2. Fabunmi, R. P., Wigley, W. C., Thomas, P. J., and DeMartino, G. N. (2000). Activity and regulation of the centrosome-associated proteasome. J. Biol. Chem. 275:409–413. Favre, M., Breitburd, F., Croissant, O., and Orth, G. (1977). Chromatin-like structures obtained after alkaline disruption of bovine and human papillomaviruses. J. Virol. 21:1205–1209. Ferran, M. C., and McBride, A. A. (1998). Transient viral DNA replication and repression of viral transcription are supported by the C-terminal domain of the bovine papillomavirus type 1 E1 protein. J. Virol. 72:796–801. Florence, B., and Faller, D. V. (2001). You bet-cha: A novel family of transcriptional regulators. Front Biosci. 6:D1008–D1018. Flores, E. R., Allen-Hoffmann, B. L., Lee, D., Sattler, C. A., and Lambert, P. F. (1999). Establishment of the human papillomavirus type 16 (HPV-16) life cycle in an immortalized human foreskin keratinocyte cell line. Virology 262:344–354.
Replication and Partitioning of Papillomavirus Genomes
195
Flores, E. R., and Lambert, P. F. (1997). Evidence for a switch in the mode of human papillomavirus type 16 DNA replication during the viral life cycle. J. Virol. 71:7167–7179. Fradet-Turcotte, A., and Archambault, J. (2007). Recent advances in the search for antiviral agents against human papillomaviruses. Antivir. Ther. 12:431–451. Frattini, M. G., Hurst, S. D., Lim, H. B., Swaminathan, S., and Laimins, L. A. (1997). Abrogation of a mitotic checkpoint by E2 proteins from oncogenic human papillomaviruses correlates with increased turnover of the p53 tumor suppressor protein. EMBO J. 16:318–331. Frattini, M. G., Lim, H. B., and Laimins, L. A. (1996). In vitro synthesis of oncogenic human papillomaviruses requires episomal genomes for differentiation-dependent late expression. Proc. Natl. Acad. Sci. USA 93:3062–3067. Freeman, L., Ragon-Alcaide, L., and Strunnikov, A. (2000). The condensin complex governs chromosome condensation and mitotic transmission of rDNA. J. Cell Biol. 149:811–824. Friedmann, T. (2001). Stanfield Rogers: Insights into virus vectors and failure of an early gene therapy model. Mol. Ther. 4:285–288. Gadi, V. K., Zou, N., Liu, J. S., Cheng, S., Broker, T. R., Sorscher, E. J., and Chow, L. T. (1999). Components of human papillomavirus that activate transcription and support plasmid replication in human airway cells. Am. J. Respir. Cell Mol. Biol. 20:1001–1006. Gammoh, N., Grm, H. S., Massimi, P., and Banks, L. (2006). Regulation of human papillomavirus type 16 E7 activity through direct protein interaction with the E2 transcriptional activator. J. Virol. 80:1787–1797. Gilbert, D. M. (2004). In search of the holy replicator. Nat. Rev. Mol. Cell Biol. 5:848–855. Gilbert, D. M., and Cohen, S. N. (1987). Bovine papilloma virus plasmids replicate randomly in mouse fibroblasts throughout S phase of the cell cycle. Cell 50:59–68. Gillison, M. L., and Lowy, D. R. (2004). A causal role for human papillomavirus in head and neck cancer. Lancet 363:1488–1489. Giroglou, T., Florin, L., Schafer, F., Streeck, R. E., and Sapp, M. (2001). Human papillomavirus infection requires cell surface heparan sulfate. J. Virol. 75:1565–1570. Goodwin, E. C., Naeger, L. K., Breiding, D. E., Androphy, E. J., and DiMaio, D. (1998). Transactivation-competent bovine papillomavirus E2 protein is specifically required for efficient repression of human papillomavirus oncogene expression and for acute growth inhibition of cervical carcinoma cell lines. J. Virol. 72:3925–3934. Gopalakrishnan, V., and Khan, S. A. (1995). E1 protein of human papillomavirus type 1a is sufficient for initiation of viral DNA replication. Proc. Natl. Acad. Sci. USA 91:9597–9601. Griffiths, R., and Whitehouse, A. (2007). Herpesvirus saimiri episomal persistence is maintained via interaction between open reading frame 73 and the cellular chromosomeassociated protein MeCP2. J. Virol. 81:4021–4032. Grundhoff, A., and Ganem, D. (2003). The latency-associated nuclear antigen of Kaposi’s sarcoma-associated herpesvirus permits replication of terminal repeat-containing plasmids. J. Virol. 77:2779–2783. Hammerschmidt, W., and Sugden, B. (1988). Identification and characterization of oriLyt, a lytic origin of DNA replication of Epstein-Barr virus. Cell 55:427–433. Han, Y., Loo, Y. M., Militello, K. T., and Melendy, T. (1999). Interactions of the papovavirus DNA replication initiator proteins, bovine papillomavirus type 1 E1 and simian virus 40 large T antigen, with human replication protein A. J. Virol. 73:4899–4907. Hartley, K. A., and Alexander, K. A. (2002). Human TATA binding protein inhibits human papillomavirus type 11 DNA replication by antagonizing E1-E2 protein complex formation on the viral origin of replication. J. Virol. 76:5014–5023. Hegde, R. S., Grossman, S. R., Laimins, L. A., and Sigler, P. B. (1992). Crystal structure at 1.7 A of the bovine papillomavirus-1 E2 DNA- binding domain bound to its DNA target. Nature 359:505–512.
196
Alison A. McBride
Hernandez-Ramon, E. E., Burns, J. E., Zhang, W., Walker, H. F., Allen, S., Antson, A. A., and Maitland, N. J. (2008). Dimerization of the human papillomavirus type 16 E2 N terminus results in DNA looping within the upstream regulatory region. J. Virol. 82:4853–4861. Hickman, A. B., Ronning, D. R., Perez, Z. N., Kotin, R. M., and Dyda, F. (2004). The nuclease domain of adeno-associated virus rep coordinates replication initiation using two distinct DNA recognition interfaces. Mol. Cell 13:403–414. Hindmarsh, P. L., and Laimins, L. A. (2007). Mechanisms regulating expression of the HPV 31 L1 and L2 capsid proteins and pseudovirion entry. Virol. J. 4:19. Hoffmann, R., Hirt, B., Bechtold, V., Beard, P., and Raj, K. (2006). Different modes of human papillomavirus DNA replication during maintenance. J. Virol. 80:4431–4439. Holt, S. E., Schuller, G., and Wilson, V. G. (1994). DNA binding specificity of the bovine papillomavirus E1 protein is determined by sequences contained within an 18-base-pair inverted repeat element at the origin of replication. J. Virol. 68:1094–1102. Hsu, C. Y., Mechali, F., and Bonne-Andrea, C. (2007). Nucleocytoplasmic shuttling of bovine papillomavirus E1 helicase downregulates viral DNA replication in S phase. J. Virol. 81:384–394. Hubert, W. G., and Lambert, P. F. (1993). The 23-kilodalton E1 phosphoprotein of bovine papillomavirus type 1 is nonessential for stable plasmid replication in murine C127 cells. J. Virol. 67:2932–2937. Huh, K. W., DeMasi, J., Ogawa, H., Nakatani, Y., Howley, P. M., and Munger, K. (2005). Association of the human papillomavirus type 16 E7 oncoprotein with the 600-kDa retinoblastoma protein-associated factor, p600. Proc. Natl. Acad. Sci. USA 102:11492–11497. Ilves, I., Kadaja, M., and Ustav, M. (2003). Two separate replication modes of the bovine papillomavirus BPV1 origin of replication that have different sensitivity to p53. Virus Res. 96:75–84. Ilves, I., Kivi, S., and Ustav, M. (1999). Long-term episomal maintenance of bovine papillomavirus type 1 plasmids is determined by attachment to host chromosomes, which Is mediated by the viral E2 protein and its binding sites. J. Virol. 73:4404–4412. Ilves, I., Maemets, K., Silla, T., Janikson, K., and Ustav, M. (2006). Brd4 is involved in multiple processes of the bovine papillomavirus type 1 life cycle. J. Virol. 80:3660–3665. Ilyina, T. V., and Koonin, E. V. (1992). Conserved sequence motifs in the initiator proteins for rolling circle DNA replication encoded by diverse replicons from eubacteria, eucaryotes and archaebacteria. Nucleic Acids Res. 20:3279–3285. Jacob, F., and Brenner, S. (1963). [On the regulation of DNA synthesis in bacteria: The hypothesis of the replicon.]. C.R.Hebd. Seances Acad. Sci. 256:298–300. Jang, M. K., Mochizuki, K., Zhou, M., Jeong, H. S., Brady, J. N., and Ozato, K. (2005). The bromodomain protein Brd4 is a positive regulatory component of P-TEFb and stimulates RNA polymerase II-dependent transcription. Mol. Cell 19:523–534. Janin, J., Rodier, F., Chakrabarti, P., and Bahadur, R. P. (2007). Macromolecular recognition in the Protein Data Bank. Acta Crystallogr. D.Biol. Crystallogr. 63:1–8. Jeckel, S., Huber, E., Stubenrauch, F., and Iftner, T. (2002). A transactivator function of cottontail rabbit papillomavirus e2 is essential for tumor induction in rabbits. J. Virol. 76:11209–11215. Jeckel, S., Loetzsch, E., Huber, E., Stubenrauch, F., and Iftner, T. (2003). Identification of the E9/E2C cDNA and functional characterization of the gene product reveal a new repressor of transcription and replication in cottontail rabbit papillomavirus. J. Virol. 77:8736–8744. Jenke, B. H., Fetzer, C. P., Stehle, I. M., Jonsson, F., Fackelmayer, F. O., Conradt, H., Bode, J., and Lipps, H. J. (2002). An episomally replicating vector binds to the nuclear matrix protein SAF-A in vivo. EMBO Rep. 3:349–354.
Replication and Partitioning of Papillomavirus Genomes
197
Jiang, Y. W., Veschambre, P., Erdjument-Bromage, H., Tempst, P., Conaway, J. W., Conaway, R. C., and Kornberg, R. D. (1998). Mammalian mediator of transcriptional regulation and its possible role as an end-point of signal transduction pathways. Proc Natl. Acad. Sci. USA 95:8538–8543. Joyce, J. G., Tung, J. S., Przysiecki, C. T., Cook, J. C., Lehman, E. D., Sands, J. A., Jansen, K. U., and Keller, P. M. (1999). The L1 major capsid protein of human papillomavirus type 11 recombinant virus-like particles interacts with heparin and cell-surface glycosaminoglycans on human keratinocytes. J. Biol. Chem. 274:5810–5822. Kapoor, P., and Frappier, L. (2003). EBNA1 partitions Epstein-Barr virus plasmids in yeast cells by attaching to human EBNA1-binding protein 2 on mitotic chromosomes. J. Virol. 77:6946–6956. Kelley-Clarke, B., Ballestas, M. E., Komatsu, T., and Kaye, K. M. (2007). Kaposi’s sarcoma herpesvirus C-terminal LANA concentrates at pericentromeric and peri-telomeric regions of a subset of mitotic chromosomes. Virology 357:149–157. Kim, K., Angeletti, P. C., Hassebroek, E. C., and Lambert, P. F. (2005). Identification of cisacting elements that mediate the replication and maintenance of human papillomavirus type 16 genomes in Saccharomyces cerevisiae. J. Virol. 79:5933–5942. Kim, K., Garner-Hamrick, P. A., Fisher, C., Lee, D., and Lambert, P. F. (2003). Methylation patterns of papillomavirus DNA, its influence on E2 function, and implications in viral infection. J. Virol. 77:12450–12459. Kim, K., and Lambert, P. F. (2002). E1 protein of bovine papillomavirus 1 is not required for the maintenance of viral plasmid DNA replication. Virology 293:10–14. Kirchmaier, A. L., and Sugden, B. (1998). Rep*: A viral element that can partially replace the origin of plasmid DNA synthesis of Epstein-Barr virus. J. Virol. 72:4657–4666. Kirnbauer, R., Booy, F., Cheng, N., Lowy, D. R., and Schiller, J. T. (1992). Papillomavirus L1 major capsid protein self-assembles into virus-like particles that are highly immunogenic. Proc. Natl. Acad. Sci. USA 89:12180–12184. Knight, J. D., Li, R., and Botchan, M. (1991). The activation domain of the bovine papillomavirus E2 protein mediates association of DNA-bound dimers to form DNA loops. Proc. Natl. Acad. Sci. USA 88:3204–3208. Kobayashi, T., Heck, D. J., Nomura, M., and Horiuchi, T. (1998). Expansion and contraction of ribosomal DNA repeats in Saccharomyces cerevisiae: Requirement of replication fork blocking (Fob1) protein and the role of RNA polymerase I. Genes Dev. 12:3821–3830. Krithivas, A., Fujimuro, M., Weidner, M., Young, D. B., and Hayward, S. D. (2002). Protein interactions targeting the latency-associated nuclear antigen of Kaposi’s sarcomaassociated herpesvirus to cell chromosomes. J. Virol. 76:11596–11604. Kruhlak, M. J., Hendzel, M. J., Fischle, W., Bertos, N. R., Hameed, S., Yang, X. J., Verdin, E., and Bazett-Jones, D. P. (2001). Regulation of global acetylation in mitosis through loss of histone acetyltransferases and deacetylases from chromatin. J. Biol. Chem. 276:38307–38319. Kuo, S. R., Liu, J. S., Broker, T. R., and Chow, L. T. (1994). Cell-free replication of the human papillomavirus DNA with homologous viral E1 and E2 proteins and human cell extracts. J. Biol. Chem. 269:24058–24065. Kurg, R., Tekkel, H., Abroi, A., and Ustav, M. (2006). Characterization of the functional activities of the bovine papillomavirus type 1 E2 protein single-chain heterodimers. J. Virol. 80:11218–11225. Lacey, C. J. (2005). Therapy for genital human papillomavirus-related disease. J. Clin. Virol. 32:S82–S90. Lai, M. C., Teh, B. H., and Tarn, W. Y. (1999). A human papillomavirus E2 transcriptional activator. The interactions with cellular splicing factors and potential function in premRNA processing. J. Biol. Chem. 274:11832–11841.
198
Alison A. McBride
Lambert, P. F., Monk, B. C., and Howley, P. M. (1990). Phenotypic analysis of bovine papillomavirus type 1 E2 repressor mutants. J. Virol. 64:950–956. Lambert, P. F., Spalholz, B. A., and Howley, P. M. (1987). A transcriptional repressor encoded by BPV-1 shares a common carboxy-terminal domain with the E2 transactivator. Cell 50:69–78. Lee, D., Kim, H., Lee, Y., and Choe, J. (1997). Identification of sequence requirement for the origin of DNA replication in human papillomavirus type 18. Virus Res. 52:97–108. Lee, K. Y., Broker, T. R., and Chow, L. T. (1998). Transcription factor YY1 represses cell-free replication from human papillomavirus origins. J. Virol. 72:4911–4917. Lehman, C. W., and Botchan, M. R. (1998). Segregation of viral plasmids depends on tethering to chromosomes and is regulated by phosphorylation. Proc. Natl. Acad. Sci. USA 95:4338–4343. Lentz, M. R. (2002). A carboxyl-terminal serine of the bovine papillomavirus E1 protein is phosphorylated in vivo and in vitro. Virus Res. 83:213–219. Lentz, M. R., Pak, D., Mohr, I., and Botchan, M. R. (1993). The E1 replication protein of bovine papillomavirus type 1 contains an extended nuclear localization signal that includes a p34cdc2 phosphorylation site. J. Virol. 67:1414–1423. Lentz, M. R., Stevens, S. M., Jr., Raynes, J., and Elkhoury, N. (2006). A phosphorylation map of the bovine papillomavirus E1 helicase. Virol. J. 3:13. Li, D., Zhao, R., Lilyestrom, W., Gai, D., Zhang, R., DeCaprio, J. A., Fanning, E., Jochimiak, A., Szakonyi, G., and Chen, X. S. (2003). Structure of the replicative helicase of the oncoprotein SV40 large tumour antigen. Nature 423:512–518. Li, R., and Botchan, M. R. (1993). The acidic transcriptional activation domains of VP16 and p53 bind the cellular replication protein A and stimulate in vitro BPV-1 DNA replication. Cell 73:1207–1221. Li, R., and Botchan, M. R. (1994). Acidic transcription factors alleviate nucleosome-mediated repression of DNA replication of bovine papillomavirus type 1. Proc. Natl. Acad. Sci. USA 91:7051–7055. Li, R., Knight, J., Bream, G., Stenlund, A., and Botchan, M. (1989). Specific recognition nucleotides and their DNA context determine the affinity of E2 protein for 17 binding sites in the BPV- 1 genome. Genes Dev. 3:510–526. Lim, D. A., Gossen, M., Lehman, C. W., and Botchan, M. R. (1998). Competition for DNA binding sites between the short and long forms of E2 dimers underlies repression in bovine papillomavirus type 1 DNA replication control. J. Virol. 72:1931–1940. Lin, B. Y., Ma, T., Liu, J. S., Kuo, S. R., Jin, G., Broker, T. R., Harper, J. W., and Chow, L. T. (2000). HeLa cells are phenotypically limiting in cyclin E/CDK2 for efficient human papillomavirus DNA replication. J. Biol. Chem. 275:6167–6174. Liu, X., Schuck, S., and Stenlund, A. (2007). Adjacent residues in the E1 initiator beta-hairpin define different roles of the beta-hairpin in Ori melting, helicase loading, and helicase activity. Mol. Cell 25:825–837. Loo, Y. M., and Melendy, T. (2004). Recruitment of replication protein A by the papillomavirus E1 protein and modulation by single-stranded DNA. J. Virol. 78:1605–1615. Lu, J. Z., Sun, Y. N., Rose, R. C., Bonnez, W., and McCance, D. J. (1993). Two E2 binding sites (E2BS) alone or one E2BS plus an A/T-rich region are minimal requirements for the replication of the human papillomavirus type 11 origin. J. Virol. 67:7131–7139. Luo, X., Sanford, D. G., Bullock, P. A., and Bachovchin, W. W. (1996). Solution structure of the origin DNA-binding domain of SV40 T-antigen. Nat. Struct. Biol. 3:1034–1039. Lusky, M., and Botchan, M. R. (1986). A bovine papillomavirus type 1-encoded modulator function is dispensable for transient viral replication but is required for establishment of the stable plasmid state. J. Virol. 60:729–742. Ma, T., Zou, N., Lin, B. Y., Chow, L. T., and Harper, J. W. (1999). Interaction between cyclindependent kinases and human papillomavirus replication-initiation protein E1 is required for efficient viral replication. Proc. Natl. Acad. Sci. USA 96:382–387.
Replication and Partitioning of Papillomavirus Genomes
199
Mannik, A., Piirsoo, M., Nordstrom, K., Ustav, E., Vennstrom, B., and Ustav, M. (2003). Effective generation of transgenic mice by Bovine papillomavirus type 1 based selfreplicating plasmid that is maintained as extrachromosomal genetic element in three generations of animals. Plasmid 49:193–204. Mannik, A., Runkorg, K., Jaanson, N., Ustav, M., and Ustav, E. (2002). Induction of the bovine papillomavirus origin ‘‘onion skin’’-type DNA replication at high E1 protein concentrations in vivo J. Virol. 76:5835–5845. Mansky, K. C., Batiza, A., and Lambert, P. F. (1997). Bovine papillomavirus type 1 E1 and simian virus 40 large T antigen share regions of sequence similarity required for multiple functions. J. Virol. 71:7600–7608. Maruyama, T., Farina, A., Dey, A., Cheong, J., Bermudez, V. P., Tamura, T., Sciortino, S., Shuman, J., Hurwitz, J., and Ozato, K. (2002). A Mammalian bromodomain protein, Brd4, interacts with replication factor C and inhibits progression to S phase. Mol. Cell Biol. 22:6509–6520. Masai, H., You, Z., and Arai, K. (2005). Control of DNA replication: Regulation and activation of eukaryotic replicative helicase, MCM. IUBMB.Life 57:323–335. Masterson, P. J., Stanley, M. A., Lewis, A. P., and Romanos, M. A. (1998). A C-terminal helicase domain of the human papillomavirus E1 protein binds E2 and the DNA polymerase alphaprimase p68 subunit. J. Virol. 72:7407–7419. McBride, A. A., Dlugosz, A., and Baker, C. C. (2000). Production of infectious BPV-1 virions from cloned viral DNA using an organotypic raft/xenograft technique. Proc. Natl. Acad. Sci. USA 97:5534–5539. McBride, A. A., Bolen, J. B., and Howley, P. M. (1989). Phosphorylation sites of the E2 transcriptional regulatory proteins of bovine papillomavirus type 1. J. Virol. 63:5076–5085. McBride, A. A., and Howley, P. M. (1991). Bovine papillomavirus with a mutation in the E2 serine 301 phosphorylation site replicates at a high copy number. J. Virol. 65:6528–6534. McBride, A. A., and Myers, G. (1997). The E2 proteins: An update. In ‘‘Human Papillomaviruses 1997’’ (G. Myers, et al.), pp. III54–III99. Los Alamos National Laboratory, Los Alamos. McBride, A. A., Oliveira, J. G., and McPhillips, M. G. (2006). Partitioning viral genomes in mitosis: Same idea, different targets. Cell Cycle 5:1499–1502. McMillan, N. A., Payne, E., Frazer, I. H., and Evander, M. (1999). Expression of the alpha6 integrin confers papillomavirus binding upon receptor-negative B-cells. Virology 261:271–279. McPhillips, M. G., Oliveira, J. G., Spindler, J. E., Mitra, R., and McBride, A. A. (2006). Brd4 is required for e2-mediated transcriptional activation but not genome partitioning of all papillomaviruses. J. Virol. 80:9530–9543. McPhillips, M. G., Ozato, K., and McBride, A. A. (2005). Interaction of bovine papillomavirus E2 protein with Brd4 stabilizes its association with chromatin. J. Virol. 79:8920–8932. McShan, G. D., and Wilson, V. G. (1997a). Casein kinase II phosphorylates bovine papillomavirus type 1 E1 in vitro at a conserved motif. J. Gen. Virol. 78:171–177. McShan, G. D., and Wilson, V. G. (1997b). Reconstitution of a functional bovine papillomavirus type 1 origin of replication reveals a modular tripartite replicon with an essential AT-rich element. Virology 237:198–208. McStay, B. (2006). Nucleolar dominance: A model for rRNA gene silencing. Genes Dev. 20:1207–1214. Mechali, F., Hsu, C. Y., Castro, A., Lorca, T., and Bonne-Andrea, C. (2004). Bovine papillomavirus replicative helicase E1 is a target of the ubiquitin ligase APC. J. Virol. 78:2615–2619.
200
Alison A. McBride
Meinke, G., Phelan, P., Moine, S., Bochkareva, E., Bochkarev, A., Bullock, P. A., and Bohm, A. (2007). The crystal structure of the SV40 T-antigen origin binding domain in complex with DNA. PLoS.Biol. 5:e23. Melendy, T., Sedman, J., and Stenlund, A. (1995). Cellular factors required for papillomavirus DNA replication. J. Virol. 69:7857–7867. Meyers, C., Frattini, M. G., Hudson, J. B., and Laimins, L. A. (1992). Biosynthesis of human papillomavirus from a continuous cell line upon epithelial differentiation. Science 257:971–973. Meyers, C., Mayer, T. J., and Ozbun, M. A. (1997). Synthesis of infectious human papillomavirus type 18 in differentiating epithelium transfected with viral DNA. J. Virol. 71:7381–7386. Mohr, I. J., Clark, R., Sun, S., Androphy, E. J., MacPherson, P., and Botchan, M. R. (1990). Targeting the E1 replication protein to the papillomavirus origin of replication by complex formation with the E2 transactivator. Science 250:1694–1699. Moody, C. A., Fradet-Turcotte, A., Archambault, J., and Laimins, L. A. (2007). Human papillomaviruses activate caspases upon epithelial differentiation to induce viral genome amplification. Proc. Natl. Acad. Sci. USA 104:19541–19546. Muller, F., Seo, Y. S., and Hurwitz, J. (1994). Replication of bovine papillomavirus type 1 origin-containing DNA in crude extracts and with purified proteins. J. Biol. Chem. 269:17086–17094. Narahari, J., Fisk, J. C., Melendy, T., and Roman, A. (2006). Interactions of the cellular CCAAT displacement protein and human papillomavirus E2 protein with the viral origin of replication can regulate DNA replication. Virology 350:302–311. Narajczyk, M., Baranska, S., Wegrzyn, A., and Wegrzyn, G. (2007). Switch from theta to sigma replication of bacteriophage lambda DNA: Factors involved in the process and a model for its regulation. Mol. Genet. Genomics 278:65–74. Negorev, D., and Maul, G. G. (2001). Cellular proteins localized at and interacting within ND10/PML nuclear bodies/PODs suggest functions of a nuclear depot. Oncogene 20:7234–7242. Newhouse, C. D., and Silverstein, S. J. (2001). Orientation of a novel DNA binding site affects human papillomavirus- mediated transcription and replication. J. Virol. 75:1722–1735. O’Connor, M. J., Stunkel, W., Koh, C. H., Zimmermann, H., and Bernard, H. U. (2000). The differentiation-specific factor CDP/Cut represses transcription and replication of human papillomaviruses through a conserved silencing element. J. Virol. 74:401–410. O’Connor, M. J., Tan, S. H., Tan, C. H., and Bernard, H. U. (1996). YY1 represses human papillomavirus type 16 transcription by quenching AP-1 activity. J. Virol. 70:6529–6539. Ohe, Y., Zhao, D., Saijo, N., and Podack, E. R. (1995). Construction of a novel bovine papillomavirus vector without detectable transforming activity suitable for gene transfer. Hum. Gene Ther. 6:325–333. Oliveira, J. G., Colf, L. A., and McBride, A. A. (2006). Variations in the association of papillomavirus E2 proteins with mitotic chromosomes. Proc. Natl. Acad. Sci. USA 103:1047–1052. Ozbun, M. A., and Meyers, C. (1998). Human papillomavirus type 31b E1 and E2 transcript expression correlates with vegetative viral genome amplification. Virology 248:218–230. Parish, J. L., Bean, A. M., Park, R. B., and Androphy, E. J. (2006). ChlR1 is required for loading papillomavirus E2 onto mitotic chromosomes and viral genome maintenance. Mol. Cell 24:867–876. Park, R. B., and Androphy, E. J. (2002). Genetic analysis of high-risk e6 in episomal maintenance of human papillomavirus genomes in primary human keratinocytes. J. Virol. 76:11359–11364.
Replication and Partitioning of Papillomavirus Genomes
201
Parkin, D. M., Bray, F., Ferlay, J., and Pisani, P. (2005). Global cancer statistics, 2002. CA Cancer J. Clin. 55:74–108. Pattison, S., Skalnik, D. G., and Roman, A. (1997). CCAAT displacement protein, a regulator of differentiation-specific gene expression, binds a negative regulatory element within the 50 end of the human papillomavirus type 6 long control region. J. Virol. 71:2013–2022. Peh, W. L., Middleton, K., Christensen, N., Nicholls, P., Egawa, K., Sotlar, K., Brandsma, J., Percival, A., Lewis, J., Liu, W. J., and Doorbar, J. (2002). Life cycle heterogeneity in animal models of human papillomavirus- associated disease. J. Virol. 76:10401–10416. Penrose, K. J., Garcia-Alai, M., Prat-Gay, G., and McBride, A. A. (2004). CK2 phosphorylationinduced conformational switch triggers degradation of the papillomavirus E2 protein. J. Biol. Chem. 279:22430–22439. Penrose, K. J., and McBride, A. A. (2000). Proteasome-mediated degradation of the papillomavirus E2-TA protein is regulated by phosphorylation and can modulate viral genome copy number. J. Virol. 74:6031–6038. Piirsoo, M., Ustav, E., Mandel, T., Stenlund, A., and Ustav, M. (1996). Cis and trans requirements for stable episomal maintenance of the BPV-1 replicator. EMBO J. 15:1–11. Pittayakhajortwut, D., and Angeletti, P. C. (2008). Analysis of cis-elements that facilitate extrachromosomal persistence of human papillomavirus genomes. Virology 374:304–314. Platt, G. M., Simpson, G. R., Mittnacht, S., and Schulz, T. F. (1999). Latent nuclear antigen of Kaposi’s sarcoma-associated herpesvirus interacts with RING3, a homolog of the Drosophila female sterile homeotic (fsh) gene. J. Virol. 73:9789–9795. Poddar, A., Reed, S. C., McPhillips, M. G., and McBride, A. A. (2008). The Human Papillomavirus Type 8 E2 Tethering Protein Targets the Ribosomal DNA Loci of Host Mitotic Chromosomes. manuscript submitted. Rangasamy, D., and Wilson, V. G. (2000). Bovine papillomavirus E1 protein is sumoylated by the host cell Ubc9 protein. J. Biol. Chem. 275:30487–30495. Rangasamy, D., Woytek, K., Khan, S. A., and Wilson, V. G. (2000). SUMO-1 modification of bovine papillomavirus E1 protein is required for intranuclear accumulation. J. Biol. Chem. 275:37999–38004. Ravnan, J. -B., Gilbert, G. M., Ten Hagen, K. G., and Cohen, S. N. (1992). Random-choice replication of extrachromosomal bovine papillomavirus (BPV) molecules in heterogeneous clonally-derived BPV-infected cell lines. J. Virol. 66:6946–6952. Rawlins, D. R., Milman, G., Hayward, S. D., and Hayward, G. S. (1985). Sequence-specific DNA binding of the Epstein-Barr virus nuclear antigen (EBNA-1) to clustered sites in the plasmid maintenance region. Cell 42:859–868. Remm, M., Brain, R., and Jenkins, J. R. (1992). The E2 binding sites determine the efficiency of replication for the origin of human papillomavirus type 18. Nucleic.Acids.Res. 20:6015–6021. Riese, D. J., II, Settleman, J., Neary, K., and DiMaio, D. (1990). Bovine papillomavirus E2 repressor mutant displays a high- copy-number phenotype and enhanced transforming activity. J. Virol. 64:944–949. Rogers, S. (1966). Shope Papilloma Virus - A passenger in man and its significance to potential control of host genome. Nature 212:1220–1222. Rosas-Acosta, G., and Wilson, V. G. (2008). Identification of a nuclear export signal sequence for bovine papillomavirus E1 protein. Virology 373:149–162. Rotenberg, M. O., Chow, L. T., and Broker, T. R. (1989). Characterization of rare human papillomavirus type 11 mRNAs coding for regulatory and structural proteins, using the polymerase chain reaction. Virology 172:489–497. Russell, J., and Botchan, M. R. (1995). cis-Acting components of human papillomavirus (HPV) DNA replication: Linker substitution analysis of the HPV type 11 origin. J. Virol. 69:651–660.
