Human Emerging and Re-emerging Infections

Emerging and re-emerging pathogens pose several challenges to diagnosis, treatment, and public health surveillance, primarily because pathogen identification is a difficult and time-consuming process due to the “novel” nature of the agent. Proper identification requires a wide array of techniques, but the significance of these diagnostics is anticipated to increase with advances in newer molecular and nanobiotechnological interventions and health information technology.

Human Emerging and Re-emerging Infections covers the epidemiology, pathogenesis, diagnostics, clinical features, and public health risks posed by new viral and microbial infections. The book includes detailed coverage on the molecular mechanisms of pathogenesis, development of various diagnostic tools, diagnostic assays and their limitations, key research priorities, and new technologies in infection diagnostics. Volume 1 addresses viral and parasitic infections, while volume 2 delves into bacterial and mycotic infections.

Human Emerging and Re-emerging Infections is an invaluable resource for researchers in parasitologists, microbiology, Immunology, neurology and virology, as well as clinicians and students interested in understanding the current knowledge and future directions of infectious diseases.

• Language: English
• Publisher: Wiley
• Release date: Nov 9, 2015
• ISBN: 9781118644645

Book preview

Human Emerging and Re-emerging Infections

Viral and Parasitic Infections

Volume I

Edited by

Sunit Kumar Singh

Laboratory of Human Molecular Virology and Immunology

Molecular Biology Unit, Faculty of Medicine, Institute of Medical Sciences

Banaras Hindu University, Varanasi, India

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Copyright © 2016 by John Wiley & Sons, Inc. All rights reserved

Published by John Wiley & Sons, Inc., Hoboken, New Jersey

Published simultaneously in Canada

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Library of Congress Cataloging-in-Publication Data:

Human emerging and re-emerging infections / edited by Sunit K. Singh.

      p. ; cm.

   Includes index.

   ISBN 978-1-118-64471-3 (cloth)

   I. Singh, Sunit K., editor.

   [DNLM: 1. Communicable Diseases, Emerging.   WA 110]

   RA643

   616.9--dc23

2015028631

oBook ISBN: 9781118644843

ePDF ISBN: 9781118644829

ePub ISBN: 9781118644645

Dedicated to my parents

Contents

List of Contributors

Preface

Acknowledgments

About the Editor

Chapter 1 Pathogenesis of the Old World Arenaviruses in Humans

1.1 The Old World arenaviruses: taxonomic and zoonotic introduction

1.2 Genome structure

1.3 Arenavirus replication strategy

1.4 Epidemiology

1.5 Pathogenesis of disease induced by Old World arenaviruses

1.6 Treatment and prevention

Notes in the Proofs

Acknowledgement

References

Chapter 2 Pathogenesis of New World Arenaviruses in Humans

2.1 Introduction

2.2 Biology of arenaviruses

2.3 Pathogenesis and immunity

2.4 Arenavirus population dynamics

2.5 Receptor use and emergence/disease potential for the NW arenaviruses

2.6 New trends in antiviral treatments

Acknowledgments

References

Chapter 3 Pathogenesis of Emerging and Novel Bunyaviruses in Humans

3.1 Introduction

3.2 Phlebovirus

3.3 Hantavirus

3.4 Orthobunyavirus

3.5 Nairovirus

References

Chapter 4 Pathogenesis of Rift Valley Fever in Humans

4.1 Introduction

4.2 Classification of RVFV

4.3 Genome structure of RVFV

4.4 Virion structure of RVFV

4.5 Viral life cycle of RVFV

4.6 Virulence factors for RVFV

4.7 Natural reservoirs and transmission cycles of RVFV

4.8 Risk factors for RVF outbreaks

4.9 Clinical features of human RVF

4.10 Animal models for RVF

4.11 Pathogenesis of RVF

4.12 Diagnosis of RVF

4.13 Treatment of RVF

4.14 RVF vaccines

4.15 Conclusions

Acknowledgments

References

Chapter 5 Pathogenesis of Hantavirus Infections

5.1 Introduction

5.2 Hantaviruses: genome organization, virion structure, and replication

5.3 Epidemiology

5.4 Hantaviruses in shrews and bats

5.5 Clinical features

5.6 Pathogenesis and immunity

5.7 Diagnosis

5.8 Treatment

5.9 Prevention and control

5.10 Conclusion

References

Chapter 6 Molecular Pathogenesis of Japanese Encephalitis Virus Infection

6.1 Introduction

6.2 Genome organization of JEV

6.3 JEV replication

6.4 JEV pathogenesis

6.5 Host response to JEV infection

6.6 Clinical manifestations and diagnosis

6.7 Treatment and preventive measures

6.8 Conclusion

Acknowledgments

References

Chapter 7 Dengue Virus Infection in Humans: Epidemiology, Biology, Pathogenesis, and Clinical Aspects