202
Alison A. McBride
Sakai, H., Yasugi, T., Benson, J. D., Dowhanick, J. J., and Howley, P. M. (1996). Targeted mutagenesis of the human papillomavirus type 16 E2 transactivation domain reveals separable transcriptional activation and DNA replication functions. J. Virol. 70:1602–1611. Sanders, C. M., Sizov, D., Seavers, P. R., Ortiz-Lombardia, M., and Antson, A. A. (2007). Transcription activator structure reveals redox control of a replication initiation reaction. Nucleic Acids Res. 35:3504–3515. Sanders, C. M., and Stenlund, A. (1998). Recruitment and loading of the E1 initiator protein: An ATP-dependent process catalysed by a transcription factor. EMBO J. 17:7044–7055. Sarafi, T. R., and McBride, A. A. (1995). Domains of the BPV-1 E1 replication protein required for origin-specific DNA binding and interaction with the E2 transactivator. Virology 211:385–396. Sarver, N., Gruss, P., Law, M. F., Khoury, G., and Howley, P. M. (1981). Bovine papilloma virus deoxyribonucleic acid: A novel eucaryotic cloning vector. Mol. Cell.Biol. 1:486–496. Schaal, T. D., Mallet, W. G., McMinn, D. L., Nguyen, N. V., Sopko, M. M., John, S., and Parekh, B. S. (2003). Inhibition of human papilloma virus E2 DNA binding protein by covalently linked polyamides. Nucleic Acids Res. 31:1282–1291. Schepers, A., Ritzi, M., Bousset, K., Kremmer, E., Yates, J. L., Harwood, J., Diffley, J. F., and Hammerschmidt, W. (2001). Human origin recognition complex binds to the region of the latent origin of DNA replication of Epstein-Barr virus. EMBO J. 20:4588–4602. Schuck, S., and Stenlund, A. (2005). Assembly of a double hexameric helicase. Mol. Cell 20:377–389. Schuck, S., and Stenlund, A. (2006). Surface mutagenesis of the bovine papillomavirus E1 DNA binding domain reveals residues required for multiple functions related to DNA replication. J. Virol. 80:7491–7499. Schweiger, M. R., You, J., and Howley, P. M. (2006). Bromodomain protein 4 mediates the papillomavirus e2 transcriptional activation function. J. Virol. 80:4276–4285. Sears, J., Ujihara, M., Wong, S., Ott, C., Middeldorp, J., and Aiyar, A. (2004). The amino terminus of Epstein-Barr Virus (EBV) nuclear antigen 1 contains AT hooks that facilitate the replication and partitioning of latent EBV genomes by tethering them to cellular chromosomes. J. Virol. 78:11487–11505. Sedman, J., and Stenlund, A. (1998). The papillomavirus E1 protein forms a DNA-dependent hexameric complex with ATPase and DNA helicase activities. J. Virol. 72:6893–6897. Sedman, T., Sedman, J., and Stenlund, A. (1997). Binding of the E1 and E2 proteins to the origin of replication of bovine papillomavirus. J. Virol. 71:2887–2896. Senechal, H., Poirier, G. G., Coulombe, B., Laimins, L. A., and Archambault, J. (2007). Amino acid substitutions that specifically impair the transcriptional activity of papillomavirus E2 affect binding to the long isoform of Brd4. Virology 358:10–17. Shafti-Keramat, S., Handisurya, A., Kriehuber, E., Meneguzzi, G., Slupetzky, K., and Kirnbauer, R. (2003). Different heparan sulfate proteoglycans serve as cellular receptors for human papillomaviruses. J. Virol. 77:13125–13135. Shire, K., Ceccarelli, D. F., Avolio-Hunter, T. M., and Frappier, L. (1999). EBP2, a human protein that interacts with sequences of the Epstein-Barr virus nuclear antigen 1 important for plasmid maintenance. J. Virol. 73:2587–2595. Si, H., Verma, S. C., Lampson, M., Cai, Q., and Robertson, E. S. (2008). KSHV Encoded LANA can Interact with the Nuclear Mitotic Apparatus Protein to Regulate Genome Maintenance and Segregation. J. Virol. 82:6734–6746. Sim, J., Ozgur, S., Lin, B. Y., Yu, J. H., Broker, T. R., Chow, L. T., and Griffith, J. (2008). Remodeling of the human papillomavirus type 11 replication origin into discrete nucleoprotein particles and looped structures by the E2 protein. J. Mol. Biol. 375:1165–1177.
Replication and Partitioning of Papillomavirus Genomes
203
Skiadopoulos, M. H., and McBride, A. A. (1998). Bovine papillomavirus type 1 genomes and the E2 transactivator protein are closely associated with mitotic chromatin. J. Virol. 72:2079–2088. Smith, J. L., Campos, S. K., and Ozbun, M. A. (2007). Human papillomavirus type 31 uses a caveolin 1- and dynamin 2-mediated entry pathway for infection of human keratinocytes. J. Virol. 81:9922–9931. Soeda, E., Ferran, M. C., Baker, C. C., and McBride, A. A. (2006). Repression of HPV16 early region transcription by the E2 protein. Virology 351:29–41. Stanley, M. A., Browne, H. M., Appleby, M., and Minson, A. C. (1989). Properties of a nontumorigenic human cervical keratinocyte cell line. Int. J. Cancer 43:672–676. Steger, G., Schnabel, C., and Schmidt, H. M. (2002). The hinge region of the human papillomavirus type 8 E2 protein activates the human p21(WAF1/CIP1) promoter via interaction with Sp1. J. Gen. Virol. 83:503–510. Stenlund, A. (2003). E1 initiator DNA binding specificity is unmasked by selective inhibition of non-specific DNA binding. EMBO J. 22:954–963. Stubenrauch, F., Colbert, A. M., and Laimins, L. A. (1998a). Transactivation by the E2 protein of oncogenic human papillomavirus type 31 is not essential for early and late viral functions. J. Virol. 72:8115–8123. Stubenrauch, F., Hummel, M., Iftner, T., and Laimins, L. A. (2000). The E8E2C protein, a negative regulator of viral transcription and replication, is required for extrachromosomal maintenance of human papillomavirus type 31 in keratinocytes. J. Virol. 74:1178–1186. Stubenrauch, F., Lim, H. B., and Laimins, L. A. (1998b). Differential requirements for conserved E2 binding sites in the life cycle of oncogenic human papillomavirus type 31. J. Virol. 72:1071–1077. Sullivan, M., Higuchi, T., Katis, V. L., and Uhlmann, F. (2004). Cdc14 phosphatase induces rDNA condensation and resolves cohesin-independent cohesion during budding yeast anaphase. Cell 117:471–482. Sun, Y., Han, H., and McCance, D. J. (1998). Active domains of human papillomavirus type 11 E1 protein for origin replication. J. Gen. Virol. 79:1651–1658. Sun, Y. N., Lu, J. Z., and McCance, D. J. (1996). Mapping of HPV-11 E1 binding site and determination of other important cis elements for replication of the origin. Virology 216:219–222. Sverdrup, F., and Khan, S. A. (1995). Two E2 binding sites alone are sufficient to function as the minimal origin of replication of human papillomavirus type 18 DNA. J. Virol. 69:1319–1323. Sverdrup, F., Sheahan, L., and Khan, S. (1999). Development of human papillomavirus plasmids capable of episomal replication in human cell lines. Gene Ther. 6:1317–1321. Swindle, C. S., Zou, N., Van Tine, B. A., Shaw, G. M., Engler, J. A., and Chow, L. T. (1999). Human papillomavirus DNA replication compartments in a transient DNA replication system. J. Virol. 73:1001–1009. Tan, S. H., Bartsch, D., Schwarz, E., and Bernard, H. U. (1998). Nuclear matrix attachment regions of human papillomavirus type 16 point toward conservation of these genomic elements in all genital papillomaviruses. J. Virol. 72:3610–3622. Thain, A., Jenkins, O., Clarke, A. R., and Gaston, K. (1996). CpG methylation directly inhibits binding of the human papillomavirus type 16 E2 protein to specific DNA sequences. J. Virol. 70:7233–7235. Thierry, F., and Yaniv, M. (1987). The BPV1-E2 trans-acting protein can be either an activator or a repressor of the HPV18 regulatory region. EMBO J. 6:3391–3397. Thomas, J. T., Hubert, W. G., Ruesch, M. N., and Laimins, L. A. (1999). Human papillomavirus type 31 oncoproteins E6 and E7 are required for the maintenance of episomes during the viral life cycle in normal human keratinocytes. Proc. Natl. Acad. Sci. USA 96:8449–8454.
204
Alison A. McBride
Titolo, S., Brault, K., Majewski, J., White, P. W., and Archambault, J. (2003). Characterization of the minimal DNA binding domain of the human papillomavirus e1 helicase: Fluorescence anisotropy studies and characterization of a dimerization-defective mutant protein. J. Virol. 77:5178–5191. Titolo, S., Pelletier, A., Pulichino, A. M., Brault, K., Wardrop, E., White, P. W., Cordingley, M. G., and Archambault, J. (2000). Identification of domains of the human papillomavirus type 11 E1 helicase involved in oligomerization and binding to the viral origin. J. Virol. 74:7349–7361. Torres-Rosell, J., Machin, F., Farmer, S., Jarmuz, A., Eydmann, T., Dalgaard, J. Z., and Aragon, L. (2005). SMC5 and SMC6 genes are required for the segregation of repetitive chromosome regions. Nat. Cell Biol. 7:412–419. Tsurumi, T., Fujita, M., and Kudoh, A. (2005). Latent and lytic Epstein-Barr virus replication strategies. Rev. Med. Virol. 15:3–15. Ustav, E., Ustav, M., Szymanski, P., and Stenlund, A. (1993). The bovine papillomavirus origin of replication requires a binding site for the E2 transcriptional activator. Proc. Natl. Acad. Sci. USA 90:898–902. Ustav, M., and Stenlund, A. (1991). Transient replication of BPV-1 requires two viral polypeptides encoded by the E1 and E2 open reading frames. EMBO J. 10:449–457. Ustav, M., Ustav, E., Szymanski, P., and Stenlund, A. (1991). Identification of the origin of replication of bovine papillomavirus and characterization of the viral origin recognition factor E1. EMBO J. 10:4321–4329. Van Tine, B. A., Dao, L. D., Wu, S. Y., Sonbuchner, T. M., Lin, B. Y., Zou, N., Chiang, C. M., Broker, T. R., and Chow, L. T. (2004). Human papillomavirus (HPV) origin-binding protein associates with mitotic spindles to enable viral DNA partitioning. Proc. Natl. Acad. Sci. USA 101:4030–4035. Viejo-Borbolla, A., Ottinger, M., Bruning, E., Burger, A., Konig, R., Kati, E., Sheldon, J. A., and Schulz, T. F. (2005). Brd2/RING3 interacts with a chromatin-binding domain in the Kaposi’s Sarcoma-associated herpesvirus latency-associated nuclear antigen 1 (LANA1) that is required for multiple functions of LANA-1. J. Virol. 79:13618–13629. Voitenleitner, C., and Botchan, M. (2002). E1 protein of bovine papillomavirus type 1 interferes with E2 protein- mediated tethering of the viral DNA to mitotic chromosomes. J. Virol. 76:3440–3451. Walboomers, J. M., Jacobs, M. V., Manos, M. M., Bosch, F. X., Kummer, J. A., Shah, K. V., Snijders, P. J., Peto, J., Meijer, C. J., and Munoz, N. (1999). Human papillomavirus is a necessary cause of invasive cervical cancer worldwide. J. Pathol. 189:12–19. Waldenstrom, M., Schenstrom, K., Sollerbrant, K., and Hansson, L. (1992). Replication of bovine papillomavirus vectors in murine cells. Gene 120:175–181. Wang, Y., Coulombe, R., Cameron, D. R., Thauvette, L., Massariol, M. J., Amon, L. M., Fink, D., Titolo, S., Welchner, E., Yoakim, C., Archambault, J., and White, P. W. (2004). Crystal structure of the E2 transactivation domain of human papillomavirus type 11 bound to a protein interaction inhibitor. J. Biol.Chem. 279:6976–6985. White, P. W., Titolo, S., Brault, K., Thauvette, L., Pelletier, A., Welchner, E., Bourgon, L., Doyon, L., Ogilvie, W. W., Yoakim, C., Cordingley, M. G., and Archambault, J. (2003). Inhibition of human papillomavirus DNA replication by small molecule antagonists of the E1–E2 protein interaction. J. Biol. Chem. 278:26765–26772. Winokur, P. L., and McBride, A. A. (1996). The transactivation and DNA binding domains of the BPV-1 E2 protein have different roles in cooperative origin binding with the E1 protein. Virology 221:44–53. Wise-Draper, T. M., and Wells, S. I. (2008). Papillomavirus E6 and E7 proteins and their cellular targets. Front Biosci. 13:1003–1017. Woodworth, C. D. (2002). HPV innate immunity. Front Biosci. 7:d2058–d2071.
Replication and Partitioning of Papillomavirus Genomes
205
Wu, H., Ceccarelli, D. F., and Frappier, L. (2000). The DNA segregation mechanism of Epstein-Barr virus nuclear antigen 1. EMBO Rep. 1:140–144. Wu, S. Y., Lee, A. Y., Hou, S. Y., Kemper, J. K., Erdjument-Bromage, H., Tempst, P., and Chiang, C. M. (2006). Brd4 links chromatin targeting to HPV transcriptional silencing. Genes Dev. 20:2383–2396. Yang, L., Mohr, I., Fouts, E., Lim, D. A., Nohaile, M., and Botchan, M. (1993). The E1 protein of bovine papillomavirus 1 is an ATP-dependent DNA helicase. Proc. Natl. Acad. Sci. USA 90:5086–5090. Yates, J. L., and Guan, N. (1991). Epstein-Barr virus-derived plasmids replicate only once per cell cycle and are not amplified after entry into cells. J. Virol. 65:483–488. Yates, J. L., Warren, N., and Sugden, B. (1985). Stable replication of plasmids derived from Epstein-Barr virus in various mammalian cells. Nature 313:812–815. You, J., Croyle, J. L., Nishimura, A., Ozato, K., and Howley, P. M. (2004). Interaction of the bovine papillomavirus E2 protein with Brd4 tethers the viral DNA to host mitotic chromosomes. Cell 117:349–360. You, J., Schweiger, M. R., and Howley, P. M. (2005). Inhibition of E2 binding to Brd4 enhances viral genome loss and phenotypic reversion of bovine papillomavirustransformed cells. J. Virol. 79:14956–14961. You, J., Srinivasan, V., Denis, G. V., Harrington, W. J., Jr., Ballestas, M. E., Kaye, K. M., and Howley, P. M. (2006). Kaposi’s sarcoma-associated herpesvirus latency-associated nuclear antigen interacts with bromodomain protein Brd4 on host mitotic chromosomes. J. Virol. 80:8909–8919. Yu, J. H., Lin, B. Y., Deng, W., Broker, T. R., and Chow, L. T. (2007a). Mitogen-activated protein kinases activate the nuclear localization sequence of human papillomavirus type 11 E1 DNA helicase to promote efficient nuclear import. J. Virol. 81:5066–5078. Yu, T., Peng, Y. C., and Androphy, E. J. (2007b). Mitotic kinesin-like protein 2 binds and colocalizes with papillomavirus E2 during mitosis. J. Virol. 81:1736–1745. Zanardi, T. A., Stanley, C. M., Saville, B. M., Spacek, S. M., and Lentz, M. R. (1997). Modulation of bovine papillomavirus DNA replication by phosphorylation of the viral E1 protein. Virology 228:1–10. Zhang, B., Chen, W., and Roman, A. (2006). The E7 proteins of low- and high-risk human papillomaviruses share the ability to target the pRB family member p130 for degradation. Proc. Natl. Acad. Sci. USA 103:437–442. Zhang, B. Y., Li, P., Wang, E., Brahmi, Z., Dunn, K. W., Blum, J. S., and Roman, A. (2003). The E5 protein of human papillornavirus type 16 perturbs MHC class II antigen maturation in human foreskin keratinocytes treated with interferon-gamma. Virology 310:100–108. Zhao, K. N., Sun, X. Y., Frazer, I. H., and Zhou, J. (1998). DNA packaging by L1 and L2 capsid proteins of bovine papillomavirus type 1. Virology 243:482–491. Zheng, P. S., Brokaw, J., and McBride, A. A. (2005). Conditional mutations in the mitotic chromosome binding function of the bovine papillomavirus type 1 E2 protein. J. Virol. 79:1500–1509. Zhou, J., Sun, X. Y., Stenzel, D. J., and Frazer, I. H. (1991). Expression of vaccinia recombinant HPV 16 L1 and L2 ORF proteins in epithelial cells is sufficient for assembly of HPV virion-like particles. Virology 185:251–257. Zobel, T., Iftner, T., and Stubenrauch, F. (2003). The papillomavirus E8-E2C protein represses DNA replication from extrachromosomal origins. Mol. Cell Biol. 23:8352–8362. Zou, N., Lin, B. Y., Duan, F., Lee, K. Y., Jin, G., Guan, R., Yao, G., Lefkowitz, E. J., Broker, T. R., and Chow, L. T. (2000). The hinge of the human papillomavirus type 11 E2 protein contains major determinants for nuclear localization and nuclear matrix association. J. Virol. 74:3761–3770.
CHAPTER
5 Rhesus Cytomegalovirus: A Nonhuman Primate Model for The Study of Human Cytomegalovirus Yujuan Yue* and Peter A. Barry*,†,‡
Contents
I. Introduction II. Background III. Discovery of RhCMV IV. Epidemiology of RhCMV V. Pathogenesis of RhCMV VI. The Coding Capacity of RhCMV VII. Susceptibility of RhCMV to Anti-HCMV Drugs VIII. Host Immunity to RhCMV IX. Modulation of the Host by RhCMV X. Vaccine Studies in Rhesus Macaques XI. Future Directions Acknowledgements References
Abstract
Human cytomegalovirus (HCMV), a member of an ancient family of viruses (Herpesviridae), has acquired the capacity to maintain a lifelong persistent infection within an immunocompetent host. Since both primary and recurrent infections are generally subclinical, host antiviral immune responses are effective at limiting the
208 209 210 211 212 213 214 215 216 218 220 221 221
* Center for Comparative Medicine, University of California, Davis, California 95616 {
Department of Medical Pathology and Laboratory Medicine, University of California, Davis, California 95817 California National Primate Research Center, University of California, Davis, California 95616
{
Advances in Virus Research, Volume 72 ISSN 0065-3527, DOI: 10.1016/S0065-3527(08)00405-3
#
2008 Elsevier Inc. All rights reserved.
207
208
Yujuan Yue and Peter A. Barry
pathogenic potential of HCMV. However, the fact that HCMV can persist in the presence of those protective immune responses indicates that host immunity is unable to prevent or eliminate long-term reservoirs of virus. The ability of HCMV to persist has important clinical implications, a fact reflected by the spectrum of pathogenic outcomes observed in those without a fully functional immune system. Recurrence of viral replication or transmission of HCMV from an infected individual to those most susceptible to primary infection during immune suppression, deficiency, or immaturity can lead to multiorgan disease and, sometimes, death. The clinical need for a protective HCMV vaccine has been recognized for decades, but due to a conspiracy of factors, there is no approved vaccine despite intensive investigations to develop one. Animal models of HCMV have been used as systems of discovery and translation to understand viral mechanisms of persistence and pathogenesis, and to test concepts and modalities for the generation of immune responses that protect from primary infection and sequelae. This review summarizes studies in a nonhuman primate model of HCMV involving infection of rhesus macaques (Macaca mulatta) with rhesus cytomegalovirus (RhCMV). The RhCMV model serves as an important complement to those in other animals, particularly small animals, and the lessons learned from RhCMV should have direct clinical relevance to HCMV and the design of protective vaccines.
I. INTRODUCTION Winston Churchill’s description of the Soviet Union as ‘‘a riddle wrapped in a mystery inside an enigma’’ could easily be applied to our current understanding of human cytomegalovirus (HCMV). The enigma is how a virus, whose pathogenic potential is so readily controlled by host antiviral immune responses, can maintain a stable, lifelong coexistence with that host in the face of those very same immune responses. The mystery revolves around the mechanisms of how HCMV evades immunemediated clearance and the specificities of the immune response that correlate with protection from disease. The riddle is how we assemble the identified pieces of the puzzle, as well as those we do not know, into an effective vaccine strategy that protects those at risk from fulminant HCMV infection. There was an important coda to Churchill’s famous quote that has generally been forgotten with time, ‘‘. . . but perhaps there is a key.’’ In Churchill’s view, ‘‘That key [was] Russian national interests.’’ For HCMV, the key to the riddle posed above may relate to the ability of HCMV to establish, maintain, and reactivate from a persistent infection. A daunting challenge in meeting the Institute of Medicine’s call for the development of an effective vaccine (Stratton et al., 2000) is that HCMV is
RhCMV: A Model for the Study of HCMV
209
exceedingly well adapted to infection in its human host. HCMV gene expression and virus production can persist despite robust humoral and cellular responses to viral antigens, demonstrating that HCMV can partially blunt immune functions. Although antiviral immune responses generally protect from disease and limit re-infection, instances of HCMV re-infection have been documented in those with prior immunity (Boppana et al., 1999). The prevalence of HCMV in humans and its ability to persist in an immunocompetent host are undoubtedly related to its evolutionary history. CMV is an ancient virus with a wide distribution in mammals. Genetic evidence indicates that all CMVs originated from a common progenitor with each CMV species coevolving with its host during mammalian speciation (Karlin et al., 1994; McGeoch et al., 1995, 2006). Relating with the long history as an obligate intracellular pathogen, HCMV and its predecessors have had millions of years to hone their craft in response to the selective pressures of innate and adaptive immunity. This review will focus on issues of designing antiviral strategies for HCMV, and the role that the rhesus macaque model of HCMV persistence and pathogenesis can play in evaluating novel concepts and strategies in a nonhuman primate model that strongly reflects the human condition.
II. BACKGROUND HCMV is a b-herpesvirus that is widespread in humans (50–90% seroprevalence in adults) and establishes a life-long persistence following primary infection. Persistence is characterized by the presence of reservoirs of cells harboring latent viral genomes that periodically reactivate to produce infectious virus. Although it is usually asymptomatic in most immunocompetent people, it can cause severe or fatal disease in immunologically immature or immunocompromised individuals, such as transplant recipients, human immunodeficiency virus (HIV)-infected patients, and congenitally infected fetuses/newborns (Alford and Britt, 1993; Stagno, 1990). Its clinical significance has increased due to the rise in organ allograft transplantation and HIV-infected individuals. Furthermore, it is the leading infectious cause of birth defects. Many approaches have been investigated since the first attempt to immunize humans against HCMV in the 1970s (Elek and Stern, 1974; Neff et al., 1979). Unfortunately, no licensed vaccine is currently available for HCMV. The development of an effective HCMV vaccine is critically important to minimize the potential for the devastating effects of HCMV infection in those at high risk for primary HCMV infection and disease, and has been placed in the top priority by the Institute of Medicine for the clinical benefits that a protective vaccine would provide (Stratton et al., 2000).
210
Yujuan Yue and Peter A. Barry
One of the obstacles in the development of a HCMV vaccine is the fact that, due to its strict species-specificity, HCMV cannot be studied directly in any animal models (Staczek, 1990). Alternatively, the exploration of indigenous CMVs in animal models is vital to understand the mechanism of HCMV pathogenesis and persistence, and to prioritize both potential vaccine strategies and anti-viral therapies for humans. As a nonhuman primate model, RhCMV and its host, rhesus macaque (Macaca mulatta), are phylogenetically close to HCMV and humans (McGeoch et al., 1995, 2006; Shanley et al., 1985). Studies of RhCMV have shown that RhCMV shares many features of HCMV, in terms of genome coding content, persistence in healthy hosts, and RhCMV-related disease in immune compromised animals (Barry and Chang, 2007; Powers and Fru¨h, 2008). Accordingly, infection of rhesus macaques with RhCMV holds promise as a highly relevant surrogate for HCMV infection in humans.
III. DISCOVERY OF RhCMV Similar to the discovery of CMV infection in other species (Alford and Britt, 1993), RhCMV infection in monkeys was initially recognized by the histological presence of typical cytomegalic cells with intranuclear inclusions. In 1929, Stewart and Rhoads first reported intranuclear inclusions in the nasal mucous membrane of monkeys experimentally infected with poliomyelitis virus (PV), although there were no PV-associated lesions in nerve plexuses and ganglion cells within the nasal glands. Inclusions were not found in all of the infected monkeys (Stewart and Rhoads, 1929). Concurrently, Hindle and Stevenson also noted intranuclear inclusions in the kidney of apparently healthy monkeys (Hindle and Stevenson, 1929– 1930). Subsequently, in addition to nasal mucosal membrane, similar intranuclear inclusions were observed in the epithelial cells of trachea, lungs, and bile ducts in both normal and diseased monkeys. Importantly, all monkeys were noted for the absence of RhCMV-related disease (Covell, 1932). In 1935, Cowdry and Scott discovered numerous and highly variable renal nuclear inclusion-laden cells in the majority of rhesus macaques treated with irradiated ergosterol (a precursor of vitamin D2), compared to untreated animals. The presence of the inclusionbearing cells suggested that there was an activation of otherwise ‘‘latent’’ RhCMV genomes in the kidneys (Cowdry and Scott, 1935). The use of the term ‘‘latent’’ in this particular manuscript may represent the first recognized association of reactivation from latency by CMV in any species. In summary, these early discoveries demonstrated that RhCMV infection in monkeys is common and subclinical, which sometimes, can be reactivated from a latent state. It should be noted that cytomegalic cells are almost never observed nowadays in a nondiseased animal examined
RhCMV: A Model for the Study of HCMV
211
post mortem, most likely reflecting the improvements in animal husbandry and significant reductions in exposure to stressful conditions and pathogens, such as Mycobacterium tuberculosis. In 1968, RhCMV was isolated from the urine of healthy adult rhesus monkeys by Asher et al. and was classified as CMV for the characteristic cytomegalic cytopathic effect as shown by other CMVs (Asher et al., 1974). The isolation of RhCMV made it possible to investigate RhCMV biology and infection.