7.1 Epidemiology

7.2 The vector

7.3 Pathogen

7.4 Clinical manifestations

7.5 Pathogenesis

7.6 Laboratory diagnosis and monitoring

7.7 Treatment

7.8 Prevention and control

References

Chapter 8 Pathogenesis of West Nile Virus in Humans

8.1 Introduction

8.2 The pathogen

8.3 Transmission, epidemiology, and surveillance

8.4 Clinical manifestations and prognosis

8.5 WNV dissemination within the host

8.6 Immune response in mammals

8.7 WNV virulence factors

8.8 WNV therapeutics and disease control

8.9 Conclusion

References

Chapter 9 Overview on Chikungunya Virus Pathogenesis

9.1 Introduction

9.2 Transmission of chikungunya virus

9.3 Genome organization and replication in CHIKV

9.4 Pathogenesis of chikungunya virus

9.5 Chikungunya virus tissue tropism

9.6 Chikungunya virus and host immune interaction

9.7 Clinical manifestations

9.8 Diagnosis, treatment and preventive measures of chikungunya infection

9.9 Conclusion

Acknowledgement

References

Chapter 10 Nipah Virus Infections in Humans

10.1 Introduction

10.2 Virological and biological characteristics of NiV

10.3 Epidemiological features of NiV

10.4 Clinical features of NiV infections

10.5 Pathological findings of NiV infections

10.6 Diagnosis of NiV infections

10.7 Current laboratory diagnostic capabilities

10.8 Diagnostic challenges

10.9 Treatment of NiV infections

10.10 NiV outbreak control and prevention

References

Chapter 11 Pathogenesis of Hendra Virus in Humans

11.1 Introduction

11.2 Epidemiology

11.3 Bats are the natural reservoir of HeV

11.4 Transmission of HeV

11.5 Clinical features of human HeV infection

11.6 Pathogenesis

11.7 Animal models of HeV infection

11.8 Diagnosis

11.9 Vaccination and treatment

11.10 Prevention and control

References

Chapter 12 Pathogenesis of Rotavirus in Humans

12.1 Introduction

12.2 Rotavirus replication cycle

12.3 Epidemiology

12.4 Clinical manifestations

12.5 Pathophysiology

12.6 Immune response

12.7 Diagnosis

12.8 Treatment

12.9 Prevention

12.10 Other RV vaccines

References

Chapter 13 Pathogenesis of Papillomaviruses in Humans

13.1 Introduction

13.2 Papillomavirus diversity and epithelial tropisms

13.3 Virus structure and genome organization

13.4 Natural history of disease

13.5 Clinical features of HPV-associated disease

13.6 Pathogenesis and immunity

13.7 Prevention, detection, and treatment of HPV-associated disease at the cervix and other sites

13.8 Conclusion

Acknowledgments

References

Chapter 14 Kaposi's Sarcoma–Associated Herpesvirus Pathogenesis

14.1 Introduction

14.2 KSHV particle

14.3 KSHV-associated diseases

14.4 Epidemiology, prevalence, and transmission

14.5 KSHV genome

14.6 KSHV replication

14.7 KSHV treatment

Acknowledgments

References

Chapter 15 Microsporidiosis in Humans

15.1 Introduction

15.2 Epidemiology

15.3 Host response

15.4 Clinical features of microsporidiosis

15.5 Central nervous system infection

15.6 Ocular infection

15.7 Musculoskeletal infection

15.8 Cardiac infection

15.9 Sinus and respiratory infection

15.10 Skin infection

15.11 Genitourinary tract infection

15.12 Diagnosis

15.13 Treatment

15.14 Prophylaxis and prevention

Acknowledgments

References

Chapter 16 Pathogenesis of Toxoplasma gondii in Humans

16.1 Pathogen epidemiology

16.2 Parasite developmental biology

16.3 Transmission

16.4 Tachyzoite, tissue cyst, and oocyst biology

16.5 Pathogenesis and clinical features

16.6 Immunity

16.7 Laboratory diagnosis

16.8 Treatment

16.9 Prevention and control

References

Chapter 17 Pathogenesis of Human African Trypanosomiasis

17.1 Introduction

17.2 Pathogen

17.3 Specific pathogenic defense mechanisms

17.4 Epidemiology

17.5 Clinical features

17.6 Pathogenesis and immunity

17.7 Diagnosis

17.8 Treatment

17.9 Prevention and control

17.10 Conclusion

References

Chapter 18 Pathogenesis of Leishmaniasis in Humans

18.1 Introduction

18.2 Life cycle of Leishmania

18.3 Genome organization

18.4 Gene regulation

18.5 Differentiation in Leishmania

18.6 Metabolism

18.7 Clinical features of leishmaniasis

18.8 Epidemiology of leishmaniasis

18.9 Pathogenesis and immunity

18.10 Diagnosis and treatment

18.11 Prevention and control

Note

References

Chapter 19 Pathogenesis of Chagas Disease in Humans

19.1 Introduction

19.2 Epidemiology of Chagas disease

19.3 Physiopathology of the infection by T. cruzi

19.4 Clinical features

19.5 Challenges in Chagas disease

References

Chapter 20 Pathogenesis of Cryptosporidium in Humans

20.1 Introduction

20.2 Life cycle

20.3 Cryptosporidium species

20.4 Genomics of Cryptosporidium species

20.5 Epidemiology

20.6 Clinical features

20.7 Pathogenesis and immunity

20.8 Immune responses to Cryptosporidium infection

20.9 Cryptosporidium and cancer (some experimental evidence)

20.10 Diagnosis

20.11 Treatment

20.12 Control and prevention

References

Chapter 21 Pathogenesis of Malarial Parasites in Humans

21.1 Current global malaria situation

21.2 Plasmodium falciparum

21.3 Epidemiology

21.4 Clinical manifestation and disease pathogenesis

21.5 Groups at risk of severe malaria

21.6 Immunity

21.7 Protective immunity in malaria

21.8 Plasmodium vivax

21.9 Plasmodium malariae

21.10 Plasmodium ovale

21.11 Plasmodium knowlesi

21.12 Detection

21.13 Diagnosis

21.14 Treatment

21.15 Prevention and control

21.16 Emerging challenges

21.17 Re-emergence of malaria

21.18 Conclusion

References

Chapter 22 Pathogenesis of Trichomonas vaginalis in Humans

22.1 Introduction

22.2 T. vaginalis morphology

22.3 T. vaginalis genome

22.4 Pathogenesis and immunity

22.5 Diagnostic methods

22.6 Treatment

22.7 Epidemiology

22.8 Clinical features

22.9 Prevention and control

Acknowledgments

References

Chapter 23 Loa loa Pathogenesis in Humans

23.1 Introduction

23.2 The pathogen: L. loa

23.3 The vector

23.4 Epidemiology of loiasis

23.5 Clinical symptoms and features

23.6 Host–parasite relationship in loiasis infection

23.7 Cellular immune response in loiasis infection

23.8 Humoral immune response in loiasis

23.9 Target antigens for immune response in loiasis

23.10 Mechanism of evasion to immune attack in loiasis

23.11 Concomitant immunity

23.12 Effector mechanisms

23.13 Protective immunity

23.14 Clinical diagnosis

23.15 Biological diagnosis

23.16 Treatment of uncomplicated loiasis

23.17 Treatment of loiasis encephalopathy

23.18 Prevention and control

23.19 Conclusion

Acknowledgment

References

Chapter 24 Nematode Larva Migrans

24.1 Introduction

24.2 Toxocariasis

24.3 Baylisascaris procyonis

24.4 Gnathostomiasis

24.5 Cutaneous larva migrans

Acknowledgments

References

Chapter 25 Pathogenesis of Human Schistosomiasis

25.1 Introduction

25.2 Historical background

25.3 Epidemiology

25.4 Parasite

25.5 The schistosome genome

25.6 Pathogenesis and immunity

25.7 Clinical features

25.8 Diagnosis

25.9 Treatment

25.10 Prevention and control

25.11 Vaccine research

References

Chapter 26 Vector-Borne Parasitic Zoonotic Infections in Humans

26.1 Introduction

26.2 Protozoan vector-borne parasitic zoonosis

26.3 Helminthic vector-borne parasitic zoonotic infections

26.4 Future trends and research needs

References

Index

EULA

List of Tables

Chapter 4

Table 4.1

Chapter 5

Table 5.1

Chapter 7

Table 7.1

Chapter 8

Table 8.1

Table 8.2

Chapter 9

Table 9.1

Chapter 11

Table 11.1

Table 11.2

Table 11.3

Chapter 12

Table 12.1

Chapter 15

Table 15.1

Table 15.2

Chapter 18

Table 18.1

Chapter 19

Table 19.1

Table 19.2

Table 19.3

Table 19.4

Table 19.5

Table 19.6

Table 19.7

Table 19.8

Table 19.9

Chapter 20

Table 20.1

Table 20.2

Table 20.3

Table 20.4

Table 20.5

Chapter 22

Table 22.1

Chapter 24

Table 24.1

Chapter 26

Table 26.1

List of Illustrations

Chapter 1

Fig. 1.1 Estimated global burden of viral hemorrhagic fevers (approximately 500,000 cases per year): DF, Dengue HF and Dengue shock syndrome; LF, Lassa Fever; YF, Yellow fever; HFRS, HF fever with renal syndrome; NWA, HF caused by OW and NW arenaviruses; RVF, Rift Valley fever; CCHF, Congo-Crimean HF; KFD, Kyasanur forest disease; OHF, Omsk HF; E/MHF, Ebola and Marburg HF; SFTS, severe fever with thrombocytopenia syndrome. From Falzarano and Feldmann, 2013. With permission from Elsevier.

Fig. 1.2 Electron microscopy of Lassa virus. (a) LASV purified in isopycnic sucrose gradient, negative staining (×200,000). (b, c) Ultrathin sections of infected Vero cells. A particle which appears to be budding from plasma membrane (B, ×170,000) and the discrete area of membrane thickening associated with viral buds (c, ×30,000). Arrows indicate LASV glycoprotein on the virus surface (a, b) or in the cell budding site; (d) arenavirus ambisense replication strategy. Genes and antigenomic RNAs are shown as open boxes separated by intergenic areas; subgenomic mRNAs are in black.