IV. EPIDEMIOLOGY OF RhCMV As with HCMV infection in humans, RhCMV infection is ubiquitous in captive and wild monkeys (Andrade et al., 2003; Jones-Engel et al., 2006; Swack and Hsiung, 1982; Swack et al., 1971; Vogel et al., 1994) and can establish a persistent, life-long infection following initial exposure (Asher et al., 1974; Huff et al., 2003). Seroprevalence rates reach 100% around 1 year of age, well before the age of sexual maturity in rhesus macaques (2.5–3 years). All nonhuman primate species analyzed to date appear to harbor their own species-specific CMV (Barry and Chang, 2007), with one exception. Barbary macaques (M. sylvanus) in Gibraltar are seronegative for any virus with antigenic relatedness to RhCMV and other normally universal viruses (Engel et al., 2008). Adult rhesus macaques can secrete virus years after primary infection (Asher et al., 1974; Huff et al., 2003) and transmit the virus to cohoused naı¨ve animals (Strelow, Yue, and Barry, unpublished). The routes of RhCMV transmission have not been fully determined. In humans, direct contact with infectious secretions, such as saliva, tears, urine, feces, semen, and milk, and intrauterine exposure is required for transmission. Similar routes appear to exist in rhesus macaques. RhCMV shedding can be persistently or intermittently detected in urine, saliva, and genital secretions of healthy rhesus monkeys (Asher et al., 1974; Huff et al., 2003). There are no reports of RhCMV in breast milk, although this route of transmission has probably never been examined. Neither vertical transmission from mother to fetus nor histological evidence of RhCMV disease in fetuses or neonates has been documented, although low rates of transmission (<5%) cannot be excluded. The high prevalence of RhCMV infection in the population of rhesus macaques may explain the absence of congenital infection. Maternal preconceptional immunity to HCMV provides substantial protection against both transplacental transmission and clinical outcomes in those fetuses that are congenitally infected. The highest risk for congenital HCMV infection is among infants born to mothers who have had primary infection during pregnancy (Adler et al., 1995; Fowler et al., 1992; Stagno et al., 1986). Since 100% of rhesus macaques are exposed to the virus and become
212
Yujuan Yue and Peter A. Barry
seropositive to RhCMV before reaching breeding age (Andrade et al., 2003; Vogel et al., 1994), macaque fetuses may be protected from both transplacental transmission and RhCMV-induced congenital abnormalities by preexisting maternal immunity.
V. PATHOGENESIS OF RhCMV HCMV infection of immunocompetent hosts is usually asymptomatic but may cause a mild febrile illness and, uncommonly, mononucleosis, pneumonia, and hepatitis. Similarly, RhCMV infection in healthy immunocompetent rhesus macaques following either natural exposure or experimental inoculation does not result in overt clinical signs of disease. However, transient hematological changes, such as lymphocytosis, monocytosis, and neutrophilia are observed in experimentally infected rhesus macaques (Lockridge et al., 1999). Histopathological, immunohistochemical examination, and viral DNA detection of necropsy and postmortem tissues show that RhCMV infects a wide range of tissues and multiple cell types within an involved organ, consistent with those identified in HCMV infection (Chang et al., 2002; Covell, 1932; Cowdry and Scott, 1935; Lockridge et al., 1999; Stewart and Rhoads, 1929). For example, RhCMVpositive cells have been identified in the ductal cells of the parotid and submandibular salivary glands, endothelial cells, and macrophages (Lockridge et al., 1999). Moreover, antigen-positive epithelial cells have been identified following the explant into culture of kidney cells isolated from macaques experimentally inoculated with a genetic variant of RhCMV that has undergone minimal passage in cultures (Schmidt, Yue, Chang, and Barry, unpublished). HCMV infection in fetuses is the most infectious cause of multiorgan developmental abnormalities, particularly within the central nervous system (CNS) and sensory and neural components of the auditory complex (Alford and Britt, 1993; Dollard et al., 2007). The neuropathological changes include lissencephaly-pachygyria, polymicrogyria, ventriculomegaly, periventricular calcifications, and microcephaly. While no cases of congenital RhCMV infection have yet been reported in naturally infected monkeys, direct intrauterine inoculation of RhCMV in monkey fetuses leads to neuropathologic outcomes and sensorineural abnormalities that are almost identical to those observed in HCMV congenital infection (Barry et al., 2006; Chang et al., 2002; London et al., 1986; Tarantal et al., 1998). Additionally, rhesus macaque represents a well-suited model of human acquired immunodeficient syndrome in the research of simian immunodeficiency virus (SIV) (Haigwood, 2004) and a frequently used model of solid organ transplantation in preclinical testing of new immunosuppressive strategies (Pearson et al., 2002), since its immune system closely
RhCMV: A Model for the Study of HCMV
213
resembles that of the humans (Bontrop et al., 1995). Disseminated RhCMV infection frequently occurs in rhesus macaques infected with SIV or treated with immunosuppressive drugs (Barry and Chang, 2007; Baskin, 1987). Characteristic RhCMV lesions can be found in multiple organs, such as lungs, gastrointestinal tract, adrenals, and CNS (Baskin, 1987; Conway et al., 1990; Kuhn et al., 1999; Pearson et al., 2002), comparable to the infection of HCMV in transplant recipients and AIDS patients (Alford and Britt, 1993; Landolfo et al., 2003; Pass, 2001). RhCMV retinitis has never been observed in an immunodeficient macaque. While this may be due to a difference in the pathogenic potential of RhCMV from HCMV, it may also be a factor of then humane endpoint for macaque experiments. SIV-infected macaques are culled early after the onset of simian AIDS, well before the onset of prolonged CD4þ T cell depletion that is characteristic of human AIDS patients. The absence of prolonged CD4þ T cell depletion may be associated with the absence of any observations of RhCMV retinitis.
VI. THE CODING CAPACITY OF RhCMV The RhCMV and HCMV genomes are similar in size, structure, and genetic organization. The genome of the prototypical RhCMV strain 68–1 is 221,459 bp in length and is predicted to contain 230 open reading frames (ORFs) of >100 amino acid (aa) residues (Hansen et al., 2003). The 180.92 variant of RhCMV encodes 258 ORFs of 21–2178 aa (Rivailler et al., 2006), whereas the HCMV has a 229,354 bp genome and comprises between 165 and 250 ORFs (Chee et al., 1990; Davison et al., 2003; Murphy et al., 2003a,b). The genomic arrangement of RhCMV is generally collinear with the HCMV genome, even though it lacks internal and terminal repeats, and does not isomerize (Chang and Barry, 2003). Almost 60% of potential RhCMV ORFs are homologous to HCMV proteins, including viral structural and immunomodulatory proteins, G proteincoupled receptors, and proteins involved in nucleic acid metabolism, DNA replication, and transcriptional regulation (Barry and Chang, 2007; Barry et al., 1996; Hansen et al., 2003; Lockridge et al., 2000; Oxford et al., 2008; Rivailler et al., 2006). Amino acid identities range from 24–87% (Barry and Chang, 2007; Hansen et al., 2003; North et al., 2004; Rivailler et al., 2006). A comparison of the full-length sequence of RhCMV isolates 180.92 and 68–1 highlights the complexity and instability of the RhCMV genome similar to that of HCMV (Hansen et al., 2003; Rivailler et al., 2006). Although the two strains exhibit 98% nucleic acid identity, a considerably divergent region was observed at the right end of the UL ORFs homologous to the HCMV UL proteins, a region of the RhCMV genome corresponding to a segment of the HCMV genome termed ULb0 . HCMV
214
Yujuan Yue and Peter A. Barry
ULb0 is a 13–15 kb DNA segment that undergoes genomic rearrangements, deletions, and point mutations during in vitro passage and is hypervariable among different clinical isolates (Cha et al., 1996; Pignatelli et al., 2004; Prichard et al., 2001). Further analysis of the ULb0 region of wild-type RhCMV DNA collected from naturally infected macaques has identified that the structure of ‘‘wild-type’’ RhCMV ULb0 in RhCMV isolates circulating in breeding populations of macaques is distinct from the ULb0 region of both 68–1 and 180.92 (Oxford et al., 2008). A total of 24 potential ORFs of >99 amino acids are contained within the 13-kb sequence, 10 of which are homologous to HCMV ORFs. This includes three novel a-chemokine-like ORFs, not present in either 68–1 or 180.92, in addition to previously listed homologues of HCMV UL144, 146, and 147. Several of the ORFs, including some of the a-chemokine-like proteins, exhibit exceedingly high sequence divergence amongst different genetic variants, comparable to the degree of divergence demonstrated for HCMV UL146 (Arav-Boger et al., 2006; Hassan-Walker et al., 2004; Lurain et al., 2006; Prichard et al., 2001). Preliminary work indicates that differences in the coding content of ULb0 contribute to differences in dissemination in experimentally inoculated macaques (Barry et al., in preparation). While many of the HCMV ORFs remain undefined by function and/or sequence homology, continued study of the RhCMV homologues in macaques would be of great use in determining gene products that contribute to viral mechanisms of persistence and pathogenesis.
VII. SUSCEPTIBILITY OF RhCMV TO ANTI-HCMV DRUGS Antiviral drugs licensed for HCMV treatment and prophylaxis play a pivotal role in controlling disseminated CMV infection in transplantation recipients and AIDS patients. However, they all are notably associated with toxicity, resistance, and/or pharmacokinetic limitations (Villarreal, 2003). New drugs that are more efficacious and less toxic are clearly needed, especially in the absence of a licensed HCMV vaccine. RhCMV and HCMV exhibit comparable susceptibility to currently approved compounds ganciclovir (GCV), foscarnet (phosphonoformic acid, PFA) (Swanson et al., 1998), and a promising new class of anti-HCMV compounds, benzimidazole nucleosides (North et al., 2004). GCV and PFA inhibit virus replication by targeting HCMV DNA polymerase (pUL54) and resistance can result from mutations in viral phosphotransferase gene (UL97) (only for GCV) and UL54 gene (Chou, 1999). Benzimidazole nucleosides block the cleavage of concatemeric HCMV DNA into genome-length pieces for packaging by targeting HCMV transport/capsid assembly protein (pUL56) and DNA packaging protein (pUL89)
RhCMV: A Model for the Study of HCMV
215
(Krosky et al., 1998; Townsend et al., 1995; Underwood et al., 1998), and resistance to these compounds can be induced by specific mutations in these genes. Sequence comparisons reveal that the UL54, UL56, UL89, and UL97 of HCMV are highly conserved with their RhCMV homologues, including conservation of amino acids associated with drug resistance (Hansen et al., 2003; North et al., 2004; Rivailler et al., 2006). Therefore, RhCMV infection of macaques may serve as a suitable model to test the efficacy and safety of novel anti-HCMV drugs, both in immunocompetent and immunocompromised animals.
VIII. HOST IMMUNITY TO RhCMV As with HCMV infection in humans, RhCMV-specific immunity must be viewed as protective because it can contain the pathogenic potential of RhCMV infection in immunocompetent macaques, whereas symptomatic infection is only associated with impaired or immature immunity. Although the nature of protective immune responses for HCMV remains incompletely defined, natural history studies and animal models have demonstrated that CMV specific immune responses, especially neutralizing antibodies and effective cytotoxic T lymphocyte (CTL) responses, play important roles for the control of CMV infection and the recovery of CMV-associated disease (Zhong and Khanna, 2007). Host immune responses to RhCMV infection parallel those of humans to HCMV infection as viewed by protection against RhCMV disease. Immunocompetent rhesus macaques do not display clinical signs of disease following natural or experimental RhCMV infection. Vigorous class I MHC restricted and CD8þ T cells mediated RhCMV-specific CTL activity and high frequency of RhCMV-specific CD4þ T cells are found in the peripheral blood of healthy seropositive monkeys (Kaur et al., 1996, 2002). Prominent viral targets for T cell responses include the immediate-early 1 and 2 proteins, and the RhCMV homologue of HCMV pp65, pp65–2 (Chan and Kaur, 2007; Yue et al., 2006). Detailed analysis of RhCMV proteins which represent the predominant CTL targets has not been reported. Following primary infection, RhCMV specific antibodies, including neutralizing antibodies are detected within a few weeks postinfection, increase over time, and remain relatively stable for the lifetime of the host (Lockridge et al., 1999; Swack and Hsiung, 1982). RhCMV glycoprotein B (gB), like HCMV gB, encodes the majority of neutralizing antibodies but not all (Yue et al., 2003), indicating that other viral glycoproteins represent additional determinants for neutralizing antibody responses. Neutralizing antibody titers remain relatively constant over the life of the host after reaching a plateau titer that can vary 20-fold or greater between animals. It is not known whether virus–host interactions
216
Yujuan Yue and Peter A. Barry
lead to the wide disparity between infected monkeys in neutralizing antibody or cellular responses. Preliminary evidence indicates that the differences in neutralizing antibody responses are not associated with differences in viral parameters of infection, such as shedding. In a large survey of RhCMV-infected macaques in a large outdoor breeding cohort, a large number of animals are shedding virus in saliva at any one time. However, there is no apparent association between neutralizing antibody titer and shedding of virus in saliva (Oxford et al., submitted for publication). As with HCMV infection in humans (Boppana et al., 1999; Novak et al., 2008), prior immunity to RhCMV does not appear to prevent reinfection by RhCMV. Multiple genetic variants can be isolated from an individual immunocompetent animals housed in outdoor breeding cohorts (Oxford et al., 2008), which is not surprising given the high percentage of RhCMV-infected animals shedding virus (Huff et al., 2003). On the other hand, severe infection is often observed in rhesus macaques without a fully functional immune system. In SIV-infected rhesus macaques, it has been noted that the magnitude and rate of decline of RhCMV-specific antibodies, CD4þ and CD8þ T cells are directly correlated to the increase of RhCMV DNA in plasma and the progression to RhCMV disease (Baroncelli et al., 1997; Kaur et al., 2002, 2003; Sequar et al., 2002). In experimentally inoculated rhesus macaque fetuses, the risk of CNS abnormalities is inversely associated with transplacental transfer of maternal IgG and the development of fetal immunity to RhCMV (Barry et al., 2006). Together, these data further confirm, in the macaque model, that the interplay between virus and host immunity determines the clinical outcomes of RhCMV infection.
IX. MODULATION OF THE HOST BY RhCMV HCMV encodes numerous immunomodulatory proteins that disrupt cell signaling, activation, trafficking, and death (Mocarski, 2002). RhCMV also contains homologues of most of these genes, although HCMV encodes a greater number of encoded ORFs that appear to target natural killer cells (Hansen et al., 2003; Oxford et al., 2008; Pande et al., 2005; Rivailler et al., 2006). Most of them share low homology with their HCMV counterparts, but exhibit similar functions as shown by in vitro studies. The product of the UL111a ORF is a highly divergent sequence homologue of cellular interleukin-10 (cIL-10). Both RhCMV IL-10 and HCMV IL-10 exhibit 25– 27% amino acid identity with their host’s cIL-10 (Kotenko et al., 2000; Lockridge et al., 2000). Despite this sequence divergence, work with HCMV IL-10 has shown that it binds to the cIL-10 high affinity receptor with higher affinity than does cIL-10 (Jones et al., 2002). Moreover, the
RhCMV: A Model for the Study of HCMV
217
functional activities of HCMV and cIL-10 are virtually indistinguishable (Chang et al., 2004, 2007; Spencer et al., 2002). In vitro analysis of RhCMV IL-10 indicates that it shares most, if not all, of the functions of HCMV IL10 (Chang et al., unpublished). Consistent with an anti-inflammatory role during viral infection, inoculation of seronegative macaques with a RhCMV variant lacking the RhCMV IL-10 gene results in a rapid and far greater inflammatory response at the site of inoculation, compared with inoculation with the parental, RhCMV IL-10-positive virus (Chang et al., in preparation). All seropositive animals in naturally infected cohorts develop anti-RhCMV IL-10 antibody responses, although it is not yet known if these antibodies neutralize RhCMV IL-10 function (Eberhardt et al., in preparation). The evolutionary force(s) driving sequence divergence between RhCMV and HCMV IL-10 is unknown, but presents an interesting conundrum. The RhCMV and HCMV IL-10 proteins are about as divergent from each other (31% identity) as they are from the host’s cIL-10 (Lockridge et al., 2000). However, the viral IL-10 exhibits exceedingly high intraspecies conservation of sequence (>98% identity). Very little sequence divergence is observed when comparing either RhCMV isolates or HCMV isolates. Presumably, the interspecies divergence was a result of co-speciation with the radiation of their primate hosts. However, the divergence in viral IL-10 proteins was not a compensatory evolution to divergence of the host’s IL-10 receptor, which are highly conserved (98% identity) between human and macaques. Most of the other immune modulating proteins in HCMV are conserved within RhCMV. However, RhCMV does not encode homologues of HCMV UL18 and 142, both of which are MHC class I-like in sequence. RhCMV encodes other ORFs associated with attenuation by HCMV of natural killer cell function, including UL40, 141, and a duplication of UL83, although functional studies have not yet been performed. HCMV UL138, important for establishment of latency in CD34þ myeloid progenitor cells in culture, is also missing from RhCMV. Preliminary evidence indicates that RhCMV can be found in CD34þ cells (Oxford and Barry, unpublished). RhCMV also encodes sequence and functional homologues of the viral inhibitor of caspase activation, UL36; the viral mitochondrial inhibitor of apoptosis, UL37; and the US2, 3, 6, and 11 ORFs that interfere with the assembly and transportation of MHC class I (McCormick et al., 2003; Pande et al., 2005). In sum, the primate CMVs contain a common set of viral proteins that modulate host cell signaling, activation, trafficking, and death. While the entire collection within each primate CMV evolved in response to co-speciation within its own host, the primate CMVs, as well as all CMVs, devote a large percentage of their coding capacity to functions targeting both innate and adaptive immune responses.
218
Yujuan Yue and Peter A. Barry
X. VACCINE STUDIES IN RHESUS MACAQUES Since HCMV was first recognized as a potential threat to the developing fetus, there have been repeated calls for a vaccine that could protect from the damaging effects of HCMV infection in those at risk for HCMV disease. The 35-year quest for a HCMV vaccine that could prevent congenital infection and fetal sequelae, as well as end-organ disease in immunosuppressed or immunodeficient individuals, remains unfulfilled. The primary objective measure for evaluating the efficiency of any vaccine is whether protective levels of immunity are generated and sustained in the vaccinees. An important issue for HCMV is the definition of what constitutes protective immunity. Using a stringent threshold, an immune response can be considered protective only if the vaccinees are absolutely protected from infection following repeated exposure to challenge virus. Alternatively, a vaccine could still be considered protective if the course of challenge virus infection was so dramatically altered that the potential for transmission (horizontal and vertical) and pathogenesis of challenge virus was essentially eliminated. The difference between the two involves the level of virus replication at the primary site of challenge and the extent of dissemination beyond. The former definition requires the generation and maintenance of sterilizing immunity with no spread of the virus, an immense hurdle considering that prior natural immunity does not necessarily protect from HCMV re-infection. The latter does not, but it does require that the immune system maintain a lifelong restriction on replication of a virus with a complex natural history of persistence in immunocompetent hosts. A hurdle for the design of an effective vaccine is that the vaccinee may be repeatedly exposed to high titers of virus. Studies of natural immunity to HCMV have clearly established that the induction of both neutralizing antibodies and HCMV-specific CD8þ and CD4þ T cell responses is critical for a successful vaccine. Based on this principle, current vaccine strategies have focused on the investigation of live attenuated whole virus vaccine, subunit vaccines consisting of proteins targeting protective immunity, such as the major neutralizing antibody target HCMV glycoprotein B (gB) and the predominant cellular antigens phosphoprotein 65 (pp65) and immediate-early protein IE1, and dense body vaccine (Gonczol and Plotkin, 2001; Plotkin, 1999, 2001). Unfortunately, to date, none of these strategies has yet achieved a protective effect against primary CMV infection and sequelae. Due to the uncertainty of vaccine strategies, extensive studies in animal models will be of great value for optimizing a potential vaccine for HCMV. As mentioned above, in addition to its genomic similarity with HCMV, RhCMV infection of macaques recapitulates HCMV infection of humans. Most likely, the investigation of vaccine approaches against RhCMV infection in macaques may provide translatable data for the design of HCMV vaccine.
RhCMV: A Model for the Study of HCMV
219
Toward this end, a number of studies have been undertaken in rhesus macaques to evaluate the protective efficacy of vaccines consisting of RhCMV homologues of HCMV, such as gB, pp65, and IE1. RhCMV gB and pp65-2 (RhCMV has a duplication of the pp65 locus) (Hansen et al., 2003; Rivailler et al., 2006) resemble their HCMV counterparts in stimulating humoral and cellular responses (Yue et al., 2003, 2006). The first attempt was to evaluate RhCMV gB- and pp65-based DNA vaccines. It has been shown that genetic immunization against the RhCMV pp65-2 and gB antigens stimulated both antigen-specific antibodies, neutralizing antibodies, and CD8 T cell responses. DNA immunization against these viral proteins significantly reduced plasma viral loads and shedding frequency in genital secretions following intravenous challenge with RhCMV 68-1 (Yue et al., 2007). It was also noted in this study that five immunizations of DNA induced only weak neutralizing antibody and CD4þ T cell responses, a result similar to findings with murine cytomegalovirus (MCMV) (Morello et al., 2002). However, a rapid and profound increase of neutralizing titers was observed within one week of intravenous RhCMV challenge, consistent with the interpretation that DNA immunization alone can effectively prime the immune system for neutralizing antibodies, but that a heterologous boost, in this case, in the form of challenge virus, can significantly boost neutralizing titers. This study demonstrates the limited protection conferred by gB and pp65based DNA vaccines and highlights that the extent of protective efficacy can be improved by enhancing vaccine-engendered immunity. Subsequent studies have been conducted in pursuit of such a goal by using heterologous prime/boost strategies. These include a DNA prime, consisting of gB, pp65-2 and supplementing with IE1, and boosting with either recombinant modified vaccinia Ankara virus (rMVA) constructs expressing the same proteins or formalin-inactivated RhCMV virions (FI-RhCMV). rMVA is avirulent in humans and monkeys, including immunodeficient individuals, and is able to efficiently express ectopic gene inserts (Drexler et al., 2004). Using its ability to accommodate foreign gene inserts and its replication-defective phenotype in vivo, rMVA holds potential for the development of an HCMV vaccine. Immunization of mice with rMVA-based HCMV vaccines can elicit both humoral and cellular responses (Wang et al., 2004, 2006, 2007). A pilot study of this approach in rhesus macaques has shown that two immunizations with rMVAs expressing RhCMV gB, pp65-2, and IE1 alone or coupled with a single DNA prime provide similar levels of protection, in terms of reducing the magnitude of plasma viral loads following intravenous RhCMV challenge compared to unvaccinated controls (Yue et al., 2008). Higher immune responses were observed in the animals that received a DNA prime and a single rMVA boost, compared to animals receiving just a single rMVA treatment. No differences were noted, however, after the second rMVA
220
Yujuan Yue and Peter A. Barry
immunization. This study also presented evidence suggesting that higher neutralizing titers at the time of challenge were associated with greater reductions in plasma viral loads following challenge, although this study needs to be substantiated with a larger number of animals. Overall, the results warrant further exploration of the rMVA–CMV vaccines and emphasize the potential protective role of neutralizing antibodies in reducing systemic viral burden during the acute phase of CMV infection. The study of the DNA prime/FI-RhCMV boost approach in rhesus macaques is based on the encouraging data obtained from a vaccine study with MCMV that demonstrated that a higher protective efficacy can be achieved by the production of augmented and broadened humoral and cellular responses through consecutive immunization of mice with DNA vaccines expressing a pool of MCMV antigens and FI-MCMV (Morello et al., 2002). In this study (Abel et al., submitted for publication), rhesus macaques were immunized four times with DNA plasmids expressing RhCMV gB, pp65-2, and IE1 followed by two immunizations with FI-RhCMV formulated in the Montanide ISA 720 water-in-oil adjuvant (Aucouturier et al., 2002). This combined immunization strategy broadened the cellular response by eliciting both antigen-specific CD4þ and CD8þ T cells and biologically relevant neutralizing antibody titers that remained elevated over the 4 weeks between the time of the second FI-RhCMV immunization and time of subcutaneous challenge. Importantly, immunization dramatically reduced the level of viral replication at the primary inoculation sites compared to unvaccinated controls. A vigorous boost in neutralizing antibody titers was observed within one week of challenge, which peaked 3 weeks post challenge. Thereafter, neutralizing titers declined such that by 22 weeks post challenge, the media neutralizing titers were significantly below those of the unvaccinated controls. This latter result is consistent with effective long-term control of virus replication in the vaccinated animals. Taken together, results from these RhCMV vaccine studies provide support for the potential of gB, pp65, and IE1 as vaccine components for human use. Future investigations should focus on defining the respective protective role conferred by elicitation of either neutralizing antibodies and/or T-cell response immunogens and assessing other potential vaccine candidates, which will be of benefit to perceive the key concepts associated with protective immunity and an optimal vaccine.
XI. FUTURE DIRECTIONS Development of protective HCMV vaccines is complicated by a complex viral natural history, continued debate about the nature of the immunogens, and large costs associated with sufficiently powered trials to
RhCMV: A Model for the Study of HCMV
221
measure significant protection. The use of animal models serves as a critical adjunct to evaluate concepts and strategies that can be translated to human critical trials. Each animal model has advantages and disadvantages, but together they comprise an effective preclinical pipeline for vaccine design and testing. This is especially important in light of the complexities involved with a DNA vaccine. Seronegative macaques are available for vaccine studies at the NIH-funded National Primate Research Centers. Juveniles can be serologically screened for RhCMV serostatus to identify seronegative animals, based on prior studies indicating that approximately 50% of juveniles are still seronegative by 6 months. Alternatively, there are increasing efforts to develop breeding cohorts of animals that are specific pathogen free for RhCMV and other herpesviruses (Barry and Strelow, 2008). Since RhCMV is frequently shed from seropositive animals, it should be possible to design vaccine challenge experiments whereby vaccinated and control animals are co-housed with RhCMV-shedding cohorts to rigorously assess protection from infection whereby natural titers of the challenge virus are transmitted by natural routes of exposure at natural frequencies of excretion. Such a challenge scheme would recapitulate an important human route of transmission involving horizontal transmission from HCMV-excreting children to seronegative parents and day care providers.
ACKNOWLEDGEMENTS The authors have been supported by funding from the National Institutes of Health to PAB (AI063356 and AI49342–06) and the California National Primate Research Center (RR000169), and the Margaret M. Deterding Infectious Disease Research Support Fund.
REFERENCES Adler, S. P., Starr, S. E., Plotkin, S. A., Hempfling, S. H., Buis, J., Manning, M. L., and Best, A. M. (1995). Immunity induced by primary human cytomegalovirus infection protects against secondary infection among women of childbearing age. J. Infect. Dis. 171:26–32. Alford, C. A., and Britt, W. J. (1993). Cytomegalovirus. In ‘‘The Human Herpesviruses’’ (B. Roizman, R. J. Whitley, and C. Lopez, eds.), pp. 227–255. Raven Press, New York. Andrade, M. R., Yee, J., Barry, P., Spinner, A., Roberts, J. A., Cabello, P. H., Leite, J. P., and Lerche, N. W. (2003). Prevalence of antibodies to selected viruses in a long-term closed breeding colony of rhesus macaques (Macaca mulatta) in Brazil. Am. J. Primatol. 59 (3):123–128. Arav-Boger, R., Foster, C. B., Zong, J. C., and Pass, R. F. (2006). Human cytomegalovirusencoded alpha-chemokines exhibit high sequence variability in congenitally infected newborns. J. Infect. Dis. 193(6):788–791. Asher, D. M., Gibbs, C. J., Jr, Lang, D. J., Gadjusek, D. C., and Chanock, R. M. (1974). Persistent shedding of cytomegalovirus in the urine of healthy rhesus monkeys. Proc. Soc. Exp. Biol. Med. 145:794–801.