Fig. 1.3 Risk factors for rodent-to-human transmission. Three major risk factors favored LASV transmission in highly endemic Gueckedou prefecture of forest Guinea: rodent infestation, open food storage, and hunting peridomestic rodents. Pita prefecture is a mountain area with low LASV prevalence. In the high prevalence region, Gueckedou, hearing problems, the most often complication of LF, were associated with rodent consumption (from ter Meulen et al., 1966, with modifications).

Fig. 1.4 Thrombomodulin (THBD) in LASV infection. LASV infects endothelial cells, which are the main THBD producers. Increased THBD expression will capture thrombin through its 4, 5, and 6 domains, activating protein C (aPC), inhibiting the coagulation pathway, and activating PAR1 membrane protein. PAR-1 has cytoprotective and anti-inflammatory effects, also inhibiting platelet activation. From Zapata et al. (2013b).

Chapter 2

Fig. 2.1 Arenavirus particle and arenavirus GPC processing. (a) Schematic of an arenavirus particle. The viral proteins GP1/GP2, Z, and L are indicated. NP and the viral RNA form the ribonucleoprotein (RNP). (b) Processing of GPC by subtilisin kexin isozyme 1 (SKI-1)/site 1 protease (S1P). For details, please see section 2.2.

Fig. 2.2 Scheme of a quasispecies. RNA viruses replicate as complex mutant clouds. Positive and negative selection with intervening bottlenecks has built the current arenavirus types.

Fig. 2.3 Scheme of a stable quasispecies and the entry in error catastrophe.

Chapter 4

Fig. 4.1 Genome structure of RVFV, and the gene expression strategy: S-segment is an ambisense genome and expresses N from viral-sense RNA and NSs from antiviral-sense RNA. M-segment encodes a single M mRNA, and NSm, 78-kD, Gn, and Gc proteins are synthesized by leaky scanning of 5 AUGs located upstream of Gn, and co-translational cleavage between NSm and Gn or Gn and Gc. L-segment encodes L-protein (RNA-dependent RNA polymerase).

Fig. 4.2 RVFV NSs (MP-12 strain) forms filaments in nuclei, while it also accumulates in cytoplasm. PKR0/0 MEF cells were transfected with in vitro synthesized RNA encoding RVFV MP-12 NSs, and stained with anti-RVFV antibody. (a) RVFV NSs, (b) DAPI, (c) merging of NSs-expressing cells, or (d) merging of control untreated cells.

Fig. 4.3 Clinical signs of RVF patients. Adapted from Ikegami and Makino (2011).

Chapter 5

Fig. 5.1 Geographical representation of approximate hantaviral disease incidence by country per year.

Fig. 5.2 Phylogenetic tree of representative hantavirus species with primary reservoir hosts. Numbers given at the nodes are posterior probability values.

Chapter 6

Fig. 6.1 Organization of the Flavivirus genus. This organization is based on phylogenetic relationships. The serological relationships and arthropod vectors are shown.

Fig. 6.2 Life cycle of Japanese encephalitis virus. JEV is maintained in a zoonotic cycle (black arrow) involving pigs, ardeid birds, and mosquitoes. Pigs are the major amplification/reservoir host, birds act as carriers, and Culex species mosquitoes are the vector. Humans are accidental hosts (dotted red arrow) and get infected through mosquito bite but cannot sustain high virus titer for further transmission. Reprinted from Unni et al. (2011) with permission from Elsevier. Copyright (2013).

Fig. 6.3 Genome organization of Flavivirus along with the products of processing of the polyprotein. NCR, noncoding region; C, capsid; prM, premembrane; E, envelope; NS, nonstructural.

Fig. 6.4 Replication in Japanese encephalitis virus. Schematic representation of the different steps involved in entry, replication, maturation, and release of JEV from the host cells: 1, interaction of JEV with host cell receptor; 2, receptor-mediated endocytosis of virus; 3, fusion of viral and host cell membrane; 4, release of the viral genome into the cytoplasm; 5, formation of RC and cyclization of JEV genome; 6, formation of dsRF; 7, synthesis of viral genome in a semi-conservative and asymmetric manner; 8, translation and processing of JEV genome producing three structural and seven nonstructural proteins; 9, JEV proteins associate with the RC and further assist viral replication; 10, maturation of virions in the Golgi complex; 11, release of mature virus. Reprinted from Unni et al. (2011) with permission from Elsevier. Copyright (2013).

Chapter 7

Fig. 7.1 Factors that contribute to the emergence and spread of DENV infection globally.

Fig. 7.2 Distribution of dengue in countries around the world. The countries with dengue and areas at risk are depicted in gray. The information is based on reports from WHO in 2011 and CDC's Dengue Map as of July 30, 2013.

Fig. 7.3 Schematic representation of DENV genome. The single ORF encodes three structural proteins (capsid (C), membrane (M), and envelope (E)) and seven nonstructural proteins (NS1, NS2A, NS2B, NS3, NS4A, NS4B, and NS5). The 5′ and 3′ ends are flanked by untranslated regions (UTRs).

Fig. 7.4 The incubation period of DENV infection following a mosquito. The incubation period following a mosquito bite is short, usually 3–5 days. The febrile phase is characterized by high fever (bi-phasic), rash, severe headache, retro-orbital pain, myalgia, arthralgia, and gastrointestinal distress. The febrile phase may be accompanied by high viremia, mild hemorrhagic manifestations like petechiae (positive tourniquet test), easy or spontaneous bruising, and mucosal membrane bleeding. The hemorrhage seen during the febrile phase is caused by increased vascular fragility (Martina et al., 2009). The critical phase is usually seen at defervescence (body temperature 38°C or less). This phase is characterized by increase in capillary permeability, resulting in significant plasma leakage localized mainly in the chest and/or abdominal cavities. Adapted with permission from Martina (2011).

Fig. 7.5 Petechiae on the forearm of a patient with dengue. Levels of viremia and cytokines as well as a disturbed hemostasis could explain hemorrhages seen in the early stage of DENV infection.

Fig. 7.6 The WHO 2009 disease classification based on severity of disease.

Fig. 7.7 Factors that influence disease outcome. Disease is the outcome of virus−host interaction. The immune response and hemostatic response to infection is influenced by the environment, host genetics, age, gender, and presence of other underlying diseases.

Fig. 7.8 Summary of laboratory diagnostic options in patients suspected with DENV infection. Day 0 is the onset of disease symptoms. In clinical practice, diagnosis of dengue is confirmed when the result of viral nucleic acid or antigen detection (NS1) or antibody (IgM seroconversion in paired sera) test is positive. During the course of infection, hematological changes can be measured.

Chapter 8

Fig. 8.1 Scale representation of the WNV (strain NY-3568) genome. The position and major functions of the structural and nonstructural proteins and the UTRs are indicated. Co- and post-translational cleavage by the indicated proteases liberates the individual proteins.

Fig. 8.2 WNV particle. (a) Schematic cross-section of a mature WNV particle. (b) Cryo-EM reconstruction of a mature WNV particle from the RCSB PDB (www.rcsb.org) of PDB ID 3j0b (Zhang et al., 2013).

Fig. 8.3 Replication cycle of WNV. The virus lifecycle begins when WNV attaches and enters a host cell. Subsequent translation of the uncoated viral genome produces a single polyprotein that is cleaved into the 10 individual proteins. The nonstructural proteins mediate the replication of the viral genome, which is assembled into virus particles at the ER membrane. Progeny virions are then exported out of the cell through the secretory pathway.

Fig. 8.4 Transmission cycle of WNV. WNV is maintained in nature in a transmission cycle between birds and mosquitos. Human and other mammals do not participate in transmission and are therefore considered dead-end hosts.

Fig. 8.5 Cell-intrinsic innate antiviral response. Schematic of the key signaling pathways contributing to the innate antiviral response to WNV. RLRs, retinoic acid-inducible gene (RIG-I)-like receptors; TLRs, Toll-like receptors; MDA5, melanoma differentiation-associated protein 5; NLRs, nucleotide-binding oligomerization domain (NOD)-like receptors; NLPR3, NLR family PYD-containing 3; TRIF, TIR-domain-containing adapter-inducing interferon-β (TRIF); MyD88, myeloid differentiation primary response 88; ISGs, interferon-stimulated genes.

Fig. 8.6 Schematic of the type I interferon signaling pathway. IFNAR, IFN-α receptor; JAK1, Janus kinase 1; Tyk2, tyrosine kinase 2; STAT, signal transducer and activation of transcription proteins; IRF-9, interferon regulatory factor 9; ISGF3, interferon stimulated gene factor 3; ISRE, interferon-sensitive response element; ISGs, interferon-stimulated genes; PKR, protein kinase R; OAS, 2′-5′-linked oligoadenylate synthase, IFITM, IFN-induced protein with tetratricopeptide repeats 1.

Chapter 9

Fig. 9.1 Maintenance of the chikungunya virus in Africa showing the interconnection between the sylvatic cycle on the left and the urban cycle on the right. Particularly in Africa, the virus is maintained in a sylvatic cycle comprising non-human primates and different species of forest-dwelling mosquitoes including Aedine mosquitoes (Ae. Africanus, Ae. furcifer-taylori, Ae. dalzielii, etc.), and non-Aedine mosquitoes (Mansonia, Culex, etc.). Adapted from Thiboutot et al., PLoS Negl Trop Dis. 2010 April;4(4):e623.