222
Yujuan Yue and Peter A. Barry
Aucouturier, J., Dupuis, L., Deville, S., Ascarateil, S., and Ganne, V. (2002). Montanide ISA 720 and 51: A new generation of water in oil emulsions as adjuvants for human vaccines. Expert Rev. Vaccines 1(1):111–118. Baroncelli, S., Barry, P. A., Capitanio, J. P., Lerche, N. W., Otsyula, M., and Mendoza, S. P. (1997). Cytomegalovirus and simian immunodeficiency virus coinfection: Longitudinal study of antibody responses and disease progression. J. Acquir. Immune Defic. Syndr. 15:5–15. Barry, P. A., and Chang, W.-L. W. (2007). Primate betaherpesviruses. In ‘‘Human Herpesviruses: Biology, Therapy and Immunoprophylaxis’’ (A. Arvin, G. Campadielli, P. Moore, E. Mocarski, B. Roizman, R. Whitley, and K. Yamanishi, eds.), pp. 1051–1075. Cambridge University Press, Cambridge. Barry, P. A., and Strelow, L. (2008). Development of breeding populations of rhesus macaques that are specific pathogen free for rhesus cytomegalovirus. Comp. Med. 58:43–46. Barry, P. A., Alcendor, D. J., Power, M. D., Kerr, H., and Luciw, P. A. (1996). Nucleotide sequence and molecular analysis of the rhesus cytomegalovirus immediate-early gene and the UL121–117 open reading frames. Virology 215:61–72. Barry, P. A., Lockridge, K. M., Salamat, S., Tinling, S. P., Yue, Y., Zhou, S. S., Gospe, S. M., Jr., Britt, W. J., and Tarantal, A. F. (2006). Nonhuman primate models of intrauterine cytomegalovirus infection. ILAR J. 47(1):49–64. Baskin, G. B. (1987). Disseminated cytomegalovirus infection in immunodeficient rhesus macaques. Am. J. Pathol. 129:345–352. Bontrop, R. E., Otting, N., Slierendregt, B. L., and Lanchbury, J. S. (1995). Evolution of major histocompatibility complex polymorphisms and T-cell receptor diversity in primates. Immunol. Rev. 143:33–62. Boppana, S. B., Fowler, K. B., Britt, W. J., Stagno, S., and Pass, R. F. (1999). Symptomatic congenital cytomegalovirus infection in infants born to mothers with preexisting immunity to cytomegalovirus. Pediatrics 104:55–60. Cha, T., Tom, E., Kemble, G., Duke, G., Mocarski, E., and Spaete, R. (1996). Human cytomegalovirus clinical isolates carry at least 19 genes not found in laboratory strains. J. Virol. 70(1):78–83. Chan, K. S., and Kaur, A. (2007). Flow cytometric detection of degranulation reveals phenotypic heterogeneity of degranulating CMV-specific CD8þ T lymphocytes in rhesus macaques. J. Immunol. Methods 325(1–2):20–34. Chang, W. L., and Barry, P. A. (2003). Cloning of the full-length rhesus cytomegalovirus genome as an infectious and self-excisable bacterial artificial chromosome for analysis of viral pathogenesis. J. Virol. 77(9):5073–5083. Chang, W. L., Tarantal, A. F., Zhou, S. S., Borowsky, A. D., and Barry, P. A. (2002). A recombinant rhesus cytomegalovirus expressing enhanced green fluorescent protein retains the wild-type phenotype and pathogenicity in fetal macaques. J. Virol. 76(18):9493–9504. Chang, W. L., Baumgarth, N., Yu, D., and Barry, P. A. (2004). Human cytomegalovirusencoded interleukin-10 homolog inhibits maturation of dendritic cells and alters their functionality. J. Virol. 78(16):8720–8731. Chang, W. L., Baumgarth, N., Eberhardt, M. K., Lee, C. Y., Baron, C. A., Gregg, J. P., and Barry, P. A. (2007). Exposure of myeloid dendritic cells to exogenous or endogenous IL-10 during maturation determines their longevity. J. Immunol. 178(12):7794–7804. Chee, M. S., Bankier, A. T., Beck, S., Bohni, R., Brown, C. M., Cerny, R., Horsnell, T., Hutchison, C. A., Kouzarides, T., Martignetti, J. A., Preddie, E., Satchwell, S. C., et al. (1990). Analysis of the protein-coding content of the sequence of human cytomegalovirus strain AD169. Curr. Top. Microbiol. Immunol. 154:125–169. Chou, S. (1999). Antiviral drug resistance in human cytomegalovirus. Transpl. Infect. Dis. 1(2):105–114.
RhCMV: A Model for the Study of HCMV
223
Conway, M. D., Didier, P., Fairburn, B., Soike, K. F., Martin, L., Murphy-Corb, M., Meiners, N., and Insler, M. S. (1990). Ocular manifestation of simian immunodeficiency syndrome (SAIDS). Curr. Eye Res. 9:759–770. Covell, W. (1932). Occurrence of intranuclear inclusions of monkeys unaccompanied by specific signs of disease. Am. J. Pathol. 8:151–158. Cowdry, W., and Scott, G. (1935). Nuclear inclusions in kidneys of Macacus rhesus monkeys. Am. J. Pathol. 11:659–668. Davison, A. J., Dolan, A., Akter, P., Addison, C., Dargan, D. J., Alcendor, D. J., McGeoch, D. J., and Hayward, G. S. (2003). The human cytomegalovirus genome revisited: Comparison with the chimpanzee cytomegalovirus genome. J. Gen. Virol. 84(Pt. 1):17–28. Dollard, S. C., Grosse, S. D., and Ross, D. S. (2007). New estimates of the prevalence of neurological and sensory sequelae and mortality associated with congenital cytomegalovirus infection. Rev. Med. Virol. 17(5):355–363. Drexler, I., Staib, C., and Sutter, G. (2004). Modified vaccinia virus Ankara as antigen delivery system: How can we best use its potential? Curr. Opin. Biotechnol. 15(6):506–512. Elek, S. D., and Stern, H. (1974). Development of a vaccine against mental retardation caused by cytomegalovirus infection in utero. Lancet 1(7845):1–5. Engel, G. A., Pizzaro, M., Cortes, J., Shaw, E., Fuentes, A., Lerche, N., Barry, P. A., Cohn, D., and Jones-Engel, L. (2008). Seroprevalence of antibodies to enzootic simian viruses among the Gibraltar macaques (M. sylvanus). Emerg. Infect. Dis. 64:1112–1115. Fowler, K. B., Stagno, S., Pass, R. F., Britt, W. J., Boll, T. J., and Alford, C. A. (1992). The outcome of congenital cytomegalovirus infection in relation to maternal antibody status. N. Engl. J. Med. 326:663–667. Gonczol, E., and Plotkin, S. (2001). Development of a cytomegalovirus vaccine: Lessons from recent clinical trials. Expert. Opin. Biol. Ther. 1(3):401–412. Haigwood, N. L. (2004). Predictive value of primate models for AIDS. AIDS Rev. 6 (4):187–198. Hansen, S. G., Strelow, L. I., Franchi, D. C., Anders, D. G., and Wong, S. W. (2003). Complete sequence and genomic analysis of rhesus cytomegalovirus. J. Virol. 77:6620–6636. Hassan-Walker, A. F., Okwuadi, S., Lee, L., Griffiths, P. D., and Emery, V. C. (2004). Sequence variability of the alpha-chemokine UL146 from clinical strains of human cytomegalovirus. J. Med. Virol. 74(4):573–579. Hindle, E., and Stevenson, A. C. (1929–1930). Hitherto undescribed intranuclear bodies in the wild rat and monkeys, compared with known virus bodies in other animals. Trans. R. Soc. Trop. Med. Hyg. 23:327. Huff, J. L., Eberle, R., Capitanio, J., Zhou, S. S., and Barry, P. A. (2003). Differential detection of B virus and rhesus cytomegalovirus in rhesus macaques. J. Gen. Virol. 84:83–92. Jones, B. C., Logsdon, N. J., Josephson, K., Cook, J., Barry, P. A., and Walter, M. R. (2002). Crystal structure of human cytomegalovirus IL-10 bound to soluble human IL-10R1. Proc. Natl. Acad. Sci. USA 99(14):9404–9409. Jones-Engel, L., Engel, G. A., Heidrich, J., Chalise, M., Poudel, N., Viscidi, R., Barry, P. A., Allan, J. S., Grant, R., and Kyes, R. (2006). Temple monkeys and health implications of commensalism, Kathmandu, Nepal. Emerg. Infect. Dis. 12(6):900–906. Karlin, S., Mocarski, E. S., and Schachtel, G. A. (1994). Molecular evolution of herpesviruses: genomic and protein sequence comparisons. J. Virol. 68(3):1886–1902. Kaur, A., Daniel, M. D., Hempel, D., Lee-Parritz, D., Hirsch, M. S., and Johnson, R. P. (1996). Cytotoxic T-lymphocyte responses to cytomegalovirus in normal and simian immunodeficiency virus-infected macaques. J. Virol. 70:7725–7733. Kaur, A., Hale, C. L., Noren, B., Kassis, N., Simon, M. A., and Johnson, R. P. (2002). Decreased frequency of cytomegalovirus (CMV)-specific CD4þ T lymphocytes in simian immunodeficiency virus-infected rhesus macaques: Inverse relationship with CMV viremia. J. Virol. 76(8):3646–3658.
224
Yujuan Yue and Peter A. Barry
Kaur, A., Kassis, N., Hale, C. L., Simon, M., Elliott, M., Gomez-Yafal, A., Lifson, J. D., Desrosiers, R. C., Wang, F., Barry, P., Mach, M., and Johnson, R. P. (2003). Direct relationship between suppression of virus-specific immunity and emergence of cytomegalovirus disease in simian AIDS. J. Virol. 77(10):5749–5758. Kotenko, S. V., Saccani, S., Izotova, L. S., Mirochnitchenko, O. V., and Pestka, S. (2000). Human cytomegalovirus harbors its own unique IL-10 homolog (cmvIL-10). Proc. Natl. Acad. Sci. 97(4):1695–1700. Krosky, P. M., Underwood, M. R., Turk, S. R., Feng, K. W., Jain, R. K., Ptak, R. G., Westerman, A. C., Biron, K. K., Townsend, L. B., and Drach, J. C. (1998). Resistance of human cytomegalovirus to benzimidazole ribonucleosides maps to two open reading frames: UL89 and UL56. J. Virol. 72(6):4721–4728. Kuhn, E. M., Stolte, N., Matz-Rensing, K., Mach, M., Stahl-Henning, C., Hunsmann, G., and Kaup, F. J. (1999). Immunohistochemical studies of productive rhesus cytomegalovirus infection in rhesus monkeys (Macaca mulatta) infected with simian immunodeficiency virus. Vet. Pathol. 36(1):51–56. Landolfo, S., Gariglio, M., Gribaudo, G., and Lembo, D. (2003). The human cytomegalovirus. Pharmacol. Ther. 98(3):269–297. Lockridge, K. M., Sequar, G., Zhou, S. S., Yue, Y., Mandell, C. M., and Barry, P. A. (1999). Pathogenesis of experimental rhesus cytomegalovirus infection. J. Virol. 73:9576–9583. Lockridge, K. M., Zhou, S. S., Kravitz, R. H., Johnson, J. L., Sawai, E. T., Blewett, E. L., and Barry, P. A. (2000). Primate cytomegaloviruses encode and express an IL-10-like protein. Virology 268(2):272–280. London, W. T., Martinez, A. J., Houff, S. A., Wallen, W. C., Curfman, B. L., Traub, R. G., and Sever, J. L. (1986). Experimental congenital disease with simian cytomegalovirus in rhesus monkeys. Teratology 33:323–331. Lurain, N. S., Fox, A. M., Lichy, H. M., Bhorade, S. M., Ware, C. F., Huang, D. D., Kwan, S. P., Garrity, E. R., and Chou, S. (2006). Analysis of the human cytomegalovirus genomic region from UL146 through UL147A reveals sequence hypervariability, genotypic stability, and overlapping transcripts. Virol. J. 3:4. McCormick, A. L., Skaletskaya, A., Barry, P. A., Mocarski, E. S., and Goldmacher, V. S. (2003). Differential function and expression of the viral inhibitor of caspase 8-induced apoptosis (vICA) and the viral mitochondria-localized inhibitor of apoptosis (vMIA) cell death suppressors conserved in primate and rodent cytomegaloviruses. Virology 316:221–233. McGeoch, D. J., Cook, S., Dolan, A., Jamieson, F. E., and Telford, E. A. (1995). Molecular phylogeny and evolutionary timescale for the family of mammalian herpesviruses. J. Mol. Biol. 247(3):443–458. McGeoch, D. J., Rixon, F. J., and Davison, A. J. (2006). Topics in herpesvirus genomics and evolution. Virus Res. 117(1):90–104. Medicine, I. O. (2000). ‘‘Vaccines for the 21st Century.’’ p. 6. National Academy Press, Washington, DC. Mocarski, E. S., Jr. (2002). Immunomodulation by cytomegaloviruses: Manipulative strategies beyond evasion. Trends Microbiol. 10(7):332–339. Morello, C. S., Ye, M., and Spector, D. H. (2002). Development of a vaccine against murine cytomegalovirus (MCMV), consisting of plasmid DNA and formalin-inactivated MCMV, that provides long-term, complete protection against viral replication. J. Virol. 76(10):4822–4835. Murphy, E., Rigoutsos, I., Shibuya, T., and Shenk, T. E. (2003). Reevaluation of human cytomegalovirus coding potential. Proc. Natl. Acad. Sci. USA 100:14976–14981. Murphy, E., Yu, D., Grimwood, J., Schmutz, J., Dickson, M., Jarvis, M. A., Hahn, G., Nelson, J. A., Myers, R. M., and Shenk, T. E. (2003). Coding potential of laboratory and clinical strains of human cytomegalovirus. Proc. Natl. Acad. Sci. USA 100:14976–14981.
RhCMV: A Model for the Study of HCMV
225
Neff, B. J., Weibel, R. E., Buynak, E. B., McLean, A. A., and Hilleman, M. R. (1979). Clinical and laboratory studies of live cytomegalovirus vaccine Ad-169. Proc. Soc. Exp. Biol. Med. 160(1):32–37. North, T. W., Sequar, G., Townsend, L. B., Drach, J. C., and Barry, P. A. (2004). Rhesus cytomegalovirus is similar to human cytomegalovirus in susceptibility to benzimidizole nucleosides. Antimicrob. Agents Chemother 48:2760–2765. Novak, Z., Ross, S. A., Patro, R. K., Pati, S. K., Kumbla, R. A., Brice, S., and Boppana, S. B. (2008). Cytomegalovirus strain diversity in seropositive women. J. Clin. Microbiol. 46(3):882–886. Oxford, K. L., Eberhardt, M. K., Yang, K. W., Strelow, L., Kelly, S., Zhou, S. S., and Barry, P. A. (2008). Protein coding content of the U(L)b0 region of wild-type rhesus cytomegalovirus. Virology 373(1):181–188. Pande, N. T., Powers, C., Ahn, K., and Fruh, K. (2005). Rhesus cytomegalovirus contains functional homologues of US2, US3, US6, and US11. J. Virol. 79(9):5786–5798. Pass, R. F. (2001). Cytomegalovirus. In ‘‘Fields Virology’’ (D. Knipe and P. Howley, eds.), pp. 2675–2705. Lippincott, Williams and Wilkins, Philadelphia. Pearson, T. C., Trambley, J., Odom, K., Anderson, D. C., Cowan, S., Bray, R., Lin, A., Hollenbaugh, D., Aruffo, A., Siadak, A. W., Strobert, E., Hennigar, R., et al. (2002). AntiCD40 therapy extends renal allograft survival in rhesus macaques. Transplantation 74 (7):933–940. Pignatelli, S., Dal Monte, P., Rossini, G., and Landini, M. P. (2004). Genetic polymorphisms among human cytomegalovirus (HCMV) wild-type strains. Rev. Med. Virol. 14 (6):383–410. Plotkin, S. A. (1999). Cytomegalovirus vaccine. Am. Heart J. 138(5 Pt. 2):S484–S487. Plotkin, S. A. (2001). Vaccination against cytomegalovirus. Arch. Virol. Suppl. 17:121–134. Powers, C., and Fru¨h, K. (2008). Rhesus CMV: An emerging animal model for human CMV. Med. Microbiol. Immunol. 197:109–115. Prichard, M. N., Penfold, M. E., Duke, G. M., Spaete, R. R., and Kemble, G. W. (2001). A review of genetic differences between limited and extensively passaged human cytomegalovirus strains. Rev. Med. Virol. 11(3):191–200. Rivailler, P., Kaur, A., Johnson, R. P., and Wang, F. (2006). Genomic sequence of rhesus cytomegalovirus 180.92: Insights into the coding potential of rhesus cytomegalovirus. J. Virol. 80(8):4179–4182. Sequar, G., Britt, W. J., Lakeman, F. D., Lockridge, K. M., Tarara, R. P., Canfield, D. R., Zhou, S. S., Gardner, M. B., and Barry, P. A. (2002). Experimental coinfection of rhesus macaques with rhesus cytomegalovirus and simian immunodeficiency virus: Pathogenesis. J. Virol. 76(15):7661–7671. Shanley, J. D., Morningstar, J., and Jordan, M. C. (1985). Inhibition of murine cytomegalovirus lung infection and interstitial pneumonitis by acyclovir and 9-(1,3-dihydroxy2-propoxymethyl)guanine. Antimicrob. Agents Chemother. 28(2):172–175. Spencer, J. V., Lockridge, K. M., Barry, P. A., Lin, G., Tsang, M., Penfold, M. E., and Schall, T. J. (2002). Potent immunosuppressive activities of cytomegalovirus-encoded interleukin-10. J. Virol. 76(3):1285–1292. Staczek, J. (1990). Animal cytomegaloviruses. Microbiol. Rev. 54(3):247–265. Stagno, S. (1990). Cytomegalovirus.. In ‘‘Infectious Diseases of the Fetus and Newborn Infant’’ (J. S. Remington and J. O. Klein, eds.), 3rd Ed., pp. 241–281. W. B. Saunders, Philadelphia. Stagno, S., Pass, R. F., Cloud, G., Britt, W. J., Henderson, R. E., Walton, P. D., Veren, D. A., Page, F., and Alford, C. A. (1986). Primary cytomegalovirus infection in pregnancy: Incidence, transmission to fetus, and clinical outcome. J. Am. Med. Assoc. 256:1904–1908. Stewart, F. W., and Rhoads, C. P. (1929). Lesions in nasal mucous membranes of monkeys with acute poliomyelitis. Proc. Soc. Exp. Biol. Med. 26:664–665.
226
Yujuan Yue and Peter A. Barry
Stratton, K. R., Durch, J. S., and Lawrence, R. S. (2000). ‘‘Vaccines for the 21st Century: A Tool for Decision making.’’ National Academy Press, Washington, DC. Swack, N. S., and Hsiung, G. D. (1982). Natural and experimental simian cytomegalovirus infections at a primate center. J. Med. Primatol. 11:169–177. Swack, N. S., Liu, O. C., and Hsiung, G. D. (1971). Cytomegalovirus infections of monkeys and baboons. Am. J. Epidemiol. 94:397–402. Swanson, R., Bergquam, E., and Wong, S. W. (1998). Characterization of rhesus cytomegalovirus genes associated with anti-viral susceptibility. Virology 240(2):338–348. Tarantal, A. F., Salamat, S., Britt, W. J., Luciw, P. A., Hendrickx, A. G., and Barry, P. A. (1998). Neuropathogenesis induced by rhesus cytomegalovirus in fetal rhesus monkeys (Macaca mulatta). J. Infect. Dis. 177:446–450. Townsend, L. B., Devivar, R. V., Turk, S. R., Nassiri, M. R., and Drach, J. C. (1995). Design, synthesis, and antiviral activity of certain 2,5,6-trihalo-1-(beta-D-ribofuranosyl)benzimidazoles. J. Med. Chem. 38(20):4098–4105. Underwood, M. R., Harvey, R. J., Stanat, S. C., Hemphill, M. L., Miller, T., Drach, J. C., Townsend, L. B., and Biron, K. K. (1998). Inhibition of human cytomegalovirus DNA maturation by a benzimidazole ribonucleoside is mediated through the UL89 gene product. J. Virol. 72(1):717–725. Villarreal, E. C. (2003). Current and potential therapies for the treatment of herpes-virus infections. Prog. Drug Res. 60:263–307. Vogel, P., Weigler, B. J., Kerr, H., Hendrickx, A., and Barry, P. A. (1994). Seroepidemiologic studies of cytomegalovirus infection in a breeding population of rhesus macaques. Lab. Anim. Sci. 44:25–30. Wang, Z., La Rosa, C., Maas, R., Ly, H., Brewer, J., Mekhoubad, S., Daftarian, P., Longmate, J., Britt, W. J., and Diamond, D. J. (2004). Recombinant modified vaccinia virus Ankara expressing a soluble form of glycoprotein B causes durable immunity and neutralizing antibodies against multiple strains of human cytomegalovirus. J. Virol. 78(8):3965–3976. Wang, Z., La Rosa, C., Lacey, S. F., Maas, R., Mekhoubad, S., Britt, W. J., and Diamond, D. J. (2006). Attenuated poxvirus expressing three immunodominant CMV antigens as a vaccine strategy for CMV infection. J. Clin. Virol. 35(3):324–331. Wang, Z., La Rosa, C., Li, Z., Ly, H., Krishnan, A., Martinez, J., Britt, W. J., and Diamond, D. J. (2007). Vaccine properties of a novel marker gene-free recombinant modified vaccinia Ankara expressing immunodominant CMV antigens pp65 and IE1. Vaccine 25 (6):1132–1141. Yue, Y., Zhou, S. S., and Barry, P. A. (2003). Antibody responses to rhesus cytomegalovirus glycoprotein B in naturally infected rhesus macaques. J. Gen. Virol. 84(Pt. 12):3371–3379. Yue, Y., Kaur, A., Zhou, S. S., and Barry, P. A. (2006). Characterization and immunological analysis of the rhesus cytomegalovirus homologue (Rh112) of the human cytomegalovirus UL83 lower matrix phosphoprotein (pp65). J. Gen. Virol. 87(Pt. 4):777–787. Yue, Y., Kaur, A., Eberhardt, M. K., Kassis, N., Zhou, S. S., Tarantal, A. F., and Barry, P. A. (2007). Immunogenicity and protective efficacy of DNA vaccines expressing rhesus cytomegalovirus glycoprotein B, phosphoprotein 65–2, and viral interleukin-10 in rhesus macaques. J. Virol. 81(3):1095–1109. Yue, Y., Wang, Z., Abel, K., Li, J., Strelow, L., Mandarino, A., Eberhardt, M. K., Schmidt, K. A., Diamond, D. J., and Barry, P. A. (2008). Evaluation of recombinant modified vaccinia Ankara virus-based rhesus cytomegalovirus vaccines in rhesus macaques. Med. Microbiol. Immunol. 197:117–123. Zhong, J., and Khanna, R. (2007). Vaccine strategies against human cytomegalovirus infection. Expert Rev. Anti Infect. Ther. 5(3):449–459.
CHAPTER
6 Drosophila Viruses and the Study of Antiviral Host-Defense Tu¨nde Huszar*,† and Jean-Luc Imler*
Contents
Abstract
I. Introduction II. Sigma Virus A. Description of the virus, relationship to other rhabdoviridae B. Interaction with D. melanogaster III. Drosophila C Virus (DCV) A. Description of the virus B. Interaction with drosophila IV. Other Drosophila Viruses A. Drosophila X virus (DXV) B. Drosophila F virus (DFV) C. Other RNA viruses D. Gypsy and infectious retrotransposons V. Antiviral Reactions in Drosophila A. RNA interference B. Inducible response to infection VI. Conclusion and Perspectives Acknowledgments References
228 229 229 234 236 236 239 240 240 243 244 246 249 250 254 256 258 258
The fruit fly Drosophila melanogaster is a powerful model to study host-pathogen interactions. Most studies so far have focused on extracellular pathogens such as bacteria and fungi. More recently, viruses have come to the front, and RNA interference was shown to play a critical role in the control of viral infections in drosophila.
* CNRS UPR 9022, Institut de Biologie Mole´culaire et Cellulaire, Strasbourg, France {
Department of Genetics, University of Szeged, Hungary
Advances in Virus Research, Volume 72 ISSN 0065-3527, DOI: 10.1016/S0065-3527(08)00406-5
#
2008 Elsevier Inc. All rights reserved.
227
228
Tu¨nde Huszar and Jean-Luc Imler
We review here our current knowledge on drosophila viruses. A diverse set of RNA viruses belonging to several families (Rhabdoviridae, Dicistroviridae, Birnaviridae, Reoviridae, Errantiviridae) has been reported in D. melanogaster. By contrast, no DNA virus has been recovered up to now. The drosophila viruses represent powerful tools to study virus-cell interactions in vivo. Analysis of the literature however reveals that for many of them, important gaps exist in our understanding of their replication cycle, genome organization, morphology or pathogenesis. The data obtained in the past few years on antiviral defense mechanisms in drosophila, which point to evolutionary conserved pathways, highlight the potential of the D. melanogaster model to study antiviral innate immunity and to better understand the complex interaction between arthropod-borne viruses and their insect vectors.
I. INTRODUCTION Like all organisms, invertebrates are plagued by viruses. Insect viruses have probably existed for as long as insect themselves, and have long been of interest to humans. Some of these viruses are of great concern because they threaten beneficial insects, such as honeybees, or human enterprises, such as silkworm industry. In addition, over 500 known varieties of arboviruses (arthropod-borne viruses) can efficiently infect and replicate in cells from both invertebrate and vertebrate hosts. Examples include members of the genus Flavivirus, such as yellow fever virus, dengue viruses, and West Nile virus. These viruses each require a blood sucking insect, the mosquito Aedes, to complete their life cycle. As exemplified in recent years for the West Nile virus in the United States, these viruses provide a spectacular example of emerging diseases of global significance (Geisbert and Jahrling, 2004). The fruit fly Drosophila melanogaster has been a favorite model of biologists since the beginning of the twentieth century. Studies conducted in this small dipteran insect have led to major discoveries in genetics, embryology, cellular and molecular biology (Rubin and Lewis, 2000). More recently, drosophila has become a popular model to decipher host-pathogen interactions (Lemaitre and Hoffmann, 2007). So far, most studies have focused on the response of drosophila to bacterial or fungal infections, and the results obtained have led to the identification of evolutionary conserved mechanisms of host defense (most spectacularly the identification of Toll-like receptors as important regulators of innate immunity), but also of molecules regulating transmission of insect borne diseases (e.g., control of the Plasmodium parasite in Anopheles mosquitoes by the complement-related molecule TEP1)(Blandin and Levashina, 2004; Ferrandon et al., 2007; Lemaitre and Hoffmann, 2007). In recent years,
Drosophila Viruses and the Study of Antiviral Host-Defense
229
interest has shifted to the study of antiviral defenses in drosophila. Although it is possible to use exogenous viruses to some extent, natural viral pathogens of D. melanogaster represent important tools to perform these studies. We review here the current state of knowledge on drosophila viruses, and how these viruses have been used to begin to address the cellular and molecular basis of resistance to virus infection.
II. SIGMA VIRUS Sigma virus (SIGMAV) is widespread in natural populations of drosophila and is one of the best-characterized virus infecting fruit-flies. SIGMAV is a member of the Rhabdoviridae, an important family of RNA viruses infecting animals and plants. These economically important viruses cause infections of crops and cattle, and are also an important threat for human health (e.g., rabies) (Hogenhout et al., 2003).
A. Description of the virus, relationship to other rhabdoviridae 1. Virion and genome structure Rhabdoviruses are enveloped RNA viruses. Their genome is composed of a single 11–15 kb single stranded RNA molecule, which is of negative polarity (ss() RNA), meaning that it has to be transcribed in infected cells before viral proteins can be translated (Table I). The RNA has a 50 -terminal triphosphate and is not polyadenylated. Its extremities contain inverted complementary sequences. The most common animal rhabdoviruses form two genera, the vesiculoviruses (Vesicular Stomatitis Virus (VSV)-like) and lyssaviruses (rabies-like). Prototypes from each genus have five genes, known as N, P, M, G, and L (from the 30 to 50 end), which encode structural and nucleocapsid proteins (Hogenhout et al., 2003; Rose and Whitt, 2001). In the virion, the RNA genome is tightly associated with the N protein. In the best studied rhabdovirus, VSV, about 1200 molecules of N associate with the RNA genome, like beads on a string, forming a tightly packed helix with a 30–70 nm diameter. The proteins encoded by the genes L and P are also associated with the nucleocapsid, each virion containing about 50 L and 500 P molecules (Fig. 1). The L protein is the RNA-dependent RNA polymerase, which mediates transcription of viral genes, as well as viral replication, in association with the P protein. The P phosphoprotein promotes interaction between the L polymerase and viral RNAs, and plays an important accessory role in the regulation of viral RNA synthesis. The P protein from rabies virus also modulates the host-defense system, by inhibiting the nuclear import of the STAT1 transcription factor (Brzozka et al., 2006; Vidy et al., 2007).
TABLE I
Drosophila viruses
Name
Family
Genome
Virion
Transmission
SIGMAV
Rhabdoviridae
RNA ss () strand; 10–15 kb?
Vertical
DCV
Discistroviridae
DXV
Birnaviridae
DFV
Reoviridae
RNA ss (þ) strand; 9264 nt, 50 VPg linked RNA ds; 2 segments A (3360 bp) and B (3243 bp), 50 VPg linked RNA ds; 10 segments
Enveloped, bullet-shape 45 nm , 100 nm length Non enveloped, 25–30 nm Non enveloped 70 nm
Gypsy
Errantiviridae
RNA ss (þ) strand; 7469 nt
Nora DPV
Unclassified Unclassified
RNA ss (þ) strand; 11879 nt RNA ss (þ) strand
Non enveloped 60–70 nm Enveloped 45 nm Non enveloped Non enveloped, 27–30 nm
Horizontal Horizontal ? Horizontal & Vertical ? Vertical
Drosophila Viruses and the Study of Antiviral Host-Defense
231
A
5⬘
3⬘ N
P
3 ?
M
G
L
B
FIGURE 1 The SIGMA virus. (A) schematic representation of the SIGMAV virion and genome. (B) Electron micrograph of a SIGMAV particle. Scale bar: 100 nm. Picture courtesy of D. Contamine.