Fig. 9.2 Life cycle of the chikungunya virus. Characteristically, there are two rounds of translation: (+) sense genomic RNA (49S9 = 11.7 kb) acts directly as mRNA and is partially translated (59 end) to produce non-structural proteins (nsp's). These proteins are responsible for replication and formation of a complementary (2) strand, the template for further (+) strand synthesis. Subgenomic mRNA (26 S = 4.1 kb) replication occurs through the synthesis of full-length (2) intermediate RNA, which is regulated by nsp4 and p123 precursor in early infection and later by mature nsp's. Translation of the newly synthesized sub-genomic RNA results in production of structural proteins such as Capsid and protein E2-6k-E1 (from 39 end of genome). Assembly occurs at the cell surface, and the envelope is acquired as the virus buds from the cell and release and maturation almost simultaneous occurred. Replication occurs in the cytoplasm and is very rapid (4 h). Adapted from Thiboutot et al., PLoS Negl Trop Dis. 2010 April;4(4):e623.

Fig. 9.3 Genome Organization of CHIKV. CHIKV genome possesses 5′ cap and 3′ poly (A) tail structures resembling to eukaryotic m RNA. The non-translated regions are existing at 5′ and 3′ ends. Junction (J) region is noncoding region. The subgenomic +ve strand mRNA (also called as 26S RNA) is a product of transcription of negative stranded RNA intermediates. The 26S RNA serves as the mRNA for the synthesis of viral structural proteins. Non-structural proteins: nsP1-nsP4 and structural proteins: C, E1, E2, E3 and 6K generate by proteolytic clevage of polyprotein precursors.

Chapter 10

Fig. 10.1 Unrooted phylogenetic tree based on complete genome nucleotide sequences of selected paramyxoviruses. Virus name (abbreviation) and GenBank accession numbers are as follows: Avian paramyxovirus 2 (APMV-2) HQ896023, Avian paramyxovirus 6 (APMV-6) AY029299, Bovine parainfluenza virus 3 (bPIV3) AF178654, Canine distemper virus (CDV) AF014953, Cedar virus (CedPV) JQ001776, Dolphin morbillivirus (DMV) NC005283, Hendra virus (HeV) AF017149, Human parainfluenza virus 1 (hPIVl)NC003461, Human parainfluenza virus 2 (hPIV2) NC003443, Human parainfluenza virus 3 (hPIV3) Z11575, Human parainfluenza virus 4a (hPIV4a) AB543336, Human parainfluenza virus 4b (hPIV4b) EU627591, Mapuera virus (MprPV) EF095490, Measles virus (MeV) AB016162, Menangle virus (MenPV) AF326114, Mumps virus (MuV) AB000388, Newcastle disease virus (NDV) AF077761, Nipah virus, Malaysian strain (NiV-M) AJ627196, Nipah virus, Bangladesh strain (NiV-B) AY988601, Peste-des-petits-ruminants virus (PPRV) X74443, Porcine mbulavirus (PorPV) BK005918, Phocine distemper virus strain (PDV) KC802221, Rinderpest virus (RPV) Z30697, Sendai virus (SeV) M19661, Simian virus 41 (SV41) X64275, Simian parainfluenza virus 5 (PIV5) AF052755, Tioman virus (TioPV) AF298895.

Fig. 10.2 Electron micrograph of positive stained thin section of Nipah virus in infected Vero cells. The virion (arrow head) is generally spherical but can be fairly pleomorphic in shape. Inclusion body consisting of viral nucleocapsids is found within the cytoplasm of infected cells (solid arrows) and budding of viral ribonueleocapsids occur at the thickened plasma membrane (broken arrows).

Fig. 10.3 Schematically shows the comparative genomic length with the respective proteins and non-coding regions of representative member of each genus within the subfamily Paramyxovirinae. Colored bars represent respective protein(s) coding regions and non-colored bars represent respective leader, trailer, and non-coding regions of the genomes.

Fig. 10.4 Magnetic resonance imaging (MRI) of the brain of a patient with acute Nipah virus infection. Small, disseminated discrete hyperintense lesions showed areas of micro-infarcts (arrows).

Fig. 10.5 Magnetic resonance imaging (MRI) of the brain of a patient suffered from relapsing Nipah virus encephalitis. MRI brain showed areas of confluent cortical involvement, rather than the small, disseminated discrete hyperintense lesions seen in acute Nipah encephalitis.

Fig. 10.6 Vasculitis associated with thrombosis and vascular occlusion in acute Nipah encephalitis (Wong and Ong, 2011).

Fig. 10.7 Endothelial syncytium (arrow) arising from a meningeal blood vessel in acute Nipah encephalitis (Wong and Ong, 2011).

Fig. 10.8 Nipah viral antigens in neurons (arrows) and neuropil in the brain of a patient succumbed to acute Nipah encephalitis (Wong and Ong, 2011).

Chapter 11

Fig. 11.1 Location of Hendra virus spillover events. Each black star represents the location of a spillover event of HeV. Each gray arrow points to a spillover event, where one or more humans were infected.