The external membrane surrounding the nucleocapsid has a composition similar to host cells plasma membranes, from which it derives by budding. The interaction between the nucleocapsid and the viral envelope is mediated by some 1800 viral matrix protein, encoded by the gene M. M is the smallest and most abundant protein in the virion and it may carry additional functions apart from its important role in virion assembly. Indeed, in the case of VSV, this protein has been shown to inhibit hostcell transcription in the nucleus and in rabies virus infected cells, M binds to the factor eIF3 and inhibits host cell translation (Komarova et al., 2007). The virion has a characteristic bullet- or cone-shape, with a diameter of 60–80 nm and a length of 180–200 nm. The outer surface of the envelope is decorated by some 400 spikes formed by trimers of the glycoprotein encoded by the gene G, which cover the whole surface of the virion, at the notable exception of the quasi-planar end. Both the M and G proteins are required for virion assembly and budding out of the cells (Rose and Whitt, 2001).
232
Tu¨nde Huszar and Jean-Luc Imler
The genome of SIGMAV has not been fully sequenced yet, nor has its replication cycle been studied with great detail in drosophila cells. SIGMAV has the characteristic bullet shape of the members of the Rhabdoviridae family (Berkaloff et al., 1965) (Fig. 1B). The sequence of a total of 6.5 kb from its genome shows a genomic organization similar to other members of the family, with the notable difference that six genes (instead of five) are observed (Teninges et al., 1993). The 6th gene, gene 3, is inserted between the P and the M gene, and the gene order in SIGMAV is 30 -NP-3-M-G-L-50 (Fig. 1A). Sequence comparison of the N gene suggests that SIGMAV occupies an intermediate evolutionary position between vesiculoviruses and lyssaviruses (Bras et al., 1994). The function of the protein encoded by gene 3 remains mysterious at this stage. As a matter of fact, even though the gene is expressed at high levels, attempts to detect the protein failed. The putative protein product of this SIGMAV-specific gene does not exhibit significant homology to any protein sequences encoded by Mononegavirales. Sequence homology searches revealed the presence of weak similarities with RNA-dependent DNA-polymerases from retroviruses or retrotransposons (Landes-Devauchelle et al., 1995). The function of SIGMAV gene 3 may be related to the adaptation of the virus to hereditary transmission in drosophila flies, which implies that the virus does not interfere with development processes. Of note, two other rhabdoviruses (rabies virus and Infectious Hematopoietic Necrosis Virus or IHNV) contain a sixth gene intercalated between G and L. In the case of the fish rhabdovirus IHNV, this gene codes the NV protein, which is involved in viral pathogenesis (Thoulouze et al., 2004).
2. Viral replication cycle The replication cycle of rhabdoviruses can be divided in several stages, entry and uncoating, transcription, replication and finally assembly and budding (reviewed in Rose and Whitt, 2001). The first step in the infection is the binding of the virion to the plasma membrane. This adsorption step may involve phospholipids or intrinsic membrane proteins, and is followed by clathrin-mediated endocytosis. The reduction of the pH in the endosomes triggers a membrane fusion reaction between the endosomal membrane and the envelope of the virion. The G protein plays a critical role in the membrane fusion event, which is accompanied by the rapid release of the ribonucleoprotein (RNP) core in the cytosol, and the dissociation of the M protein. Upon release in the cytosol, the negative-strand RNA viral genome is rapidly transcribed by the L polymerase associated with its cofactor P, which are part of the RNP. Transcription begins at the 30 end of the genome, where the polymerase first synthesizes a small leader RNA, and then proceeds to the synthesis of the other mRNAs encoding the viral proteins, in the order they appear from the 30 end of the genome. The data available
Drosophila Viruses and the Study of Antiviral Host-Defense
233
account for a model whereby the polymerase terminates transcription at the leader-N gene junction, and reinitiates at a conserved site (30 GUUGU[A/U]G-50 for SIGMAV). Transcription terminates at the conserved stop signal (30 -GUACUUUUUUU-50 for SIGMAV), which triggers polyadenylation by reiterative copying of the seven U residues (Landes-Devauchelle et al., 1995; Rose and Whitt, 2001; Teninges et al., 1993). The stop-polyadenylation signal can occasionally be by-passed to yield bicistronic mRNAs, as reported for genes P and 3 in SIGMAV (Teninges et al., 1993). When the polyadenylation is over, the polymerase scans the intergenic region until the next transcription initiation site. Premature dissociation of the polymerase in intergenic regions presumably accounts for the reduced expression of downstream genes. The main difference between transcription and replication of the viral genome is that the signal sequences that regulate the termination and reinitiation of RNA synthesis during transcription are ignored by the polymerase during replication, thus enabling the synthesis of a complete genomic RNA of positive polarity. This RNA, known as the replicative intermediate (RI), is then copied to form the genomic negative strand RNA. Differences in the sequences promoting replication at the 50 and 30 end of the genome explains the higher efficiency of synthesis of the genome over the antigenome, and accounts for 20- to 50-fold excess of genome versus RI in infected cells. Thus, the L polymerase can act both as a transcriptase and a replicase. The differentiation between these two functions remains a matter of controversy (Banerjee, 2008; Curran and Kolakofsky, 2008; Whelan, 2008). In the case of VSV, it reflects the existence of two distinct RNA polymerase complexes: in the transcriptase complex, L is complexed with P proteins, and phosphorylation of the aminoterminal I domain of P is essential for RNA synthesis (Pattnaik et al., 1997), whereas in the replicase complex, L associates with newly synthesized N protein and phosphorylation of the C-terminal domain II of P is important for activity (Hwang et al., 1999). An important aspect of replication is the coupling of RNA synthesis with encapsidation by N protein, which initiates at the 50 terminal regions of the genomic and the RI RNA molecules. Hence, RNA molecules produced by the replicase complex tightly associate with the N protein as they are elongated. After encapsidation, the RNP complex formed by the genomic RNA and the newly synthesized N, P, and L proteins associates with the M protein. This association triggers condensation of the RNP, and promotes association with the plasma membrane. Localization of the RNP below the plasma membrane initiates the budding process, whereby the nucleocapsid will become enveloped and released from the cell. The membrane envelope contains some 1200 molecules of the viral G glycoprotein, that associates with components of the RNP through its
234
Tu¨nde Huszar and Jean-Luc Imler
cytoplasmic tail (reviewed in (Hogenhout et al., 2003; Rose and Whitt, 2001). It will be important in future studies to verify that the replication cycle of SIGMAV in drosophila cells is similar to that of the better studied VSV and rabies virus.
B. Interaction with D. melanogaster Rhabdoviruses have a broad host-range, and most have two natural hosts, either insect and plant or insect and vertebrate (Hogenhout et al., 2003). Insects therefore play a central role in the horizontal transmission of these viruses. It is likely that insects were the primary hosts for the rhabdovirus ancestors, which later acquired the ability to infect secondary hosts. SIGMAV is atypical, in that it has no known vertebrate or plant hosts, and only infects drosophila. SIGMAV is widespread in natural populations of drosophila, and flies infected with the virus suffer few adverse effects, including reduced viability of infected eggs and lower survival over winter (Fleuriet, 1981a,b). In fact, the readout used in the laboratory to monitor infection is the sensitivity to exposure to pure CO2, a treatment routinely used for brief anesthesia of the flies. This artificial treatment has a dramatic effect on SIGMAV infected flies, which become irreversibly paralyzed, possibly as a result of viral proliferation in the central nervous system (L’heritier and Teissier, 1937). Uninfected flies survive this treatment. In agreement with the mild effect of the virus on the fly, the virus is not cytopathogenic in drosophila tissue-culture cells, and establishes persistent infections (Echalier, 1997; Ohanessian and Echalier, 1967). Five host loci are involved in the control of SIGMAV infection, such as ref(1)H, ref(2)P and ref(3)D (Gay, 1978). The best characterized is ref(2)P, a strongly polymorphic gene from the second chromosome (Contamine et al., 1989; Dru et al., 1993; Wayne et al., 1996). Ref(2)P is an evolutionary conserved gene, which encodes a protein containing three domains: PB1 (Phox and Bem1p), ZZ (atypical zinc finger) and UBA (ubiquitin-associated), from N- to C-terminus (CarreMlouka et al., 2007). The mammalian orthologue of Ref(2)P, known as p62, is a protein adaptor involved in the activation of the NF-kB pathway by members of the atypical protein kinase C subfamily (aPKC)(Moscat et al., 2007). In drosophila cells, Ref(2)P and the drosophila aPKC (DaPKC) have been proposed to function in the Toll pathway, and to regulate the NF-kB proteins Dorsal and DIF (Avila et al., 2002). Natural populations of flies contain two types of alleles (Contamine et al., 1989): permissive alleles of ref(2)P allow efficient SIGMAV multiplication, whereas restrictive alleles limit the replication of the virus. The probability of infection, which may reach 100% in a permissive context, drops to 0.01% in a restrictive context, at least for some viral strains (Gay, 1978).
Drosophila Viruses and the Study of Antiviral Host-Defense
235
Indeed, the ability of a restrictive allele to contain the infection depends of the viral strain, and two viral genotypes can be distinguished according to their capacity to infect flies that have the restrictive ref(2)P allele: infective strains can infect these flies, whereas avirulent strains can only infect flies having the permissive allele (Contamine, 1981; Fleuriet, 1999, 2001; Fleuriet and Periquet, 1993). Evolutionary genomics studies suggest that the restrictive allele appeared several thousands of years ago, and spread in the population as a result of the selective advantage it confers (Bangham et al., 2007). Sequence analysis of SIGMAV isolates from Europe and North America revealed a much lower level of diversity that seen for other RNA viruses, pointing to a common ancestor around 200 years ago (Carpenter et al., 2007). This common ancestor might be the first variant that successfully infected flies, possibly upon contamination by mites or parasitic wasps. Alternatively, the low genetic diversity of SIGMAV isolates might result from a selective sweep associated with a better adaptation to drosophila as a host. The appearance of the viral strain able to infect flies with the restrictive ref(2)P allele probably occurred much more recently (25 years), and rapidly spread across Europe (Carpenter et al., 2007). While ref(2)P does not appear to be essential for development of drosophila, it is required for male fertility, although the molecular mechanism involving Ref(2)P is not known(Contamine et al., 1989; Dezelee et al., 1989). Similarly, the mechanism by which Ref(2)P interacts with SIGMAV remains unclear. One important genetic observation is that flies containing a permissive allele of ref(2)P are more susceptible to infection than flies that are deficient for the gene, suggesting that SIGMAV uses the permissive allele to infect flies (Carre-Mlouka et al., 2007). An alternative explanation for these data might be a general unspecific decreased resistance to SIGMAV infection as a result of altered cellular and/or physiological functions in flies expressing the permissive allele. Interestingly, the mutations associated with the restrictive phenotype were recently shown to map to the PB1 domain of the protein. Of note, this domain contains the atypical PKC interaction domain (AID). However, the three mutations associated with the restrictive/permissive phenotype are located in the N-terminus of the PB1 domain, upstream of the AID motif (Carre-Mlouka et al., 2007). In addition, the dominant relationship between restrictive and permissive alleles varies according to the viral strain used, and one of the three restrictive mutations is virus strain specific. These genetic data suggest a direct interaction between Ref(2)P and a viral protein. Ref(2)P may for example function as a scaffolding protein during assembly of viral protein complexes. Some evidence for this scenario exists, since Ref(2)P has been shown to share conformationdependent epitopes with the SIGMAV N protein, and to interact with the P protein (Wyers et al., 1993).
236
Tu¨nde Huszar and Jean-Luc Imler
III. DROSOPHILA C VIRUS (DCV) DCV is the best studied drosophila virus. It was first reported in 1972 in a laboratory stock that exhibited unusually high and unexplained lethality. The virus was found to be similar in size and morphology to the previously characterized P and iota viruses, from which it differed serologically and by its high virulence, killing flies in as little as three days instead of 15 days (Jousset et al., 1972). The virus was later found in several laboratory stocks, and from wild populations of D. melanogaster (Gateff et al., 1980; Plus et al., 1975b). It belongs to the Dicistroviridae family, genus Cripavirus, together with many other insect viruses (e.g., Aphid Lethal Paralysis Virus in Hemiptera, Black Queen Cell Virus from Hymenoptera and Cricket Paralysis Virus from Lepidoptera)(Christian et al., 2000). One of them, Cricket Paralysis Virus (CrPV), replicates efficiently in drosophila cells and is pathogenic when injected into flies (Moore et al., 1980; Wang et al., 2006).
A. Description of the virus 1. Virion and genome structure DCV is a non-enveloped RNA virus that shares many properties with picornaviruses (Fig. 2). It was considered an insect picornavirus until its genome was sequenced and published in 1998. Among the similarities with picornaviruses, (1) the icosaedral viral particles have a 25–30 nm diameter, and a buoyant density of 1.34g/ml (Fig. 2B); (2) the capsid is composed of the three major proteins VP1 (33 kDa), VP2 (29 kDa) and VP3 (28 kDa); viral proteins of 37 kDa (VP0) and 8,5 kDa (VP4) are also detected in lesser amounts in DCV particles, VP0 being the precursor for VP3 and VP4 (VP2 and VP4 in picornaviruses, see Fig. 2A); (3) a virally encoded protease processes the viral proteins from large polyprotein precursors; (4) the genome is a single stranded positive strand RNA molecule (ss(þ)RNA), which is linked at its 50 end to a viral VPg protein, and polyadenylated at its 30 end (Jousset et al., 1977; King and Moore, 1988) (Table I). DCV also resembles Picornaviridae by many aspects of its replication cycle (see below). It therefore came as a surprise that the genome of DCV revealed significant differences with that of Picornaviridae (Johnson and Christian, 1998). The genome of picornaviruses contains a single ORF, with the sequences coding the capsid proteins at the 50 end, and the non-structural proteins (helicase, proteases, and RNA-dependent RNA polymerase) at the 30 end. By contrast, the 9264 nt DCV genome contains two ORFs. ORF1 is located at the 50 end and encodes a 202 kDa polyprotein containing the domains for the replication machinery, arranged in the same order as in
Drosophila Viruses and the Study of Antiviral Host-Defense
237
A Picornavirus VP4 VP2 VP3
VP1 Hel
5⬘VPg
IRES
Pro
RdRP
An
3⬘
An
3⬘
An
3⬘
Non structural proteins
Iflavirus L
VP2 VP4
VP3
VP1 Hel
5⬘VPg
Pro
RdRP
Non structural proteins
IRES Dicistrovirus
VP2 VP4
L Hel
5⬘VPg
IRES 1
Pro
RdRP
VP3
VP1
IRES 2 Non structural proteins
B
FIGURE 2 Drosophila C virus. (A) schematic representation of the genome structure of different types of picorna-like viruses. Hel: helicase; Pro: protease; RdRP: RNA-dependent RNA polymerase. (B) Purified particles of DCV. Scale bar: 100 nm.
picornaviruses (Fig. 2A). The 30 ORF2 encodes a 100 kDa polyprotein from which the capsid proteins are produced. Other differences with members of the Picornaviridae are the absence of one of the two proteases (2A), and of the 2B and 3A proteins. DCV from then on became the prototype
238
Tu¨nde Huszar and Jean-Luc Imler
member of a new family of RNA viruses, the Dicistroviridae, which belongs to the order Picornavirales (Johnson and Christian, 1999; Le Gall et al., 2008). No subgenomic RNA is produced and translation of both ORFs proceeds from the genome-long bicistronic RNA, using two internal ribosomal entry sites (IRES) located at the 50 end of the genome for ORF1, and in the 191 nt intergenic region for ORF2 (Sasaki and Nakashima, 2000; Wilson et al., 2000). The intergenic IRES can directly assemble 80S ribosomes in the absence of canonical translation initiation factors and initiator tRNA, unlike the poliovirus IRES (Schuler et al., 2006; Spahn et al., 2004). It is more active than the IRES located at the 50 end of the RNA molecule, thus explaining the abundance of the structural VP proteins compared to the non-structural proteins in infected cells (Wilson et al., 2000). The crystal structure of CrPV has been solved, providing interesting structural informations about this new family of viruses (Tate et al., 1999). The proteins VP1, VP2, and VP3 are arranged in a pseudo T ¼ 3 lattice to form the capsid. VP1, -2 and -3 each have a b-barrel core, a property shared with the structural proteins of picornaviruses. The smaller VP4 protein is located on the interior of the virion, at the interface between the protein capsid and the RNA genome. Unlike VP4 from picornaviruses, which have an elongated shape, VP4 from Dicistroviridae has a compact structure. Another difference is the lack, on the surface of CrPV, of the deep depression known as the canyon, and the burying of the residues critical for receptor binding in rhinoviruses and enteroviruses. In addition, the cavity known as ‘‘the pocket’’ in the VP1 b-barrel of picornaviruses, and thought to play a critical role in the receptor mediated destabilization that leads to release of the viral RNA in the cytosol, is absent in VP1 from CrPV. Overall, these differences suggest that Dicistroviridae and Picornaviridae use different sites and mechanisms of receptor attachment. In spite of these differences, the capsid proteins of CrPV adopt conformations strikingly similar to those of classical picornaviruses (Tate et al., 1999).
2. Viral replication cycle The viral replication cycle of DCV has been extensively studied by Cherry and Perrimon. In vitro experiments using DL2 cells indicated that viral particles are internalized by clathrin-mediated endocytosis. These findings were confirmed genetically in vivo, as flies with mutations in a-adaptin, clathrin heavy chain, nucleoside diphosphate kinase (encoded by the gene abnormal wing discs) or synaptotagmin exhibited significant resistance to infection with DCV (Cherry and Perrimon, 2004). Using genome-wide RNAi screens, Cherry, Perrimon and collaborators further established that DCV replicates on cellular vesicles derived from the Golgi apparatus, and that translation of viral proteins was highly sensitive to the levels of ribosomes in the cells (Cherry et al., 2005, 2006). Once in the drosophila cells, DCV protein synthesis triggers a remodeling of the
Drosophila Viruses and the Study of Antiviral Host-Defense
239
Golgi apparatus, and apparition of cytosolic vesicles with a mean diameter of 115 nm. The coat protein complex I (COPI), which is required for retrograde transport of proteins and membrane from the Golgi to the endoplasmic reticulum (ER), is required for the formation of these vesicles, and for efficient DCV replication. By contrast, neither COPII (mediating anterograde transport from ER to the Golgi) nor autophagy are required for the DCV replication cycle. Fatty acid biosynthesis is also required for DCV replication in DL2 cells, and probably contributes to the formation of the vesicles supporting DCV replication, which are larger than the 50 nm diameter COPI vesicles (Cherry et al., 2006). The DCV RNA replication complex associates with this new cytoplasmic organelle in infected cells to perform RNA replication. Importantly, both the sensitivity to ribosome levels and the COPI activity coupled with fatty acid biosynthesis also affect replication of poliovirus, a bona fide picornavirus, in mammalian cells, confirming the close relationship between Dicistroviridae and at least some Picornaviridae, and establishing the relevance of the drosophila system to investigate the basic mechanisms of virus-host cell interactions (Cherry et al., 2005, 2006). The RNA-linked VPg protein is thought to prime RNA synthesis, as shown for picornaviruses. Like picornaviruses, DCV inhibits CAP-dependent translation of cellular mRNAs in infected cells, leading to preferential translation of viral mRNAs (Cherry et al., 2005). The mechanism of assembly of the viral particles after replication and translation is still poorly characterized. Unlike CrPV, DCV is not lytic, and can persistently infect drosophila cells without apparent cytopathic effects.
B. Interaction with drosophila The outcome of the infection differs strikingly depending on the infection route. DCV is extremely pathogenic when injected in the body cavity. Injection of a few particles leads to the rapid multiplication and spreading of DCV to multiple organs, including the fat body, trachea, visceral muscles along the midgut, a subset of somatic muscles and, in female flies, the epithelial sheath surrounding the egg chamber (Cherry and Perrimon, 2004; Lautie-Harivel and Thomas-Orillard, 1990; Sabatier et al., 2003). The large number of viral particles assembled in infected cells form typical para-crystalline arrays easily recognized. As a result of this rapid and massive spreading of DCV through the organism, flies rapidly succumb to the infection. By contrast, natural infection (by the oral route or also possibly the respiratory tract) does not lead to major symptoms of infection, and is almost non pathogenic. The virus does however affect the reproductive capacity of drosophila, and infected females produce more eggs and more offspring than control females (Gomariz-Zilber et al., 1995). In addition, contamination at the larval
240
Tu¨nde Huszar and Jean-Luc Imler
stage has an accelerating effect on the development, and adult flies emerge more rapidly than normal. DCV is not transmitted vertically from mother to offspring, and infection exclusively occurs between individuals, either at the larval or adult stages (Jousset and Plus, 1975). Overall, these data point to the existence of a complex network of interactions between DCV and its host, and indicate that injection of the virus bypasses important host-defense mechanisms.
IV. OTHER DROSOPHILA VIRUSES A. Drosophila X virus (DXV) DXV was first identified as a contaminant in a series of experiments with Sigma virus. Two flies out of 72 uninfected controls were found to be sensitive to CO2. These flies were confirmed to be SIGMAV free, but contained instead DXV (Teninges et al., 1979). The virus was later found in many drosophila cell lines, although it has never been found in wild populations of flies. The name DXV reflects the enigmatic origin of this virus.
1. Virion and genome structure
DXV belongs to the Birnaviridae family. These viruses are characterized by a double stranded (ds) RNA genome, and owe their name to their bipartite genome. The virions are non envelopped, icosahedral particles with a diameter of 70 nm and a triangulation T ¼ 13(Coulibaly et al., 2005; Delmas et al., 2004; Teninges et al., 1979) (Table I). The two dsRNA molecules that compose the genome of DXV are called segments A and B. Segment A is 3360 bp long, and contains two ORFs. The first one is very large, and covers most of the segment (3096 nucleotides). The second ORF is 711 nt long, and overlaps with the 30 end of the large ORF, but in a different reading frame (Chung et al., 1996). The large ORF encodes a 128 kDa polyprotein, which is cotranslationally processed to form the major structural proteins of the viral particle (VP2 and VP3) and the proteolytic enzyme (VP4) responsible for processing of the precursor protein. The 49 kDa precursor pVP2 undergoes a slow posttranslational cleavage during particle assembly to generate the 45 kDa VP2 final product. The small peptides produced during this maturation are associated with the viral particles, and in the case of the avian birnavirus Infectious Bursal Disease Virus (IBDV), one of them (pep46) was shown to play an important role in viral entry into cells (Galloux et al., 2007). VP2 is the only component of the viral icosaedral capsid. The 34 kDa VP3 interacts with both the C-terminal end of the precursor pVP2, and with VP1, and controls birnavirus particle morphology. It forms the inner layer of the virion. Finally, the 27 kDa VP4 protein is the protease that processes
Drosophila Viruses and the Study of Antiviral Host-Defense
241
the 128 kDa polyprotein. VP4 is a serine protease, which contains a catalytic serine-lysine dyad in its active site. Other proteins belonging to this group of serine proteases include signal peptidase and the Lon proteases from bacteria (Birghan et al., 2000; Feldman et al., 2006). The small ORF encodes a putative 27 kDa basic protein. Interestingly, the size and location of this ORF is the major difference between the genetic organization of DXV and other birnaviruses. In infectious pancreatic necrosis virus (IPNV) and IBDV, which infect, respectively, fishes and birds, the small ORF overlaps the 50 end of the polyprotein, and its product is 10 kDa smaller than predicted for DXV. The only similarity between the small ORF products of DXV, IPNV and IBDV is the high number of lysine and arginine residues, which suggest that these molecules might interact with RNA (Chung et al., 1996). This ORF has not been functionally characterized in DXV, but is not required for IBDV replication in tissue-culture, suggesting that it could be involved in host-virus interactions in vivo. The 3243-bp B segment encodes VP1, the viral RNA-dependent RNA polymerase, which is found both free and covalently attached to the genomic RNA segments (Shwed et al., 2002). Like polymerases from other dsRNA viruses, VP1 catalyzes both replication and transcription of the viral genome. VP1 contains a GTP-binding site, involved in the self-guanylation of the protein. This results in the formation of a VP1pG complex bound to the 50 end of both genome fragments that acts as a primer during RNA synthesis and remains covalently linked to the 50 end of the RNA. Birnavirus polymerase initiates RNA synthesis via protein priming, a mechanism shared with other viruses, including picornaviruses. In general, the protein primer and the polymerase are separate molecules, but in the case of birnaviruses both functions are carried out by VP1.The VP1 proteins of birnaviruses form a defined subgroup of viral RdRPs. These enzymes contain five conserved motifs (A-E) that define the catalytic domain. The recent resolution of the structure of the VP1 polymerase from IBDV revealed a unique active site topology for the birnavirus RdRP. A first difference with standard RdRPs is the position of motif C, which is found on the N-terminal side of the other catalytic motifs of VP1 (C-A-B-D-E), instead of the classical arrangement (A-B-C-D-E). The second difference is that the classical Gly-Asp-Asp (GDD) sequence found in motif C of most RdRPs is replaced by a Ala-Asp-Asn (ADN) sequence in VP1 from IBDV, IPNV and DXV. This substitution in the tripeptide, which results in an active site containing only two Asp instead of three, leads to suboptimal function of the polymerase (Pan et al., 2007; Shwed et al., 2002). The reduced activity of the VP1 polymerase may represent an adaptation of birnaviruses to their hosts, to modulate their virulence and facilitate their dissemination.
242
Tu¨nde Huszar and Jean-Luc Imler
In summary, although birnaviruses share several characteristics with other dsRNA viruses (dsRNA genome, T ¼ 13 capside structure), they also exhibit several important differences, which suggest an evolutionary link with ss (þ) RNA viruses (VPg-linked genome and protein priming, polyprotein coding strategy).
2. Viral replication cycle Interaction of DXV with host-cell membranes is mediated by VP2, which is the only viral capsid protein in birnaviruses. A likely model for DXV entry into cells, based on IBDV, is that virus binding to the plasma membrane of the cell leads to endocytosis. After internalization, the environment of the endosome may trigger the release of a 43 amino-acid residue peptide produced from the pVP2 precursor, and equivalent to pep46 from IBDV (Galloux et al., 2007). This amphiphilic peptide then induces destabilization and pore-formation in the endosomal membrane, allowing the virion to access the cytosolic environment. Like other dsRNA viruses, DXV keeps its genome hidden from the cellular defense mechanisms, inside the capsid. Indeed, intact birnavirus particles are replication competent, and they are capable of producing viral messengers in a semiconservative manner. The crystal structure of the birnavirus IBDV reveals a pentameric a-helical hydrophilic channel that probably mediates extrusion of the newly transcribed birnavirus mRNAs from the capsid into the cytosol (Coulibaly et al., 2005). Viral mRNAs are then translated and the viral polyproteins assemble around them to form new virions. The newly encapsidated (þ) strands are then replicated by VP1. In drosophila cells at 25 C DXV viral proteins accumulate between 4 and 12 h post-infection, and infection results in cell lysis after 24 h (Nagy and Dobos, 1984; Teninges et al., 1979).
3. Interaction with drosophila
In vivo interactions between DXV and its presumed host, drosophila, are poorly characterized. Adult flies injected with a suspension of DXV die 10–20 days after the injection, depending on the inoculum concentration (Teninges et al., 1979; Zambon et al., 2005). One symptom of infection by DXV is sensitivity to anoxia, which becomes apparent 5–7 days after infection: infected flies are killed by exposure for several minutes to CO2 or nitrogen, whereas a similar treatment of uninfected flies leads to anesthesia, followed by rapid recovery after return to a normal atmosphere. Viral particles in dead flies are found in most organs, including the digestive tract, brain, muscles, ovaries, testis and Malpighian tubules. Earlier in infection, after the appearance of the anoxia sensitivity phenotype, viral particles are found in the trachea cells, gut cells and the muscle sheath surrounding several organs, indicating that these tissues are primary sites of viral amplification in vivo (Teninges et al., 1979; Zambon
Drosophila Viruses and the Study of Antiviral Host-Defense
243
et al., 2005). In addition, these data suggest that natural infection by DXV may occur through the digestive or respiratory tract. Indeed, contact transmission between flies has been documented. By contrast, DXV did not multiply and was not pathogenic in several cell lines or primary cell cultures from vertebrates (Teninges et al., 1979). Altogether, these data support the idea that DXV is an insect virus.
B. Drosophila F virus (DFV) DFV has been identified in laboratory stocks and natural populations of drosophila. It belongs to the Reoviridae family (Table I).