Fig. 11.2 Progression of Hendra virus infection in the seven human cases. This chart follows the progression of major events in the course of the disease for each of the seven human cases of Hendra virus infection. The case number of each patient corresponds with the case numbers described in Table 11.2.

Fig. 11.3 Chest x-ray of a patient with acute respiratory distress from Hendra virus infection. Reprinted with permission from Selvey et al. (1995).

Fig. 11.4 MRI of a patient with fatal Hendra encephalitis. Axial T2-weighted scans from case #3. Top image is day 6 and bottom image is day 14 of hospitalization. Reprinted with permission from O'Sullivan et al. (1997).

Fig. 11.5 Lesion regression in a survivor of Hendra encephalitis. Axial T2-FLAIR scans from case #6. Left: Day 22, showing subtle focal hyperintense lesion in the left precuneus with white matter sparing. Right: Week 6, demonstrating subsequent regression of lesion. Reprinted with permission from Nakka et al. (2012).

Fig. 11.6 Progressive MRI images track progression of disease in fatal Hendra encephalitis. Axial T2-FLAIR scans from case #5. (a) Two days after hospitalization, only a few cortical lesions were evident. (b) By day 15, the cortical lesions demonstrated marked progression in number and extent, with deep white matter sparing. (c) By day 22, the cortical lesions had progressed further, with early widespread white matter lesions. (d) By day 36, the cortical lesions regressed while the white matter lesions progressed. Reprinted with permission from Nakka et al. (2012).

Fig. 11.7 IHC staining for HeV antigen in AGM tissues after HeV challenge. (a) HeV antigen in endothelium of vessel in the frontal cortex of brain. (b) Strong immunolabeling of HeV antigen in the alveolar tissues. (c) Strong immunolabeling of HeV antigen in the spleen. (d) HeV antigen in the endothelium of the kidney glomeruli.

Chapter 12

Fig. 12.1 Structure of the rotavirus particle. The 11 dsRNA viral segments code for six structural proteins and six non-structural proteins.

Fig. 12.2 Rotavirus replication cycle. After the entry of viral particles into the cell, the virus follows processes of uncoating, transcription, viroplasms formation, replication, assembly, and release of new viral particles.

Fig. 12.3 Rotavirus mortality in children younger than 5 years. Reprinted with permission from Tate et al. (2012). Copyright © 2012, Elsevier.

Fig. 12.4 Mechanisms of RV-induced diarrhea. On the one hand, hyperosmotic diarrhea is generated by RV-induced death of infected enterocytes, which cause villi atrophy. Also, NSP4 increases intracytoplasmic Ca²+ and alters the cytoskeleton and the expression of disaccharidases. On the other hand, secretory diarrhea is generated by the release of 5HT from enterochromaffin cells (EC), which enhances secretion of Cl− and H2O regulated by the central nervous system.

Fig. 12.5 Effects of RV vaccination in children younger than 5 years of age. Median % reduction in RV hospital admission, all RV illness including any visit to a healthcare provider (outpatient and emergency department visits, and hospital admissions), acute gastroenteritis hospital admissions, and acute gastroenteritis deaths, 1–2 years after introduction of RV vaccination (data from 10 countries). Error bars show minimum and maximum values. n = number of studies used to obtain data. Reprinted with permission from Patel et al. (2012). Copyright © 2012, Elsevier.

Chapter 13

Fig. 13.1 (a) Evolutionary tree showing the proposed appearance of an ancestral papillomavirus between the branch point leading to amphibians and reptiles. It is thought that virus/host co-evolution has occurred during speciation, and that this has led to the widespread distribution of papillomaviruses in organisms as diverse as snakes, birds, and mammals. (b) The HPV types found in humans fall into five genera, with the Alpha and the Beta/Gamma genera representing the largest groups. HPV types from the Alpha Genus are often classified as low-risk cutaneous, low-risk mucosal, or high-risk. The evolutionary tree is based on alignment of the E1, E2, L2, and L1 genes (Doorbar et al., 2012). (c) Percentage of cervical cancers that are causally attributed to infection with members of the Alpha genus. Members of the Alpha 9 and 7 species have been studied most thoroughly.

Fig. 13.2 (a) Typical genome organization of the high-risk Alpha, Mu, and Beta HPV genomes. While all share a common genetic organization, the size and position of the major ORFs can vary, with Beta HPV types lacking an E5 ORF. The positions of the major promoters are marked with arrows on the high-risk Alpha HPV genome map, with early and late polyadenylation sites marked as PAL (late) and PAE (early). (b) Electron Micrograph of negatively stained papillomavirus particles. Individual capsomeres within the capsid structure can just be visualized. Papillomavirus particles are approximately 55nm diameter and are non-enveloped.