1. Virion and genome structure Reoviruses have dsRNA genomes, like birnaviruses. Birnaviruses and reoviruses are the only dsRNA viruses known to infect vertebrates, and only reoviruses infect mammals (the name reovirus derives from respiratory and enteric orphan virus), causing pathology only in the very young (in particular rotaviruses, the most important cause of severe diarrhea in children worldwide). These non-enveloped viruses have large particles of about 70–80 nm diameter, which contain a dsRNA genome segmented in 10–12 fragments, divided in three classes based on their sizes, large (L1–3), medium (M1–3) and small (S1–4). The L, M and S genome segments encode the viral proteins l, m and s, respectively. The genome is surrounded by two concentric protein shells. The proteins l1–3, m2, and s2 form the inner capsid or core (T ¼ 1), which is surrounded by the icosahedral outer capsid formed by m1, s1, and s3 (T ¼ 13)(Chandran and Nibert, 2003). The virus DFV has been identified in flies collected from laboratory stocks, and from some natural populations of D. melanogaster. It is also present in drosophila tissue-culture cells, in particular the l(2)mbn cell line, where it is abundant (Gateff et al., 1980; Plus et al., 1975a). DFV virions are spherical particles of 60–70 nm diameter, with a capsid composed of two layers comprising eight polypeptides. These polypeptides range in size from 40 to 150 kDa, in agreement with typical reoviral proteins, and are encoded by 10 segments of dsRNA.
2. Viral replication cycle The replication cycle of DFV in drosophila cells has not been studied. Most likely, it resembles the viral cycle of mammalian reoviruses. The replication cycle of Reoviridae involves receptor-mediated endocytosis of viral particles, which is mediated by interaction of the protein s1 with host-cell plasma membrane receptors. Upon delivery of the virions to the lysosomal compartment, the protein s3 is cleaved by acid-dependent proteases, and this cleavage exposes m1. The latter mediates by still unknown mechanisms the penetration of the viral core in the cytosol.
244
Tu¨nde Huszar and Jean-Luc Imler
Once in the cytosol, the viral RNA-dependent RNA polymerase encoded by l3 and contained in the inner capsid produces capped mRNAs, which are released in the cytosol, and translated. The next step in the replication cycle is the assortment and packaging of the viral mRNAs, which involve the structural protein s3 and the non-structural proteins sNS and mNS. The synthesis of the minus-strand of the RNA genome is then initiated, within these nascent viral particles. Completion of the minus strand RNA synthesis leads either to secondary transcription to produce more viral mRNAs and proteins, or to assembly of the inner and outer capsid proteins around the double-stranded RNA fragments to form complete virion particles. The assembly of the viral particles, the encapsidation of the ten genomic RNA molecules, and the exit of the virions from host cells involve still poorly understood mechanisms (Chandran and Nibert, 2003). Clearly, it would be interesting to know more about DFV replication in drosophila cells.
3. Pathogenesis in drosophila
DFV has been identified as a latent virus in D. melanogaster, and does not seem to cause major adverse effects to the flies. Injection of a DFV viral suspension in flies has been reported to cause lethality within one to two weeks, but the presence of contaminating DCV particles in the inoculum cannot be excluded (Gateff, 1994). Another reovirus, Drosophila S virus or DSV, has been isolated from populations of the closely related species D. simulans (Lopez-Ferber et al., 1989; Louis et al., 1988). This reovirus, which is different from DFV, is transmitted vertically, mostly by maternal transmission. DSV is the probable agent of the S character in D. simulans, which is characterized by the absence or abnormalities of bristles mainly at the dorsocentral and scutellar region. The phenotype and its severity correlate with the presence and density of viral particles in subcuticular cells, including chaetal forming cells, thus providing an explanation for the morphogenesis problems associated with the infection. DSV can also be detected in the male and female gonads, allowing vertical transmission, and in the trachea and some muscles. Infection by DSV is associated with a small reduction in fitness. The DSV viral particles appear to be very fragile, which probably explains the high thermosensitivity of the S character and lack of infectivity of DSV upon injection into normal flies (Lopez Ferber et al., 1997).
C. Other RNA viruses 1. Unclassified RNA viruses Other RNA viruses have been described in drosophila (Table I). Most of them share the properties of picorna-like viruses, in particular a singlestranded coding RNA genome and small (27–30 nm diameter)
Drosophila Viruses and the Study of Antiviral Host-Defense
245
non-enveloped capsids, but differ significantly from DCV on the basis of serology, pathogenesis and/or physicochemical properties. These include drosophila P virus (DPV), found in laboratory stocks and wild populations of flies, mostly from tropical areas. The P virus is much less virulent than the C virus (DCV), killing injected flies after more than two weeks (versus as low as 3 days for DCV)(Jousset et al., 1972; Plus et al., 1975b). DPV replicates mainly in the intestine and the malpighian tubules, but even in these tissues the density of viral particles observed is low compared to DCV infected tissues. DPV can also be found in tracheae, periovarian sheath, and follicular cells. The virus, which is serologically related to iota virus isolated from D. immigrans, can be transmitted vertically through the female germ-line (Plus et al., 1975b). DPV has not been characterized molecularly at this stage, nor is its replicative mechanism known. This virus therefore cannot be classified at this stage. Another poorly characterized virus is the recently described Nora virus. This virus causes persistent infection in D. melanogaster and is present in both laboratory stocks and wild populations, at a titer varying between 104 and 1010 genome copies per fly. It does not cause any obvious pathological effect. The viral particles are non-enveloped, with a diameter of about 30 nm, and contain a polyadenylated positive-sense single stranded RNA genome. Unlike other picorna-like viruses, the 11,879 nt RNA genome of Nora virus exhibits four ORFs instead of one or two. Only the largest of these ORFs, ORF2, bears significant sequence similarity with picornavirus-like genes, and includes sequences coding an RNA helicase, a protease and an RNA-dependent RNA polymerase (Habayeb et al., 2006). Finally, another atypical non envelopped RNA virus has been identified on one occasion in a drosophila cell line. The genome of this virus, HPS-1, is composed of a single 6 kb double-stranded RNA molecule, contained in a 36 nm diameter virion. This virus has not been described from wild-caught flies, and its origin is unknown (Scott et al., 1980).
2. Other insect RNA viruses
Two groups of small RNA viruses, the unassiagned genus Iflavirus and the family Nodaviridae, have been isolated from insects, but not from drosophila so far. Iflaviruses closely resemble Picornaviridae. The genome is translated as a single ORF, with the capsid proteins at the 50 end and the non structural proteins at the 30 end. These viruses (e.g., Infectious Flacherie Virus from Lepidoptera or Sacbrood virus from Hymenoptera) differ from picornaviruses by the arrangement of the capsid proteins, which is more similar to that observed in members of the Dicistroviridae (Fig. 2A)(Le Gall et al., 2008). The nodavirus FHV (Flock house virus) has been isolated from the Coleoptera Costelytra zealandica near the Flock House Agricultural research station in New Zealand in 1983 (Scotti et al., 1983). Its genome consists of two positive-strand RNA molecules, which are both packaged
246
Tu¨nde Huszar and Jean-Luc Imler
in a non-enveloped icosaedral virion. RNA1 (3.1 kb) encodes the 112 kDa replicase, whereas RNA2 (1.4 kb) encodes the 43 kDa capsid protein precursor (Schneemann et al., 1998). In infected cells, the subgenomic RNA3 (0.4 kb) is produced from RNA1. It encodes the protein B2, a potent suppressor of RNA interference (Chao et al., 2005; Li et al., 2002). Although isolated from a Coleoteran insect, FHV has a broad host range, and replicates efficiently in drosophila cells in tissue culture and in vivo, providing an interesting model to study host-virus interaction (GalianaArnoux et al., 2006; Wang et al., 2006).
D. Gypsy and infectious retrotransposons 1. Retrotransposable elements and errantiviruses Retrotransposons are transposable elements that replicate by reverse transcription of an RNA intermediate, followed by integration of the resulting DNA into the genome of host-cells. Retrotransposons are widespread in eukaryotes, and belong to different classes (Fig. 3A)(Kaminker A Non LTR retrotransposons (I factor) ORF 1 ORF 2 RT-RH
gag
pol
LTR retrotransposons (copia) 5⬘LTR
3⬘LTR Pro - Int - RT - RH
gag
pol
Retroviruses (gypsy) 5⬘LTR
3⬘LTR Pro - RT - RH - Int
gag
pol
env
B
FIGURE 3 Retroviruses. (A) schematic representation of the genome structure of retrotransposons and the retrovirus gypsy. RT: reverse transcriptase; RH: RNase H; Pro: protease; Int: integrase; LTR: long terminal repeat. (B) virions of the retrovirus ZAM in the cytosol of a follicular cell. The virions are indicated by arrowheads. Scale bar: 100 nm. Picture courtesy of C. Vaury.
Drosophila Viruses and the Study of Antiviral Host-Defense
247
et al., 2002). A first class, the non-LTR retrotransposons, is composed of retroelements that contain two large open reading frames similar to the gag and pol genes from retroviruses, but do not contain long terminal repeats (LTR). These genes encode, respectively, the capsid and nucleocapsid components and the factors required for replication (Protease, reverse transcriptase, RNaseH, integrase). The second class, the LTR retrotransposons, do contain LTRs, like retroviruses. Most of them contain the same two ORFs as non-LTR retrotransposons and are not infectious. Finally, some LTR retrotransposons contain a third ORF, similar to the env gene of retroviruses found in vertebrates, and can form infectious particles. The env gene typically encodes a transmembrane glycoprotein, which mediates binding to a host-cell receptor, and membrane fusion leading to the intracytoplasmic delivery of the viral capsid (Kim et al., 1994; Song et al., 1994). The drosophila LTR retrotransposons containing an env gene are classified in the family Errantiviridae. Whereas vertebrate retroviruses are predominantly transmitted horizontally by cell-to-cell infection, Errantiviridae, also known as endogenous retroviruses, are mainly transmitted vertically from mother to offspring as integrated copies in the host cell genome (Chalvet et al., 1999). The genome of D. melanogaster contains a large number of LTR retrotransposable elements (up to 304, belonging to 49 families), of which only a few contain an env-like gene. The endogenous retroviruses from drosophila include gypsy, the best characterized Errantivirus, 17.6, 297, tom, roo and ZAM/Idefix. In addition to the presence of an env gene, Errantiviruses also present the pol gene domain arrangement typical of retroviruses (protease-reverse transcriptase-RNaseH-integrase), instead of the arrangement found in classical LTR retrotransposons like copia (proteaseintegrase-reverse transcriptase-RNaseH)(Bucheton, 1995; Terzian et al., 2001). However, the nucleocapsid-like region encoded by the gypsy gag gene differs from the gag sequences found in classical simple retroviruses (Gabus et al., 2006).
2. The gypsy endogenous retrovirus
Gypsy has a genetic organization reminiscent of that from classical vertebrate gammaretroviruses. Its 7469 bp genome contains the three canonical ORFs gag, pol and env, flanked by two 482 nucleotides LTRs (Marlor et al., 1986). Most drosophila stocks contain a few (less than 5, mostly 1 or 2) gypsy elements, located on chromosome arms. In addition, a large number of defective copies are present in pericentromeric heterochromatin. Gypsy is normally repressed by the flamenco locus (see below). When this control fails, gypsy can transpose at high rates, resulting in induced mutability. This property provides a test to monitor gypsy activity, taking advantage of the existence of hot-spots for gypsy insertions, for example in the gene ovo. Thus, reversion of the phenotype of female flies carrying the
248
Tu¨nde Huszar and Jean-Luc Imler
dominant ovoD1 mutations (sterility due to developmental problems in the ovaries) can be used to monitor gypsy activity (Bucheton, 1995; Prud’homme et al., 1995). Importantly, gypsy is the only endogenous errantivirus that has been demonstrated to be capable of exogenous infection in drosophila. Three lines of evidence support the claim that gypsy is a retrovirus. First, feeding larvae in which gypsy is inactive with extracts from flies exhibiting high gypsy activity, or with purified viral particles, leads to the insertion of new copies of gypsy in their progeny, indicating that gypsy is infectious (Kim et al., 1994; Song et al., 1994). Second, a Moloney murine leukemia virus (Mo-MLV) pseudotyped by the gypsy envelope can infect drosophila cells, indicating that the env gene of gypsy plays a critical role in the infectivity of the virus (Teysset et al., 1998). Third, ex vivo experiments indicate that gypsy Env has fusogenic activity, demonstrating that Env is functional and can mediate membrane fusion between the viral particle envelope and the plasma membrane of the host-cell (Misseri et al., 2004). Interestingly however, the invasion of the female germline by gypsy was shown to be independent of the env gene. Thus, gypsy appears to rely on a double strategy to maintain itself in the drosophila populations. On one hand, derepression of gypsy in the soma appears sufficient to ensure transfer of gypsy to the germline, by a non-infectious, env-independent route. On the other hand, the env gene probably ensures that a minimal number of active gypsy provirus is maintained in the genome of natural populations (Chalvet et al., 1999). This hypothesis is supported by genomic analysis of gypsy sequences in drosophila species, which point to occasional envmediated horizontal transfer of gypsy (Ludwig et al., 2008; Mejlumian et al., 2002; Terzian et al., 2000). It will be interesting to address the function of the env gene from other drosophila errantiviruses. For example, in the case of the endogenous retrovirus ZAM, the 8.4 kb RNA and the Gag and Env polypeptides are expressed in a small set of follicle cells surrounding the oocyte, and ZAM is thought to be transferred to the germline through the endosome/exosome pathway, concomitantly with yolk transfer (Fig. 3B). The role of env in this process, if any, is not know at this stage, although it has been suggested that ZAM could bud intracellularly. Should this be the case, Env could mediate release of ZAM at the apical side of the follicle cells together with yolk proteins and precursors of the vitellin membrane (Brasset et al., 2006; Leblanc et al., 2000).
3. Origins of the infectious abilities of drosophila retroviruses The question of the origin of retroviruses has long been of interest for virologists. Phylogenetic analysis of reverse transcriptase sequences indicate that retroviruses most likely have derived from LTR retrotransposons, from which they differ by the acquisition of an env gene (Malik et al., 2000; Terzian et al., 2001). The origin of this env gene therefore reflects the
Drosophila Viruses and the Study of Antiviral Host-Defense
249
origin of retroviruses. It is difficult to ascertain the origin of the env gene from vertebrate retroviruses, since this gene is rapidly diverging, as a result of a strong selection pressure to avoid recognition by the host immune system. Tracing the history of env in insect errantiviruses is easier. Interestingly, these studies reveal significant levels of sequence homology, as well as conservation of several motifs and cystein arrays, between the env gene product from gypsy and the F envelope protein from baculoviruses infecting lepidopterans (Malik et al., 2000; Rohrmann and Karplus, 2001). Thus, a likely scenario is that gypsy acquired its env gene from a baculovirus genome. This hypothesis is supported by the identification of LTR retrotransposons inserted in the genome of some baculoviruses. Interestingly, similar analysis for endogenous retroviruses from a nematode and a plant also suggest that their env genes originate from the genome of other viruses (Malik et al., 2000). Transposons are known to undergo horizontal transfer, presumably using vectors such as DNA viruses. The acquisition by LTR retrotransposons of an env gene from these vectors would enable them to bypass the requirement for a vector, thus increasing their probability of transfer across individuals and species. Interestingly, the drosophila genome contains a gene, Iris, encoding a protein related to gypsy Env and baculovirus F proteins (Lung and Blissard, 2005; Malik and Henikoff, 2005). Careful analysis of insect genome sequences suggests that this cellular gene has been acquired (‘‘domesticated’’) from retroviral genomes (roo in particular), and that this happened in at least two independent occasions, in drosophila and mosquitoes. The function of Iris is unknown at this stage. In mammals, retroviral genes have been domesticated to exploit enzymatic functions from transposons (e.g., RAG proteins mediating DNA recombination at antigen receptor loci in lymphocytes) or to counter viral proteins and participate in host-defense (e.g., Fv1 in mice, which encodes a Gag-like protein and is thought to interfere with some retroviral infections by interacting with incoming capsid proteins)(Goff, 2004). An interesting aspect of Iris is that the gene has been subject to strong positive selection. This argues for a role of Iris, by a mechanism that remains to be characterized, in the restriction of env-mediated infection of dipteran insects by retroviruses and baculoviruses (Malik and Henikoff, 2005).
V. ANTIVIRAL REACTIONS IN DROSOPHILA Viral infection is a major burden for all eukaryotic (and prokaryotic) cells. As a result, both plants and animals possess efficient mechanisms to detect and counter viral infections. Although still in its infancy, the study of drosophila antiviral immunity has already provided evidence
250
Tu¨nde Huszar and Jean-Luc Imler
for two types of host-defense mechanisms, an intrinsic cell defense mechanism based on RNA interference, and an inducible response (Beutler et al., 2007).
A. RNA interference RNA interference, or RNAi, was first identified as a potent system of hostdefense in plants. It was recently shown to also play an important role in host defense in invertebrates (Ding and Voinnet, 2007).
1. The RNAi pathways RNAi is initiated by the RNaseIII enzyme Dicer-2, which recognizes double stranded RNA molecules, and processes them (‘‘dicing’’) into 21–25 base pair small interfering (si) RNA duplexes. With the help of the double-stranded RNA binding protein R2D2, one strand of the duplex, the ‘‘guide’’ strand, is then transferred to an effector complex, the RNA-induced silencing complex or RISC, while the second strand (‘‘passenger’’ strand) is degraded. The RISC complex contains a member of the Argonaute family, AGO2, which contains an RNaseH domain and cleaves the RNA molecules targeted by the guide strand (‘‘slicing’’)(Tolia and Joshua-Tor, 2007). This scheme enables host cells to detect dsRNAs, a hallmark of replication of many viruses, and to activate a nuclease that relies on the base-pairing of complementary sequences to specifically degrade viral genomes (Fig. 4A)(Ding and Voinnet, 2007; Marques and Carthew, 2007). The drosophila genome encodes a second Dicer gene, Dicer-1, which mediates the production of a second type of small regulatory RNAs, the micro (mi) RNAs and plays essential developmental functions (Lee et al., 2004). R3D1 and AGO1 are paralogues of R2D2 and AGO2, respectively, and mediate silencing by miRNAs. Three other closely related Argonaute family members are encoded by the drosophila genome. These proteins, Piwi, Aubergine (Aub) and AGO3, define a third silencing pathway, characterized by a distinct class of small regulatory RNAs, the Piwi-associated RNAs or piRNAs (Aravin et al., 2007; O’Donnell and Boeke, 2007). As detailed below, production of piRNAs does not involve enzymes of the Dicer family.
2. Control of RNA virus infections by RNAi in drosophila RNAi plays a critical role in antiviral host-defense in drosophila, and flies deficient for the genes dicer-2, r2d2, or Ago2 exhibit increased sensitivity to infection by the dicistroviridae DCV and CrPV, the birnavirus DXV, and other exogenous RNA viruses such as the nodavirus Flock house virus (FHV) and the alphavirus Sindbis virus (SINV)(Galiana-Arnoux et al., 2006; van Rij et al., 2006; Wang et al., 2006; Zambon et al., 2006).
Drosophila Viruses and the Study of Antiviral Host-Defense
251
A Uncoating Replication ssRNA Slicing Dicing Cleaved viral genome
AGO2
dsRNA
RISC
Dcr2
RISC Guide strand
AGO2
Dcr2
Dcr2 R2D2
Passenger strand
B
piRNAs
Gypsy
Flamenco
X
Gypsy II
Gypsy III
C Piwi/Aub 1 5⬘U A 10
3⬘ ?
10 N N 1
Primary piRNA 3⬘ 5⬘ Gipsy mRNA
Slicer
3⬘end processing
Aub
AGO3
5⬘U
3⬘
Antisense piRNA
3⬘
AGO3 3⬘ 5⬘
A U Slicer
5⬘
A
5⬘
Sense piRNA
piRNA cluster transcript 3⬘ ?
FIGURE 4 RNA interference and the control of viral infection in drosophila. (A) siRNA mediated antiviral defense. The RNase III enzyme Dicer-2 recognizes double stranded RNA in the cytosol of infected cells, and processes them into 21–25 nt siRNA fragments. R2D2 separates the two strands of the siRNA duplex, and the guide strand is incorporated in the RISC complex. The guide strand will target the RISC complex to
252
Tu¨nde Huszar and Jean-Luc Imler
Increased lethality of mutant flies is coupled with increased viral load, in agreement with the proposed role of the RNAi machinery on the degradation of viral RNAs. In addition, siRNAs of viral origin can be detected in virus infected flies, and participate in the protection against the infection. Indeed, transgenic flies expressing a replicating viral RNA from FHV (and thus producing siRNAs) exhibit some protection against a challenge by this virus. As expected for this sequence-specific type of immunity, the protection is virus-specific, and FHV-transgenic flies are not protected against a challenge by DCV (Galiana-Arnoux et al., 2006). RNAi also plays an important role in the control of viral infections in mosquitoes, including species vector of arboviruses, and in the nematode Caenorhabditis elegans (Keene et al., 2004; Lu et al., 2005; Schott et al., 2005; Wilkins et al., 2005). Not surprisingly given its importance, insect viruses have adapted to RNAi, and evolved mechanisms to counter this host-defense. This was first shown for FHV, which encodes a small protein known as B2 that is a potent suppressor of RNAi (Li et al., 2002). B2 is a dsRNA binding protein, which associates with viral double-stranded RNA replication intermediates, and protects them from recognition by Dicer-2 (Chao et al., 2005). Indeed, replication of FHV RNA1 in transgenic flies is completely suppressed in the absence of B2, and this suppression is Dicer-2 dependent (Galiana-Arnoux et al., 2006; Wang et al., 2006). DCV also encodes a suppressor of RNAi, at the N-terminus of ORF1, which contains a bona fide dsRNA binding motif (van Rij et al., 2006). Interestingly, its sequence is completely different from that of the suppressor of RNAi of CrPV, which is also located at the N-terminus of ORF1 (Wang et al., 2006). The mode of action of this suppressor is not known at this stage. Thus it appears that DCV and CrPV, which share 58% of amino acid identity in ORF1, have adopted different strategies to counter RNAi. There is no doubt that other (all?) insect viruses also encode suppressors of RNAi. Their identification and the detailed characterization of their mode of action will provide useful information on the mechanisms of RNA interference in the control of viral infections in insects. single stranded RNA molecules of complementary sequence, which will be cleaved by the RNaseH enzyme AGO2. (B) The locus flamenco produces piRNAs, which silence gypsy expression on drosophila chromosomes (only chromosomes I to III are shown). Redrawn with modifications from (O’Donnell and Boeke, 2007). (C) The ping-pong model for synthesis of piRNAs. Primary piRNAs associate with the Argonaute family members Piwi or Aub, and guide them to complementary RNA molecules. Piwi/Aub cleaves the gypsy mRNA (slicer) to generate the 50 end of a new sense piRNA. An unknown nuclease processes the 30 end of the piRNA. The newly formed sense piRNA associates with AGO3, and guides it to RNAs transcribed from the piRNA cluster, to generate antisense piRNAs. Redrawn with modifications from (Aravin et al., 2007).
Drosophila Viruses and the Study of Antiviral Host-Defense
253
By contrast, vertebrates do not appear to rely on RNAi to control viral infections. Vertebrates contain a single Dicer gene, involved in the production of miRNAs, and carrying essential developmental functions, like Dicer-1 in drosophila. miRNAs in mammals are known to participate in the regulation of the immune response, through their roles on development and differentiation of leukocyte subsets. Some miRNAs targeting viral sequences, or produced by viruses in infected cells have also been described (Muller and Imler, 2007). However, mice carrying an hypomorphic mutation in the dicer gene (null mutants are embryonic lethal) are not dramatically impaired in their resistance to virus infection (Otsuka et al., 2007). These data are supported by the fact that siRNAs of viral origin could not be detected in infected mammalian cells (Pfeffer et al., 2005). Thus, the important antiviral role of RNAi appears to be restricted to plants and invertebrates.
3. Control of endogenous retroviruses by piRNAs
The spread of gypsy, ZAM and idefix is controlled by the flamenco locus (Prud’homme et al., 1995). This locus maps to the pericentromeric heterochromatin on the X chromosome, and consists of a large number of truncated or defective retrotransposons (Desset et al., 2003; Mevel-Ninio et al., 2007). Flamenco is a major piRNA cluster, and is thought to produce a mixture of sense and antisense piRNAs (Pelisson et al., 2007) (Fig. 4B). Siomi, Hannon and collaborators sequenced the small RNAs associated with Piwi, Aub, and AGO3, and observed that Piwi and Aub bind antisense-strand piRNAs, while AGO3 binds sense-strand piRNAs. In addition, Piwi- and Aub-interacting piRNAs have a U bias for their 50 end nucleotide, whereas AGO3-interacting piRNAs have an A bias at the tenth nucleotide from their 50 end. Finally, the first ten nucleotides of piRNAs associated with AGO3 can be complementary to the first ten nucleotides of piRNAs interacting with Aub. These observations led to the ping-pong model whereby sense piRNAs associated with AGO3 recognize and cleave long antisense RNAs, produced for example at the flamenco locus, thus generating the 50 end of new antisense piRNAs (Brennecke et al., 2007; Gunawardane et al., 2007). The 30 end of the piRNAs might be generated by the nucleases Squash and/or Zucchini, which are both required for transposon repression and piRNA synthesis in drosophila (Pane et al., 2007). The antisense piRNAs then associate with Aub or Piwi, and these complexes cleave sense RNAs, such as those produced from retrotransposons. Again, the cleaved RNAs can be used to generate the sense piRNAs that will be loaded on AGO3, thus amplifying the production of piRNAs (Fig. 4C). This amplification loop using two slicer enzymes associated with piRNAs of different strand complementarity provides an efficient mechanism to generate piRNAs independently from Dicer enzymes. The piRNAs generated in clusters such as
254
Tu¨nde Huszar and Jean-Luc Imler
the flamenco locus can then silence transcription of retrotransposons anywhere in the genome (Fig. 4B). This model is supported by the observation that, in flamenco mutant flies, piRNAs originating from the cluster are lost, and, most importantly, gypsy is de-repressed. A fascinating aspect of this host-defense mechanism against endogenous retroviruses, which appears to be conserved in vertebrates, is that it involves both exquisite specificity (driven by nucleotide strand complementarity), and memory of previous exposure to a given retrotransposon once fragments of its sequences have found their way in piRNA clusters (Aravin et al., 2007; O’Donnell and Boeke, 2007).
B. Inducible response to infection As mentioned above, RNAi is not a major weapon in the antiviral arsenal of mammalian cells. Rather, the hallmark of antiviral immunity in vertebrates is the induction of cytokines of the interferon family, which is triggered by the detection of viral RNAs in the cytosol of infected cells. Interferons then trigger induction of several hundreds of genes in neighboring cells, which mediate an antiviral state (e.g., protein kinase R, 20 -50 oligo (A) synthase, MxA)(Galiana-Arnoux and Imler, 2006). Interestingly, viral infection also triggers an inducible response in flies, although it is still poorly characterized at this stage.
1. JAK/STAT-dependent immune responses Infection of drosophila with bacteria or fungi triggers a strong humoral response, and secretion by the fat body in the hemolymph of a large number of molecules, including potent antimicrobial peptides. Infection of drosophila with DCV does not trigger a humoral response, which may reflect the fact that this type of response would not be efficient against intracellular pathogens like viruses (Sabatier et al., 2003). Microarray analysis nevertheless revealed that about 150 genes are induced by a factor of at least two in DCV infected flies (Dostert et al., 2005). This finding raises two important questions, relating to the signaling pathway(s) mediating this inducible response on one hand, and to the function of the induced genes on the other hand. The list of the genes induced by DCV did not provide any signature pointing to a particular signaling pathway involved in the induction of these genes. In particular, typical target genes for the Toll pathway (e.g., drosomycin), the Imd pathway (e.g., diptericin), the JNK pathway (e.g., flightin) or the JAK-STAT pathway (e.g., Turandot (Tot) genes) are not induced by DCV infection, at least not in the first 48 h. A detailed analysis of the promoter of the gene vir-1, which is strongly induced by DCV infection, mapped the virus-response element to two 15 bp motifs corresponding to consensus binding sites for the transcription
Drosophila Viruses and the Study of Antiviral Host-Defense
255
factor D-STAT. Such motifs were also found in the proximal promoters of many genes induced by DCV. Genetic analysis confirmed that induction of vir-1, as well as several other genes, by DCV depended on the gene hopscotch, which encodes the only JAK kinase found in drosophila. Importantly, hopscotch mutant flies contain an increased viral load, suggesting that some of the genes induced by DCV encode factors that oppose viral replication (Dostert et al., 2005). The identification of these genes and the characterization of the function of the proteins they encode represent a major goal for future studies. Interestingly, hopscotch mutant flies do not succumb more rapidly to a high dose DCV challenge than wild-type controls, even though they contain more virus. This suggests that the inducible response contributes to the development of the pathology, a situation reminiscent of the toxic effects of inflammation in mammals. Recent results indeed revealed that KATP channels, which are expressed in the coronary arteries in mice, where they play an important role in the homeostasis of the immune response, are also required in the heart (dorsal vessel) of drosophila for the resistance to at least some viruses (Croker et al., 2007). The data available at this stage suggest that DCV infection triggers expression of a cytokine of the Unpaired family in infected cells. This cytokine then activates the receptor Domeless, an homologue of the gp130 subunit of the interleukine-6 receptor in mammals, and the JAK/STAT pathway (Agaisse and Perrimon, 2004). This leads to expression of vir-1 and other genes in non-infected cells, possibly triggering an antiviral state. The identification of a function of the JAK/STAT pathway in the control of DCV infection provides an interesting parallel with innate antiviral immunity in mammals, since the JAK/STAT pathway plays important roles in cytokine signaling in vertebrates, and was initially characterized for its essential role in interferon signaling. At this stage, one must however bear in mind that the role of the JAK-STAT pathway in drosophila antiviral immunity was so far only established for DCV, and the generality of this finding remains to be established for other viruses. In addition, the fact that DCV does not induce expression of known JAKSTAT target genes in adult flies such as Tot genes, and that vir-1 is not expressed in flies carrying the gain of function allele hopTum-l (unlike Tot genes), clearly indicate that the JAK/STAT pathway is required, but not sufficient for the antiviral response in flies (Dostert et al., 2005).
2. Evidence for JAK/STAT-independent immune responses The other pathways activated during viral infection are not known at this stage. The gene Vago, which does not contain consensus STAT binding sites in its proximal promoter and remains fully inducible in hopscotch mutant flies, provides a good tool to identify a second pathway activated in virus infected flies (Croker et al., 2007). Wu and colleagues also
256
Tu¨nde Huszar and Jean-Luc Imler
proposed that the Toll pathway might be involved in an antiviral response. Indeed, infection by DXV leads to a strong induction of drosomycin expression, a marker of the Toll pathway (Zambon et al., 2005). This induction appears to be specific to DXV infection, since we did not detect significant induction of antimicrobial peptide expression in flies infected with DCV, FHV or SINV (Dostert et al., 2005; Sabatier et al., 2003). In addition, flies with a loss of function mutation in the gene encoding the NF-kB transcription factor DIF or with a gain of function mutation in the receptor Toll, succumb more rapidly to DXV infection than control flies. Curiously however, flies with loss of function mutations in Toll, spaetzle, tube or pelle do not exhibit a DXV-sensitivity phenotype (Zambon et al., 2005). Thus, DIF might be regulated by a pathway different from the classical Toll pathway in DXV infected flies. The hypothesis that NF-kB pathways participate in antiviral defenses in insects is supported by the presence in the genomes of some DNA viruses (e.g., polydnaviruses) of genes encoding IkB-like proteins that inhibit DIF and Relish in drosophila cells (Thoetkiattikul et al., 2005). In addition, the gene ref(2)P, which mediates refractoriness to SIGMAV infection (see above), has been proposed to encode a component of the Toll pathway and to regulate the activity of DIF and Dorsal (Avila et al., 2002). This hypothesis however has not been confirmed in vivo yet using ref(2)P mutant flies. In conclusion, much remains to be done to characterize in detail the antiviral inducible response, depending or not on the JAK/STAT pathway.
VI. CONCLUSION AND PERSPECTIVES At the end of this review of the literature on drosophila viruses, two major facts emerge. The first is that D. melanogaster is a host to several viruses belonging to different families (Rhabdoviridae, Dicistroviridae, Birnaviridae, Reoviridae, Errantiviridae). These viruses provide interesting experimental systems to analyze how a model organism like drosophila recognizes and controls viruses with different types of RNA genomes (single-stranded of negative or positive polarity, double-stranded; depending on IRES or protein-primer for translation; polyadenylated or not). This diversity contrasts with the recently reviewed situation for honeybees, where all viruses reported so far belong to the Picornavirales order (Dicistroviridae family or Iflavirus genus)(Chen and Siede, 2007). The reason for this difference is not clear, but may reflect the high pathogenicity of these viruses, which led to their identification in bees. Indeed, DCV is so far the most virulent drosophila virus in the intrathoracic injection model of infection. This artificial route of infection may be achieved by Varroa mites in honeybees (Shen et al., 2005; Yue and Genersch, 2005). On a
Drosophila Viruses and the Study of Antiviral Host-Defense
257
different note, the number of RNA viruses reported to infect drosophila strikingly contrasts with the absence of DNA viruses. Several families of DNA viruses (e.g., Iridoviridae, Baculoviridae, Poxviridae, Parvoviridae) have been reported in insects, including for some of them Dipteran insects, but not in drosophila (Friesen and Miller, 2001). The sensing of DNA viruses by mammalian cells clearly differs from the sensing of RNA viruses, and is at present an area of intense investigation (Ishii et al., 2008). Thus, it would be interesting to establish a DNA virus infection model in drosophila. The second fact that emerges is that many of the drosophila viruses remain poorly characterized. Much basic virology remains to be done in order to fully exploit these viruses as tools to understand host-virus interactions. For example, if a great deal is now known on the structure and genome of DXV, little is known about its replication cycle and interaction with drosophila. Similarly, it is surprising that SIGMAV, for which interaction with drosophila has been extensively characterized, remains so poorly characterized molecularly, and regarding its replication cycle in drosophila cells. Finally, DPV, DFV and Nora virus represent promising novel tools that deserve to be better characterized. D. melanogaster is a unique and powerful model to use complementary approaches such as genetics, genome-wide screens, transcriptomics, proteomics, systems biology to analyze in vivo host-virus interactions. The data already at hand point to the importance of using a complementary set of viruses to fully appreciate the spectrum of host-cell responses to viral infection. For example, FHV induces the JAK/STAT target gene Tot, whereas DCV does not. Similarly, DXV infection was reported to induce antimicrobial peptide gene expression whereas DCV, FHV and Sindbis virus (SINV) do not. Finally, the gene dSUR, which encodes the regulatory component of a KATP channel, is required for resistance to FHV infection, but not to DCV infection. These observations illustrate the interest of comparing different viruses and argue for the necessity to gain more information on drosophila viruses, which unlike the exogenous viruses used in some studies (e.g., FHV, CrPV, SINV) have co-evolved with their host. The identification of two completely different suppressors of RNAi in the strongly related DCV and CrPV viruses further illustrates the interest of looking for new viruses, even if they belong to similar families. In summary, in drosophila as in other organisms, there is much to learn from viruses. One foreseeable bottleneck in the nascent field of antiviral innate immunity in drosophila in the coming years will be the insufficient level of characterization of drosophila viruses. These considerations argue for the necessity to invest more on the identification of new drosophila viruses, and on the in-depth analysis of those already at hand.
258
Tu¨nde Huszar and Jean-Luc Imler
ACKNOWLEDGMENTS We thank Cordula Kemp and Stefanie Muller for critical reading of the manuscript. We also express our gratitude to Didier Contamine and Chantal Vaury for providing pictures for Figs. 1 and 3. Our work on RNAi and the control of viral infections in drosophila is supported by a grant from the Agence Nationale de la Recherche (ANR). T.H. is supported by a fellowship from the Ministe`re de l’Education Nationale, de la Recherche et de la Technologie (MENRT).
REFERENCES Agaisse, H., and Perrimon, N. (2004). The roles of JAK/STAT signaling in Drosophila immune responses. Immunol. Rev. 198:72–82. Aravin, A. A., Hannon, G. J., and Brennecke, J. (2007). The Piwi-piRNA pathway provides an adaptive defense in the transposon arms race. Science 318(5851):761–764. Avila, A., Silverman, N., Diaz-Meco, M. T., and Moscat, J. (2002). The Drosophila atypical protein kinase C-ref(2)p complex constitutes a conserved module for signaling in the toll pathway. Mol. Cell Biol. 22(24):8787–8795. Banerjee, A. K. (2008). Response to ‘‘Non-segmented negative-strand RNA virus RNA synthesis in vivo’’ Virology 371(2):231–233. Bangham, J., Obbard, D. J., Kim, K. W., Haddrill, P. R., and Jiggins, F. M. (2007). The age and evolution of an antiviral resistance mutation in Drosophila melanogaster. Proc. Biol. Sci. 274 (1621):2027–2034. Berkaloff, A., Bregliano, J. C., and Ohanessian, A. (1965). Demonstration of virions in Drosophila infected by the hereditary sigma virus. C. R. Acad. Sci. Hebd. Seances Acad. Sci. D 260(22):5956–5959. Beutler, B., Eidenschenk, C., Crozat, K., Imler, J. L., Takeuchi, O., Hoffmann, J. A., and Akira, S. (2007). Genetic analysis of resistance to viral infection. Nat. Rev. Immunol. 7(10):753–766. Birghan, C., Mundt, E., and Gorbalenya, A. E. (2000). A non-canonical lon proteinase lacking the ATPase domain employs the ser-Lys catalytic dyad to exercise broad control over the life cycle of a double-stranded RNA virus. Embo J. 19(1):114–123. Blandin, S., and Levashina, E. A. (2004). Thioester-containing proteins and insect immunity. Mol. Immunol. 40(12):903–908. Bras, F., Teninges, D., and Dezelee, S. (1994). Sequences of the N and M genes of the sigma virus of Drosophila and evolutionary comparison. Virology 200(1):189–199. Brasset, E., Taddei, A. R., Arnaud, F., Faye, B., Fausto, A. M., Mazzini, M., Giorgi, F., and Vaury, C. (2006). Viral particles of the endogenous retrovirus ZAM from Drosophila melanogaster use a pre-existing endosome/exosome pathway for transfer to the oocyte. Retrovirology 3:25. Brennecke, J., Aravin, A. A., Stark, A., Dus, M., Kellis, M., Sachidanandam, R., and Hannon, G. J. (2007). Discrete small RNA-generating loci as master regulators of transposon activity in Drosophila. Cell 128(6):1089–1103. Brzozka, K., Finke, S., and Conzelmann, K. K. (2006). Inhibition of interferon signaling by rabies virus phosphoprotein P: Activation-dependent binding of STAT1 and STAT2. J. Virol. 80(6):2675–2683. Bucheton, A. (1995). The relationship between the flamenco gene and gypsy in Drosophila: How to tame a retrovirus. Trends Genet. 11(9):349–353. Carpenter, J. A., Obbard, D. J., Maside, X., and Jiggins, F. M. (2007). The recent spread of a vertically transmitted virus through populations of Drosophila melanogaster. Mol. Ecol. 16 (18):3947–3954.
Drosophila Viruses and the Study of Antiviral Host-Defense
259
Carre-Mlouka, A., Gaumer, S., Gay, P., Petitjean, A. M., Coulondre, C., Dru, P., Bras, F., Dezelee, S., and Contamine, D. (2007). Control of sigma virus multiplication by the ref(2)P gene of Drosophila melanogaster: An in vivo study of the PB1 domain of Ref(2)P. Genetics 176(1):409–419. Chalvet, F., Teysset, L., Terzian, C., Prud’homme, N., Santamaria, P., Bucheton, A., and Pelisson, A. (1999). Proviral amplification of the Gypsy endogenous retrovirus of Drosophila melanogaster involves env-independent invasion of the female germline. Embo J. 18 (9):2659–2669. Chandran, K., and Nibert, M. L. (2003). Animal cell invasion by a large nonenveloped virus: Reovirus delivers the goods. Trends Microbiol. 11(8):374–382. Chao, J. A., Lee, J. H., Chapados, B. R., Debler, E. W., Schneemann, A., and Williamson, J. R. (2005). Dual modes of RNA-silencing suppression by Flock House virus protein B2. Nat. Struct. Mol. Biol. 12:952–957. Chen, Y. P., and Siede, R. (2007). Honey bee viruses. Adv. Virus Res. 70:33–80. Cherry, S., and Perrimon, N. (2004). Entry is a rate-limiting step for viral infection in a Drosophila melanogaster model of pathogenesis. Nat. Immunol. 5(1):81–87. Cherry, S., Doukas, T., Armknecht, S., Whelan, S., Wang, H., Sarnow, P., and Perrimon, N. (2005). Genome-wide RNAi screen reveals a specific sensitivity of IRES-containing RNA viruses to host translation inhibition. Genes Dev. 19(4):445–452. Cherry, S., Kunte, A., Wang, H., Coyne, C., Rawson, R. B., and Perrimon, N. (2006). COPI activity coupled with fatty acid biosynthesis is required for viral replication. PLoS Pathog. 2(10):e102. Christian, P., Castens, E., Domier, L., Johnson, K., Nakashima, N., Scotti, P., and van der Wilk, F. (2000). In ‘‘Virus taxonomy.’’ (M. Regenmortel, C. Fauquet, D. Bishop, E. Carstens, M. Estes, S. Lemon, J. Maniloff, M. Mayo, D. McGeoch, C. Pringle, and R. Wickner, Eds.). Academic Press, New York, NY. Chung, H. K., Kordyban, S., Cameron, L., and Dobos, P. (1996). Sequence analysis of the bicistronic Drosophila X virus genome segment A and its encoded polypeptides. Virology 225(2):359–368. Contamine, D. (1981). Role of the drosophila genome in Sigma virus multiplication. I. Role of ref(2)P: Selection of host-adapted mutants at the non-permissive allele ref(2)PP. Virology 114:474–488. Contamine, D., Petitjean, A. M., and Ashburner, M. (1989). Genetic resistance to viral infection: The molecular cloning of a Drosophila gene that restricts infection by the rhabdovirus sigma. Genetics 123(3):525–533. Coulibaly, F., Chevalier, C., Gutsche, I., Pous, J., Navaza, J., Bressanelli, S., Delmas, B., and Rey, F. A. (2005). The birnavirus crystal structure reveals structural relationships among icosahedral viruses. Cell 120(6):761–772. Croker, B., Crozat, K., Berger, M., Xia, Y., Sovath, S., Schaffer, L., Eleftherianos, I., Imler, J. L., and Beutler, B. (2007). ATP-sensitive potassium channels mediate survival during infection in mammals and insects. Nat. Genet. 39(12):1453–1460. Curran, J., and Kolakofsky, D. (2008). Nonsegmented negative-strand RNA virus RNA synthesis in vivo. Virology 371(2):227–230. Delmas, B., Kibenge, F., Leong, J., Mundt, E., Vakharia, V., and Wu, J. (2004). Birnaviridae. In ‘‘Virus Taxonomy’’ (C. Fauquet, M. Mayo, J. Maniloff, U. Desselberger, and A. Ball, Eds.), pp. 561–569. Acedemic Press, London. Desset, S., Meignin, C., Dastugue, B., and Vaury, C. (2003). COM, a heterochromatic locus governing the control of independent endogenous retroviruses from Drosophila melanogaster. Genetics 164(2):501–509. Dezelee, S., Bras, F., Contamine, D., Lopez-Ferber, M., Segretain, D., and Teninges, D. (1989). Molecular analysis of ref(2)P, a Drosophila gene implicated in sigma rhabdovirus multiplication and necessary for male fertility. Embo J. 8(11):3437–3446.
260
Tu¨nde Huszar and Jean-Luc Imler
Ding, S. W., and Voinnet, O. (2007). Antiviral immunity directed by small RNAs. Cell 130 (3):413–426. Dostert, C., Jouanguy, E., Irving, P., Troxler, L., Galiana-Arnoux, D., Hetru, C., Hoffmann, J. A., and Imler, J. L. (2005). The Jak-STAT signaling pathway is required but not sufficient for the antiviral response of drosophila. Nat. Immunol. 6(9):946–953. Dru, P., Bras, F., Dezelee, S., Gay, P., Petitjean, A. M., Pierre-Deneubourg, A., Teninges, D., and Contamine, D. (1993). Unusual variability of the Drosophila melanogaster ref(2)P protein which controls the multiplication of sigma rhabdovirus. Genetics 133(4):943–954. Echalier, G. (1997). ‘‘Drosophila cells in culture.’’ Academic Press, New York. . Feldman, A. R., Lee, J., Delmas, B., and Paetzel, M. (2006). Crystal structure of a novel viral protease with a serine/lysine catalytic dyad mechanism. J. Mol. Biol. 358(5):1378–1389. Ferrandon, D., Imler, J. L., Hetru, C., and Hoffmann, J. A. (2007). The Drosophila systemic immune response: Sensing and signalling during bacterial and fungal infections. Nat. Rev. Immunol. 7(11):862–874. Fleuriet, A. (1981a). Comparison of various physiological traits in flies (Drosophila melanogaster) of wild origin, infected or uninfected by the hereditary Rhabdovirus sigma. Arch. Virol. 69(3–4):261–272. Fleuriet, A. (1981b). Effect of overwintering on the frequency of flies infected by the rhabdovirus sigma in experimental populations of Drosophila melanogaster. Arch. Virol. 69(3–4): 253–260. Fleuriet, A. (1999). Evolution of the proportions of two sigma viral types in experimental populations of Drosophila melanogaster in the absence of the allele that is restrictive of viral multiplication. Genetics 153(4):1799–1808. Fleuriet, A. (2001). Evolution of the proportions of two sigma viral types in experimental populations of Drosophila melanogaster. Genetics 157(1):455–456. Fleuriet, A., and Periquet, G. (1993). Evolution of the Drosophila melanogaster-sigma virus system in natural populations from Languedoc (southern France). Arch. Virol. 129 (1–4):131–143. Friesen, P. D., and Miller, L. K. (2001). Insect viruses. In ‘‘Fields Virology’’ (D. Knipe and P. Howley, Eds.), Vol. 1, pp. 599–628. 2 vols. Lippincott Williams & Wilkins, Philadelphia. Gabus, C., Ivanyi-Nagy, R., Depollier, J., Bucheton, A., Pelisson, A., and Darlix, J. L. (2006). Characterization of a nucleocapsid-like region and of two distinct primer tRNALys, 2 binding sites in the endogenous retrovirus Gypsy. Nucleic Acids Res. 34(20):5764–5777. Galiana-Arnoux, D., and Imler, J. (2006). Toll-like receptors and antiviral innate immunity. Tissue Antigens 67:267–276. Galiana-Arnoux, D., Dostert, C., Schneemann, A., Hoffmann, J. A., and Imler, J. L. (2006). Essential function in vivo for Dicer-2 in host defense against RNA viruses in drosophila. Nat. Immunol. 7(6):590–597. Galloux, M., Libersou, S., Morellet, N., Bouaziz, S., Da Costa, B., Ouldali, M., Lepault, J., and Delmas, B. (2007). Infectious bursal disease virus, a non-enveloped virus, possesses a capsid-associated peptide that deforms and perforates biological membranes. J. Biol. Chem. 282(28):20774–20784. Gateff, E. (1994). Tumor-suppressor genes, hematopoietic malignancies and other hematopoietic disorders of Drosophila melanogaster. In ‘‘Primordial Immunity: Foundation for the Vertebrate Immune System.’’ (G. Beck, E. Cooper, and G. Habicht, Eds), Vol. 712, pp. 260–279. Annals of the New York Academy of Sciences. Gateff, E., Gissmann, L., Shresta, R., Plus, N., Pfister, H., Schro¨der, J., and Zur Hausen, H. (1980). Characterization of two tumorous blood cell lines of Drosophila melanogaster and the viruses they contain. In ‘‘Invertebrate systems in vitro’’ (E. Kurstak, K. Maramorosch, and A. Du¨bendorfer, Eds.), pp. 517–533. Elsevier/North Holland Biomedical Press, Amsterdam. Gay, P. (1978). Drosophila genes which intervene in multiplication of sigma virus (author’s transl. Mol. Gen. Genet. 159(3):269–283.
Drosophila Viruses and the Study of Antiviral Host-Defense
261
Geisbert, T. W., and Jahrling, P. B. (2004). Exotic emerging viral diseases: Progress and challenges. Nat. Med. 10(Suppl. 12):S110–S121. Goff, S. P. (2004). Retrovirus restriction factors. Mol. Cell 16(6):849–859. Gomariz-Zilber, E., Poras, M., and Thomas-Orillard, M. (1995). Drosophila C virus: Experimental study of infectious yields and underlying pathology in Drosophila melanogaster laboratory populations. J. Invertebr. Pathol. 65(3):243–247. Gunawardane, L. S., Saito, K., Nishida, K. M., Miyoshi, K., Kawamura, Y., Nagami, T., Siomi, H., and Siomi, M. C. (2007). A slicer-mediated mechanism for repeat-associated siRNA 50 end formation in Drosophila. Science 315(5818):1587–1590. Habayeb, M. S., Ekengren, S. K., and Hultmark, D. (2006). Nora virus, a persistent virus in Drosophila, defines a new picorna-like virus family. J. Gen. Virol. 87(Pt. 10):3045–3051. Hogenhout, S. A., Redinbaugh, M. G., and Ammar el, D. (2003). Plant and animal rhabdovirus host range: A bug’s view. Trends Microbiol. 11(6):264–271. Hwang, L. N., Englund, N., Das, T., Banerjee, A. K., and Pattnaik, A. K. (1999). Optimal replication activity of vesicular stomatitis virus RNA polymerase requires phosphorylation of a residue(s) at carboxy-terminal domain II of its accessory subunit, phosphoprotein P. J. Virol. 73(7):5613–5620. Ishii, K. J., Kawagoe, T., Koyama, S., Matsui, K., Kumar, H., Kawai, T., Uematsu, S., Takeuchi, O., Takeshita, F., Coban, C., and Akira, S. (2008). TANK-binding kinase-1 delineates innate and adaptive immune responses to DNA vaccines. Nature 451 (7179):725–729. Johnson, K. N., and Christian, P. D. (1998). The novel genome organization of the insect picorna-like virus Drosophila C virus suggests this virus belongs to a previously undescribed virus family. J. Gen. Virol. 79(Pt. 1):191–203. Johnson, K. N., and Christian, P. D. (1999). Molecular characterization of Drosophila C virus isolates. J. Invertebr. Pathol. 73(3):248–254. Jousset, F. X., and Plus, N. (1975). [Study of the vertical transmission and horizontal transmission of ‘‘Drosophila melanogaster’’ and ‘‘Drosophila immigrans’’ picornavirus (author’s transl)] Ann. Microbiol. (Paris) 126(2):231–249. Jousset, F. X., Plus, N., Croizier, G., and Thomas, M. (1972). Existence chez Drosophila de deux groupes de Picornavirus de proprie´te´s se´rologiques et biologiques diffe´rentes. C. R. Acad. Sc. Paris 275:3043–3046. Jousset, F. X., Bergoin, M., and Revet, B. (1977). Characterization of the Drosophila C virus. J. Gen. Virol. 34(2):269–283. Kaminker, J. S., Bergman, C. M., Kronmiller, B., Carlson, J., Svirskas, R., Patel, S., Frise, E., Wheeler, D. A., Lewis, S. E., Rubin, G. M., Ashburner, M., and Celniker, S. E. (2002). The transposable elements of the Drosophila melanogaster euchromatin: A genomics perspective. Genome Biol. 3(12):RESEARCH0084. Keene, K. M., Foy, B. D., Sanchez-Vargas, I., Beaty, B. J., Blair, C. D., and Olson, K. E. (2004). RNA interference acts as a natural antiviral response to O’nyong-nyong virus (Alphavirus; Togaviridae) infection of Anopheles gambiae. Proc. Natl. Acad. Sci. USA 101 (49):17240–17245. Kim, A., Terzian, C., Santamaria, P., Pelisson, A., Purd’homme, N., and Bucheton, A. (1994). Retroviruses in invertebrates: The gypsy retrotransposon is apparently an infectious retrovirus of Drosophila melanogaster. Proc. Natl. Acad. Sci. USA 91(4):1285–1289. King, L., and Moore, N. (1988). Evidence for the presence of a genome-linked protein in two insect picornaviruses, cricket paralysis virus and Drosophila C virus. FEMS Microbiol. Lett. 50:41–44. Komarova, A. V., Real, E., Borman, A. M., Brocard, M., England, P., Tordo, N., Hershey, J. W., Kean, K. M., and Jacob, Y. (2007). Rabies virus matrix protein interplay with eIF3, new insights into rabies virus pathogenesis. Nucleic Acids Res. 35(5):1522–1532.
262
Tu¨nde Huszar and Jean-Luc Imler
Landes-Devauchelle, C., Bras, F., Dezelee, S., and Teninges, D. (1995). Gene 2 of the sigma rhabdovirus genome encodes the P protein, and gene 3 encodes a protein related to the reverse transcriptase of retroelements. Virology 213(2):300–312. Lautie-Harivel, N., and Thomas-Orillard, M. (1990). Location of Drosophila C virus target organs in Drosophila host population by an immunofluorescence technique. Biol. Cell 69 (1):35–39. Leblanc, P., Desset, S., Giorgi, F., Taddei, A. R., Fausto, A. M., Mazzini, M., Dastugue, B., and Vaury, C. (2000). Life cycle of an endogenous retrovirus, ZAM, in Drosophila melanogaster. J. Virol. 74(22):10658–10669. Lee, Y. S., Nakahara, K., Pham, J. W., Kim, K., He, Z., Sontheimer, E. J., and Carthew, R. W. (2004). Distinct roles for Drosophila Dicer-1 and Dicer-2 in the siRNA/miRNA silencing pathways. Cell 117(1):69–81. Le Gall, O., Christian, P., Fauquet, C. M., King, A. M., Knowles, N. J., Nakashima, N., Stanway, G., and Gorbalenya, A. E. (2008). Picornavirales, a proposed order of positivesense single-stranded RNA viruses with a pseudo-T ¼ 3 virion architecture. Arch. Virol. 153:715–727. Lemaitre, B., and Hoffmann, J. (2007). The host defense of Drosophila melanogaster. Annu. Rev. Immunol. 25:697–743. L’heritier, P., and Teissier, G. (1937). Une anomalie physiologique he´re´ditaire chez la drosophile. Comptes Rendus de l’Acade´mie des Sci. 205:1099–1101. Li, H., Li, W. X., and Ding, S. W. (2002). Induction and suppression of RNA silencing by an animal virus. Science 296(5571):1319–1321. Lopez-Ferber, M., Veyrunes, J. C., and Croizier, L. (1989). Drosophila S virus is a member of the Reoviridae family. J. Virol. 63(2):1007–1009. Lopez Ferber, M., Ferreiro Rios, A., Kuhl, G., Comendador, M. A., and Louis, C. (1997). Infection of the gonads of the SimES strain of Drosophila simulans by the hereditary reovirus DSV. J. Invertebr. Pathol. 70(2):143–149. Louis, C., Lopez-Ferber, M., Comendador, M., Plus, N., Kuhl, G., and Baker, S. (1988). Drosophila S virus, a hereditary reolike virus, probable agent of the morphological S character in Drosophila simulans. J. Virol. 62(4):1266–1270. Lu, R., Maduro, M., Li, F., Li, H. W., Broitman-Maduro, G., Li, W. X., and Ding, S. W. (2005). Animal virus replication and RNAi-mediated antiviral silencing in Caenorhabditis elegans. Nature 436(7053):1040–1043. Ludwig, A., Valente, V. L., and Loreto, L. S. (2008). Multiple invasions of Errantivirus in the genus Drosophila. Insect Mol. Biol. 17:113–124. Lung, O., and Blissard, G. W. (2005). A cellular Drosophila melanogaster protein with similarity to baculovirus F envelope fusion proteins. J. Virol. 79(13):7979–7989. Malik, H. S., and Henikoff, S. (2005). Positive selection of Iris, a retroviral envelope-derived host gene in Drosophila melanogaster. PLoS Genet. 1(4):e44. Malik, H. S., Henikoff, S., and Eickbush, T. H. (2000). Poised for contagion: Evolutionary origins of the infectious abilities of invertebrate retroviruses. Genome Res. 10(9):1307–1318. Marlor, R. L., Parkhurst, S. M., and Corces, V. G. (1986). The Drosophila melanogaster gypsy transposable element encodes putative gene products homologous to retroviral proteins. Mol. Cell Biol. 6(4):1129–1134. Marques, J. T., and Carthew, R. W. (2007). A call to arms: Coevolution of animal viruses and host innate immune responses. Trends Genet. 23(7):359–364. Mejlumian, L., Pelisson, A., Bucheton, A., and Terzian, C. (2002). Comparative and functional studies of Drosophila species invasion by the gypsy endogenous retrovirus. Genetics 160(1):201–209. Mevel-Ninio, M., Pelisson, A., Kinder, J., Campos, A. R., and Bucheton, A. (2007). The flamenco locus controls the gypsy and ZAM retroviruses and is required for Drosophila oogenesis. Genetics 175(4):1615–1624.
Drosophila Viruses and the Study of Antiviral Host-Defense
263
Misseri, Y., Cerutti, M., Devauchelle, G., Bucheton, A., and Terzian, C. (2004). Analysis of the Drosophila gypsy endogenous retrovirus envelope glycoprotein. J. Gen. Virol. 85(Pt. 11): 3325–3331. Moore, N. F., Kearns, A., and Pullin, J. S. (1980). Characterization of cricket paralysis virusinduced polypeptides in Drosophila cells. J. Virol. 33(1):1–9. Moscat, J., Diaz-Meco, M. T., and Wooten, M. W. (2007). Signal integration and diversification through the p62 scaffold protein. Trends Biochem. Sci. 32(2):95–100. Muller, S., and Imler, J. L. (2007). Dicing with viruses: MicroRNAs as antiviral factors. Immunity 27(1):1–3. Nagy, E., and Dobos, P. (1984). Synthesis of Drosophila X virus proteins in cultured Drosophila cells. Virology 134:358–367. O’Donnell, K. A., and Boeke, J. D. (2007). Mighty Piwis defend the germline against genome intruders. Cell 129(1):37–44. Ohanessian, A., and Echalier, G. (1967). Multiplication of the Sindbis virus in Drosophila melanogaster (dipterous insect) in experimental conditions. C. R. Acad. Sci. Hebd. Seances Acad. Sci. D 264(10):1356–1358. Otsuka, M., Jing, Q., Georgel, P., New, L., Chen, J., Mols, J., Kang, Y. J., Jiang, Z., Du, X., Das, S. C., Pattnaik, A. S., Beutler, B., et al. (2007). Hypersusceptibility to vesicular stomatitis virus infection in dicer-deficient mice is due to impaired miR24 and miR93 expression. Immunity 27:123–134. Pan, J., Vakharia, V. N., and Tao, Y. J. (2007). The structure of a birnavirus polymerase reveals a distinct active site topology. Proc. Natl. Acad. Sci. USA 104(18):7385–7390. Pane, A., Wehr, K., and Schupbach, T. (2007). Zucchini and squash encode two putative nucleases required for rasiRNA production in the Drosophila germline. Dev. Cell 12 (6):851–862. Pattnaik, A. K., Hwang, L., Li, T., Englund, N., Mathur, M., Das, T., and Banerjee, A. K. (1997). Phosphorylation within the amino-terminal acidic domain I of the phosphoprotein of vesicular stomatitis virus is required for transcription but not for replication. J. Virol. 71(11):8167–8175. Pelisson, A., Sarot, E., Payen-Groschene, G., and Bucheton, A. (2007). A novel repeatassociated small interfering RNA-mediated silencing pathway downregulates complementary sense gypsy transcripts in somatic cells of the Drosophila ovary. J. Virol. 81 (4):1951–1960. Pfeffer, S., Sewer, A., Lagos-Quintana, M., Sheridan, R., Sander, C., Grasser, F. A., van Dyk, L. F., Ho, C. K., Shuman, S., Chien, M., Russo, J. J., Ju, J., et al. (2005). Identification of microRNAs of the herpesvirus family. Nat. Methods 2(4):269–276. Plus, N., Croizier, G., Duthoit, J. L., David, J., Anxolabehere, D., and Periquet, G. (1975a). The discovery, in Drosophila, of viruses belonging to three new groups. C. R. Acad. Sci. Hebd. Seances Acad. Sci. D 280(12):1501–1504. Plus, N., Croizier, G., Jousset, F. X., and David, J. (1975b). Picornaviruses of laboratory and wild Drosophila melanogaster: Geographical distribution and serotypic composition. Ann. Microbiol. (Paris) 126(1):107–117. Prud’homme, N., Gans, M., Masson, M., Terzian, C., and Bucheton, A. (1995). Flamenco, a gene controlling the gypsy retrovirus of Drosophila melanogaster. Genetics 139(2):697–711. Rohrmann, G. F., and Karplus, P. A. (2001). Relatedness of baculovirus and gypsy retrotransposon envelope proteins. BMC Evol. Biol. 1:1. Rose, J., and Whitt, M. (2001). Rhabdoviridae: The viruses and their replication. In Fields Virology, (D. Knipe and P. Howley, Eds.), 4th Ed., Vol. 1, pp. 1221–1244. 2 vols. Lippincott Williams & Wilkins, Philadelphia, PA. Rubin, G. M., and Lewis, E. B. (2000). A brief history of Drosophila’s contributions to genome research. Science 287(5461):2216–2218.
264
Tu¨nde Huszar and Jean-Luc Imler
Sabatier, L., Jouanguy, E., Dostert, C., Zachary, D., Dimarcq, J. L., Bulet, P., and Imler, J. L. (2003). Pherokine-2 and -3: Two Drosophila molecules related to pheromone/odor-binding proteins induced by viral and bacterial infections. Eur. J. Biochem. 270:3398–3407. Sasaki, J., and Nakashima, N. (2000). Methionine-independent initiation of translation in the capsid protein of an insect RNA virus. Proc. Natl. Acad. Sci. USA 97(4):1512–1515. Schneemann, A., Reddy, V., and Johnson, J. E. (1998). The structure and function of nodavirus particles: A paradigm for understanding chemical biology. Adv. Virus Res. 50:381–446. Schott, D. H., Cureton, D. K., Whelan, S. P., and Hunter, C. P. (2005). An antiviral role for the RNA interference machinery in Caenorhabditis elegans. Proc. Natl. Acad. Sci. USA 102 (51):18420–18424. Schuler, M., Connell, S. R., Lescoute, A., Giesebrecht, J., Dabrowski, M., Schroeer, B., Mielke, T., Penczek, P. A., Westhof, E., and Spahn, C. M. (2006). Structure of the ribosome-bound cricket paralysis virus IRES RNA. Nat. Struct. Mol. Biol. 13 (12):1092–1096. Scott, M. P., Fostel, J. M., and Pardue, M. L. (1980). A new type of virus from cultured Drosophila cells: Characterization and use in studies of the heat-shock response. Cell 22 (3):929–941. Scotti, P. D., Dearing, S., and Mossop, D. W. (1983). Flock House virus: A nodavirus isolated from Costelytra zealandica (White) (Coleoptera: Scarabaeidae). Arch. Virol. 75(3):181–189. Shen, M., Cui, L., Ostiguy, N., and Cox-Foster, D. (2005). Intricate transmission routes and interactions between picorna-like viruses (Kashmir bee virus and sacbrood virus) with the honeybee host and the parasitic varroa mite. J. Gen. Virol. 86(Pt. 8):2281–2289. Shwed, P. S., Dobos, P., Cameron, L. A., Vakharia, V. N., and Duncan, R. (2002). Birnavirus VP1 proteins form a distinct subgroup of RNA-dependent RNA polymerases lacking a GDD motif. Virology 296(2):241–250. Song, S. U., Gerasimova, T., Kurkulos, M., Boeke, J. D., and Corces, V. G. (1994). An env-like protein encoded by a Drosophila retroelement: Evidence that gypsy is an infectious retrovirus. Genes Dev. 8(17):2046–2057. Spahn, C. M., Jan, E., Mulder, A., Grassucci, R. A., Sarnow, P., and Frank, J. (2004). Cryo-EM visualization of a viral internal ribosome entry site bound to human ribosomes: The IRES functions as an RNA-based translation factor. Cell 118(4):465–475. Tate, J., Liljas, L., Scotti, P., Christian, P., Lin, T., and Johnson, J. E. (1999). The crystal structure of cricket paralysis virus: The first view of a new virus family. Nat. Struct. Biol. 6(8):765–774. Teninges, D., Ohanessian, A., Richard-Molard, C., and Contamine, D. (1979). Isolation and biological properties of drosophila X virus. J. Gen. Virol. 42:241–254. Teninges, D., Bras, F., and Dezelee, S. (1993). Genome organization of the sigma rhabdovirus: Six genes and a gene overlap. Virology 193(2):1018–1023. Terzian, C., Ferraz, C., Demaille, J., and Bucheton, A. (2000). Evolution of the Gypsy endogenous retrovirus in the Drosophila melanogaster subgroup. Mol. Biol. Evol. 17 (6):908–914. Terzian, C., Pelisson, A., and Bucheton, A. (2001). Evolution and phylogeny of insect endogenous retroviruses. BMC Evol. Biol. 1:3. Teysset, L., Burns, J. C., Shike, H., Sullivan, B. L., Bucheton, A., and Terzian, C. (1998). A Moloney murine leukemia virus-based retroviral vector pseudotyped by the insect retroviral gypsy envelope can infect Drosophila cells. J. Virol. 72(1):853–856. Thoetkiattikul, H., Beck, M. H., and Strand, M. R. (2005). Inhibitor kappaB-like proteins from a polydnavirus inhibit NF-kappaB activation and suppress the insect immune response. Proc. Natl. Acad. Sci. USA 102(32):11426–11431.
Drosophila Viruses and the Study of Antiviral Host-Defense
265
Thoulouze, M. I., Bouguyon, E., Carpentier, C., and Bremont, M. (2004). Essential role of the NV protein of Novirhabdovirus for pathogenicity in rainbow trout. J. Virol. 78 (8):4098–4107. Tolia, N. H., and Joshua-Tor, L. (2007). Slicer and the argonautes. Nat. Chem. Biol. 3(1):36–43. van Rij, R. P., Saleh, M. C., Berry, B., Foo, C., Houk, A., Antoniewski, C., and Andino, R. (2006). The RNA silencing endonuclease Argonaute 2 mediates specific antiviral immunity in Drosophila melanogaster. Genes Dev. 20(21):2985–2995. Vidy, A., El Bougrini, J., Chelbi-Alix, M. K., and Blondel, D. (2007). The nucleocytoplasmic rabies virus P protein counteracts interferon signaling by inhibiting both nuclear accumulation and DNA binding of STAT1. J. Virol. 81(8):4255–4263. Wang, X. H., Aliyari, R., Li, W. X., Li, H. W., Kim, K., Carthew, R., Atkinson, P., and Ding, S. W. (2006). RNA interference directs innate immunity against viruses in adult Drosophila. Science 312(5772):452–454. Wayne, M. L., Contamine, D., and Kreitman, M. (1996). Molecular population genetics of ref (2)P, a locus which confers viral resistance in Drosophila. Mol. Biol. Evol. 13(1):191–199. Whelan, S. P. (2008). Response to ‘‘Non-segmented negative-strand RNA virus RNA synthesis in vivo. Virology 371(2):234–237. Wilkins, C., Dishongh, R., Moore, S. C., Whitt, M. A., Chow, M., and Machaca, K. (2005). RNA interference is an antiviral defence mechanism in Caenorhabditis elegans. Nature 436 (7053):1044–1047. Wilson, J. E., Powell, M. J., Hoover, S. E., and Sarnow, P. (2000). Naturally occurring dicistronic cricket paralysis virus RNA is regulated by two internal ribosome entry sites. Mol. Cell. Biol. 20(14):4990–4999. Wyers, F., Dru, P., Simonet, B., and Contamine, D. (1993). Immunological cross-reactions and interactions between the Drosophila melanogaster ref(2)P protein and sigma rhabdovirus proteins. J. Virol. 67(6):3208–3216. Yue, C., and Genersch, E. (2005). RT-PCR analysis of deformed wing virus in honeybees (Apis mellifera) and mites (Varroa destructor). J. Gen. Virol. 86(Pt. 12):3419–3424. Zambon, R. A., Nandakumar, M., Vakharia, V. N., and Wu, L. P. (2005). The Toll pathway is important for an antiviral response in Drosophila. Proc. Natl. Acad. Sci. USA 102 (20):7257–7262. Zambon, R. A., Vakharia, V. N., and Wu, L. P. (2006). RNAi is an antiviral immune response against a dsRNA virus in Drosophila melanogaster. Cell Microbiol. 8(5):880–889.
INDEX A Abu rokab, 12 Actinobacillus pleuropneumonia, 132 Acute viscerotropic disease (YEL-AVD), 83, 85 Adeno-Associated virus (AAV), 185 Adenovirus (Ad) vectors, 92 Aedes spp. Ae. aegypti, 3, 5, 8, 13, 17–18, 21–22, 25–28, 31, 41, 44, 50–54 Ae. aegypti aegypti, 22–23 Ae. aegypti formosus, 22, 24 Ae. africanus, 24 Ae. albolateralis, 23 Ae. albopictus, 21, 23, 25–28, 31, 44, 50 Ae.(Finlaya) niveus s. l., 23 Ae. furcifer, 22, 24, 44–45, 50 Ae. (Gymnometopa) mediovittatus, 25, 28 Ae. luteocephalus, 22, 24, 50 Ae. mediovittatus, 26 Ae. niveoides, 23 Ae. novoniveus, 23 Ae. opok, 24 Ae. (Protomacleaya) triseriatus, 26 Ae. pseudoniveus, 23 Ae. (Stegomyia) polynensiensis, 26 Ae. (Stegomyia) scutellaris, 26, 28 Ae. subniveus, 23 Ae. taylori, 22, 24 Ae. vanus, 23 Ae. vitattus, 22 Alouatta palliata, 25 Alpha-papillomaviruses, 175–176 Alphavax, 105 Antibody-dependent enhancement theory (ADE), 18, 32, 39 Antigenic drift, 128–129, 134 Antigenic shift, 128 Aotus trivirgatus, 25 Aphid lethal paralysis virus, 236 Arboviruses, 27, 32–33, 35, 79, 228
Ateles fusciceps, 25 Ateles geoffroyi, 25 Autogenous SIV vaccines, 139 Avian influenza virus (AIV), 136, 137 B Barbary macaques (Macaca sylvanus), 211 Beijing–1 strain, 86–87 Benzimidazole nucleosides, 214–215 Birnaviridae, 240 Birnaviruses. See Drosophila X virus (DXV) Black Queen Cell Virus, 236 Bovine diarrhea virus, 81 C Caenorhabditis elegans, 252 Canyon, 238 CCAAT displacement protein (CDP), 171–172 Cebus capucinus, 25 ChimeriVax-JE, 101–102 ChimeriVax-WN02, 103 Chimerization, 99–100 Closed mitosis, 178 Costelytra zealandica, 245 Cricket paralysis virus (CrPV), 236, 238, 250, 252 Cytotoxic T lymphocyte (CTL), 215 D Dakar bat, 4 DEN fever (DF), 12, 20–21, 41, 43, 45, 49, 79 Dengue epidemics from 1824–1916, 14 epidemics worldwide from 1922–2007, 15–16 terms used for, 12 Dengue hemorrhagic fever (DHF), 2, 15, 17, 19–21, 39–41, 79
267
268
Index
Dengue shock syndrome (DSS), 79 Dengue viruses (DENV), 2–4, 78–80. See also Sylvatic DENV animal models, 37–39 ChimeriVax-Dengue vaccine, 102 classification of, 5–8 dengue LAV-based chimeras, 99–101 emergence of serotypes, 31–32 evolution origins and emergence, 29–35 rates of, 35–37 of virulence, 37–42 genotypes, by nucleic acid sequencing, 5 history of, 8, 11–13 DEN hyperendemism and DHF, rise of, 19–21 diagnostics and transmission, 13–16 sylvatic DENV, discovery of, 19 World War II, effect of, 17–19 LAVs, for four DENV serotypes, 96–97 population shifts and lineage replacements, 33 primary receptors for, 44–45 recombinant dengue vaccine candidates, 97–99 recombination, 33–35 transmission cycles, 21–22 endemic/epidemic DENV cycles, 26–29 sylvatic DENVcycles, 22–26 vaccines for, 83 virulence, 39–40 DENV–1, 5, 15, 17–19, 22–23, 25, 28–31, 35–37, 45 genotypes of, 5–6 DENV–2, 15–19, 22–26, 25–38, 28–31, 40–43, 54 ex vivo replication profile of, 53 genotypes of, 5–7 in vivo replication profile of, 50–51 DENV–3, 7–9, 15, 18, 20, 23, 28–30, 35, 40, 48 DENV–4, 8, 10, 15, 18, 20, 23, 28, 30–31, 35–37, 48 DENV transovarial transmission (TOT), 27–28 Dicistroviridae, 236–238 D. immigrans, 245 DNA vaccines, 93–96
for dengue, 95 Drosophila, antiviral reactions in inducible response JAK/STAT-dependent immune responses, 254–255 JAK/STAT-independent immune responses, 255–256 RNA interference (RNAi) control of RNA virus infections by, 250–253 endogenous retroviruses control, by piRNAs, 253–254 RNAi pathways, 250 Drosophila C virus (DCV), 236, 250, 252, 254–255 interaction with drosophila, 239–240 replication cycle of, 238–239 virion and genome structure, 236–238 Drosophila F virus (DFV) pathogenesis in drosophila, 244 replication cycle of, 243–244 virion and genome structure, 243 Drosophila melanogaster, 228–229, 234, 236, 243–245, 247, 256–257 Drosophila P virus (DPV), 245 Drosophila S virus (DSV), 244 Drosophila viruses, 230 Drosophila X virus (DXV), 250, 256 and drosophila, interactions between, 242–243 replication cycle, 242 virion and genome structure, 240–242 D. simulans, 244 E EBNA–1 and LANA, herpesvirus tethering proteins, 176–178 Endemic DENV cycle, 26 Endogenous retroviruses, 247 Entebbe bat, 3 Epstein-Barr virus (EBV), 176, 183, 185, 188 Ergosterol, 210 Errantiviridae, 247 Erythrocebus patas, 22, 24–25 F Flamenco, 253–254 Flaviviridae, 78, 81
Index
Flavivirus diseases currently licensed vaccines for, 83 INV and LAV for JE, 86–87 INV for tick-borne encephalitis, 87–88 LAV for yellow fever, 84–86 development of, new vaccines for, 88 ChimeriVax vaccines ChimeriVax-Dengue, 102 ChimeriVax JE, 101–102 ChimeriVax-WN, 103–104 dengue LAV-based chimeras, 99–101 DNA vaccines, 93–96 recombinant LAVs, for dengue, 97–99 subunit vaccines, 89–91 traditional LAVs, 96–97 Vero cell-derived INV for JE, 88–89 virus-vectored vaccines, 91–93 epidemiology of, 79–81 Flaviviruses, 2, 228 animal, infections in, 3 biological structure, 81–83 envelope (E) protein, 82 non-structural proteins, 82 RNA genome, 81 truncated forms of the E protein (trE), 82–83 classification of, 3 human infections, 2–3 phylogenetic tree of, 4 Flock house virus (FHV), 245–246, 250, 252 Formalin-inactivated RhCMV virions (FI-RhCMV), 219–220 Foscarnet (phosphonoformic acid, PFA), 214 French neurotropic vaccine (FNV), 84 G Ganciclovir (GCV), 214 Gypsy and infectious retrotransposons errantiviruses, 247 gypsy endogenous retrovirus, 247–248 infectious abilities, of drosophila retroviruses, 248–249 retrotransposable elements, 246–247
269
H HAd5-SIV vaccine, 146 Hawaii Biotech vaccine, 91 Hemagglutination inhibition (HI) titer, 140 Hemagglutinin (HA), 128–130 Hepacivirus, 81 Hepatitis C virus, 81 Herpes simplex viruses (HSV), 104 Herpesvirus saimiri (HVS), 176 Heterosubtypic immunity (Het-I), 140 Homotypic immunity, 15, 17, 45–46, 140 Host-pathogen interactions, 228 Human adenovirus serotype 5 (HAd5), 145 Human cytomegalovirus (HCMV), 208–209, 220–221. See also Rhesus cytomegalovirus (RhCMV) anti-HCMV drugs, 214–215 genomes, 213 immune modulating proteins, 216–217 infection and disease by, 209 infection in fetuses, 212 transmission of, 211 vaccine, development of, 209–210 Human influenza viruses, 132 I Iflaviruses, 245 IFN a/b signaling cascade, 139 Ilheus virus (ILHV), 3 Imd pathway, 254 Inactivated viral vaccines (INV), 79 Infectious bursal disease virus (IBDV), 240–242 Infectious hematopoietic necrosis virus (IHNV), 232 Infectious pancreatic necrosis virus (IPNV), 241 Influenza A viruses, 128–129, 136 cross-species transmissions in, 136 H5N1 viruses, 136 immune response of mice to, 140–141 nonstructural (NS) proteins, 130 structure of, 129–130 Intercell vaccine. See Vero cell-derived INV Internal ribosomal entry sites (IRES), 238
270
Index
Intranasal (IN) influenza vaccine, 143 Intranuclear inclusions, 210 J JAK-STAT pathway, 254–255 Japanese encephalitis virus ( JEV), 2, 78, 80 JNK pathway, 254 K Kaposi’s sarcoma associated Herpesvirus (KSHV), 176 Kedougou virus, 3 Kunjin virus, 89 Kyasanur Forest disease virus (KFDV), 3 L LaCrosse virus, 27 Laminin 5, 158 Langat virus (LGT), 99 LGT/DEN4 chimera, 99–100 Live-attenuated virus vaccine (LAV), 79 Long control region (LCR), 159 Long terminal repeats (LTR), 247 Louping ill virus (LIV), 3 Lyssaviruses, 229, 232
Monocyte derived dendritic cell (moDC) model, 38–39, 54 Mononegavirales, 232 Mosquito-borne viruses, 3–4. See also Dengue viruses (DENV) Murine cytomegalovirus (MCMV), 219–220 Murine gamma herpesvirus–68 (MHV–68), 176 Murine models, 37 Mycobacterium tuberculosis, 211 Mycoplasma hyopneumoniae, 131–132 N Nakayama-NIH, 86 Neuraminidase (NA), 128–130 NF-kb pathway, 234 Nodaviridae, 245 Non-human primates (NHPs), 85 Nora virus, 245, 257 NS1-mutated SIVs, 143 Nuclear export protein (NEP), 130 O Omsk hemorrhagic fever virus (OHFV), 3 Open reading frames (ORFs), 213–214, 216, 236–238, 240–241, 245 Orthomyxoviridae, 128
M
P
Macaca spp., 23 M. fascicularis, 22 M. mulatta, 45 M. nemestrina, 22 Mammalian virus groups, 3–4 Marikina geoffroyi, 25 Maternal derived antibody (MDA), 144–145 Matrix attachment region (MAR), 178 Maximum likelihood (ML) method, 36, 47 Measles virus (MV), 93 Micro RNAs (miRNAs), 250, 253 Minichromosome Maintenance Element (MME), 178 Mixing vessel, 132 Modified live-virus vaccine (MLV), 141, 143 Modoc virus, 3–4
Pan American Health Organization (PAHO) program, 18 Papillomaviruses, 156 anti-viral replication therapies, 189 chromosomal tethering targets Brd4, 179–182 ChLR1, 182 mitotic spindle, 183 rDNA loci, 182 disease caused by, 156 E1 and E2 binding sites in LCR, 167 E1 and E2 proteins, domain structure of, 162 E1 binding origin sequences, 169 E2 transactivation domain in complex, structure of, 165, 167 genome organization and expression, 159
Index
genome partitioning, mechanism of, 174 life cycle, 157–159 maintenance replication, 172–173 cis elements, role in genome partitioning, 178–179 E1 protein, role of, 173 genome copy number and partitioning, regulation of, 183–185 genome partitioning, in different papillomaviruses, 175–176 other viral tethering proteins, 176–178 replication licensing, 183 role of E2 protein, 173–175 papillomavirus-based vectors, 190 replication cellular DNA replication and, 188 competent environment for, 186–187 differences in strategies of different types, 187–188 modes of, 157, 161 p53 protein and, 171 and replication of other viruses, 188–189 in Saccharomyces cerevisiae,, 189 replication initiation, 161–163 E1 initiator protein, 162–164 E2 loading factor, 164–168 regulation of, 169–172 replication origin, 168–169 S-phase-like state, of host cell, 160 vegetative replication, 184–186 viral proteins, functions of, 160–161 E1, E2, and E4 proteins, 161 E5, E6, and E7 proteins, 160 L1 and L2,capsid antigens, 161 viral tethering proteins, chromosomal targets of, 177 Pasteurella multocida, 132 Pestivirus, 81 Picornaviridae, 236–239, 245 Piwi-associated RNAs (piRNAs), 250–254 Plasmodium falciparum, 42 Porcine circovirus type 2 (PCV–2), 132 Porcine reproductive and respiratory syndrome virus (PRRSV), 132 Porcine respiratory disease complex (PRDC), 131
271
Powassan virus (POWV), 3 Poxviruses, 91 Presbytis spp., 23 P. cristata, 22 P. melaphos, 22 P. obscura, 22 Primary dog kidney (PDK) cells, 87 Primary hamster kidney (PHK) cells, 87 Promyelocytic leukemia protein (PML), 158 R Rabies virus, 229, 231–232 Recombinant modified vaccinia Ankara virus (rMVA), 219 Ref(2)P gene, 234–235 Reoviridae, 228, 243, 256 Reoviruses. See Drosophila F virus (DFV) Replication-defective virus vaccines, 104–105 Replication protein A (RPA), 164 Replicative intermediate (RI), 233 RepliVAX, 106–110 growth, by using two-component genome system, 110–112 infected cell, 107–108 production in C-expressing cells, 107–109 utility of, 109–110 Retrotransposons, 246–247 Reverse genetics approach, 140, 142–143, 147 Rhabdoviridae, 229 Rhabdoviruses. See Sigma virus (SIGMAV) Rhesus cytomegalovirus (RhCMV), 210 coding capacity of, 213–214 discovery of, 210–211 epidemiology of, 211–212 host immunity to, 215–216 modulation of host by, 216–217 pathogenesis of, 212–213 route of transmission, 211 susceptibility to anti-HCMV drugs, 214–215 vaccine studies, 218–220
272
Index
Rhesus macaque (Macaca mulatta), 210 DNA prime/FI-RhCMV boost approach, study of, 219–220 Ribonucleoprotein (RNP), 232–233 Rio Bravo virus, 4 RNA-induced silencing complex (RISC), 250–251 RNase III enzyme Dicer–2, 250–251 Rolling circle replication (RCR), 185 S Saimiri orstedii, 25 Sanofi Pasteur, 102 SA14–14–2 vaccine, 87 SCID-xenograft model, 37–38 Seabird virus groups, 4 Sentinel monkeys, 22–23, 42 Sigma virus (SIGMAV), 229 and drosophila, 234–235 replication cycle of, 232–234 virion and genome structure, 229–232 Simian immunodeficiency virus (SIV), 212 Sindbis virus (SINV), 250 Single-cycle nucleic acid vaccine candidates, 106 Single-cycle virus vaccines, 104–105 Small interfering (si) RNA, 250–253 Sokoluk viruses group, 3 Spanish flu, 130, 134 Stegomyia fasciata, 27 Subunit vaccines, 89–91 Subviral particle (SVP), 81–82 Swine fever virus, 81 Swine influenza (SI). See Swine influenza virus (SIV) Swine influenza virus (SIV), 128 autogenous vaccine, use of, 139 H1 and H3 subtype, evolution in North America of, 132–135 cH1N1 subtype, 132 double reassortant H3N2 virus, 132–133 human-like swine H1 (hu-H1) viruses, spread of, 135 reassortant viruses, evolution of, 133–134 TRIG cassette, acquisition of, 133–134
triple reassortant H3N2 virus, 132–133 in humans, 136–137 novel SI subtypes, in North America H2N3 virus, 138 H4N6 virus, 137–138 in pigs, 130 pneumonia by H1N1 lineage, 130–131 vaccinating pigs against influenza virus, 139–147 Sylvatic DENV, 19, 22, 42–45 amplification cycles, 24–25, 42, 46 reemergence adaptation for transmission, 49–54 epidemics and human contact, 42–44 natural immunity or vaccination, influence of, 44–47 selection pressures, 47–49 spillover outbreaks of, 49 transmission cycles, 22–26, 29, 43 in Americas, 25 in Malaysia, 22–23 in West Africa, 24–25 T TATA-binding protein (TBP), 171 TBE INV by, Neudorfl strain, 87 The pocket, cavity in picornaviruses, 238 Tick-borne encephalitis virus (TBEV), 2–3, 78, 80–81 Tick-borne viruses, 3–4 Toll pathway, 234, 254, 256 Topotypes, 5 Transovarial transmission (TOT), 27 Triple reassortant internal gene (TRIG) cassette, 134 T1 RNase fingerprinting, 5, 7 TX98 NS1D126, as MLV vaccine, 143–145 U Upstream regulatory region (URR). See Long control region (LCR) V Vector-based subunit vaccines, 141, 143 Vector control programs, 46–47 Vegetative DNA replication, 184–186
Index
Venezuelan equine encephalitis virus replicon (VEErep), 107–109 Vero cell-derived INV, 88–89 Vesicular stomatitis virus (VSV), 27, 229, 233 Vesiculoviruses, 229, 232 Viral load, 41, 135, 219–220, 252, 255 Viruses with no known arthropod vector (NKV), 3–4 Virus-vectored vaccines Ad-vectored dengue vaccine, 92–93 ALVAC-JEV, 92 MV-vectored vaccine, 93 NYVAC-JEV, 91–92 W Water poison, 11 Wesselsbron virus (WESSV), 3
273
West Nile virus (WNV), 78, 80, 228 WN/DEN–2 PDK53 chimera, 100–101 World War II, and dengue, 17–19 Y Yellow fever vaccine-associated neurotropic disease (YEL-AND), 84–85 Yellow fever virus (YFV), 2–3, 25, 78, 80 YFV–17D vaccine, 84–86 Yin-yang 1 (YY1), 171–172 Yokose, 3 Z Zika virus (ZIKV), 3 Zone of emergence, 23, 25