Fig. 13.3 (a) The E6 and E7 proteins of the high and low-risk HPV types have different functions that reflect their different biologies. The ability of the high-risk HPV types to drive cell division in neoplasia is thought to reflect the ability of their E7 protein to bind and degrade multiple members of pRb protein family, as well as the ability of E6 to efficiently degrade p53 and to compromise the function of PDZ-domain proteins that regulate cell contact and signalling pathways. (b) High-Risk HPV infection can lead to a silent or asymptomatic infection in which viral genomes persist in the basal layer without the development of disease, or alternatively to the development of a productive lesion such as CIN1, in which viral gene expression is regulated as the infected cells differentiate. In some instances, and at some epithelial sites, infection can lead to higher-grade neoplasia, with deregulated viral gene expression leading to secondary genetic changes in the host cell and possible integration of the viral genome into the cellular chromosome. The deregulated gene expression seen in CIN2 and 3, which are considered to be precancerous lesions, predisposes to the development of cancer.

Fig. 13.4 Lesion formation is thought to be facilitated by the presence of microwounds, which allows the virus to infect epithelial basal cells (e.g., an epithelial stem cell (1)). At particular sites, such as the SCJ of the cervical TZ, basal cells, reserve cells, and stem-like/stem cells are close to the epithelial surface and may be more prone to infection. At other sites, the development of a lesion may be facilitated by wound repair (2). Once a lesion has become established, basal and parabasal epithelial cells can be driven into the cell cycle, either to mediate basal cell division (i.e., cell proliferation) or to drive cell cycle re-entry (but not mitosis) in the upper epithelial layers in order to support viral genome amplification (3). Clearance of disease involves activation of a cell-mediated immune response and a suppression of viral gene expression as activated T cells accumulate in the vicinity of the lesion. It is thought that viral genomes can persist in the basal epithelial cells with very limited gene expression, allowing possible reactivation under some circumstances, such as can occur following immunosuppression (Maglennon et al., 2014).

Fig. 13.5 High-risk HPV infection disrupts the molecular pathways that regulate epithelial differentiation and cell proliferation. Cell cycle progression is regulated in the different epithelial layers by members of the pRb (Retinoblastoma) family of proteins. The E7 proteins of high-risk HPV types can target members of this protein family for degradation (shown in b). This releases members of the E2F transcription factor family, which allows basal and parabasal cells to enter S-phase. In uninfected epithelium (shown in a), the release of E2F is dependent on external growth factors, which stimulate cyclin D/cyclin-dependent kinase activity to allow pRb phosphorylation and E2F release. The expression of cellular proteins involved in cell cycle progression is regulated by p16, which is involved in a negative feedback loop by suppressing the activity of the cyclin D/cdk. The inability of low-risk HPV types, to drive robust basal cell proliferation, is thought to be because these types can only efficiently target the p130 Rb family member, which controls suprabasal, but not basal cell cycle entry. The high-risk E7 proteins are thought to target all members of the pRB family.

Chapter 14

Fig. 14.1 General processes of the KSHV infectious cycle and supporting factors which lead to pathogenesis. Certain risk behaviors are shown to act as catalysts for initiating a successful infection. Following primary infection is the persistent and potentially long-term state of latency, in which the virus exercises tactics to evade host immune responses. Any of the numerous stressors are presumed to act as a trigger causing the virus' switch from latent to lytic infection in a process called reactivation. It is during lytic replication that mature virions are generated. Subsequently, virus egress occurs, in which virus is spread to other cells and/or individuals, and this ongoing cycle repeats. Images courtesy of Microsoft Office Powerpoint via Bing.com.

Chapter 15

Fig. 15.1 Electron micrograph of Encephalitozoon hellem spores in a conjunctival biopsy of a patient with ocular microsporidiosis. Arrows point to polar tubes seen in cross-section.

Fig. 15.2In vitro culture of Encephalitozoon hellem in rabbit kidney (RK13) cells.

Fig. 15.3 Intestinal microsporidiosis. (a) Biopsy from a patient with Enterocytozoon bieneusi. Spores (arrow) are found on the apical side of the enterocyte nucleus. Section stained with Toluidine blue. (b) Biopsy from a patient with Encephalitozoon intestinalis. Spores (arrows) are found on the apical and basal side of enterocyte nuclei as well as in the lamina propria. Section stained with Toluidine blue.

Fig. 15.4 Microsporidia in a stool sample stained with modified Chromotrope 2R stain. Stool sample is from a patient with documented infection with Enterocytozoon bieneusi. Spores stain red and can display a clothes pin-like staining pattern.

Fig. 15.5 Ocular microsporidiosis. (a) Patient with punctuate keratoconjunctivitis due to Encephalitozoon hellem. Arrow points to punctuate ulcers demonstrated by fluorescein staining. (b) Conjunctival smear stained with Gram stain. Microsporidian spores (arrows) are gram positive with this stain. (c) Conjunctival smear stained with calcofluor white. Microsporidian spores (arrows) are fluorescent with this stain.

Fig. 15.6 Myositis due to microsporidiosis. Muscle biopsy from a patient with Anncaliia algerae infection. Microsporidian spores (arrow) and proliferating forms (arrow) are present within muscle fibers undergoing myolysis. A surrounding inflammatory infiltrate is also present.

Chapter 16

Fig. 16.1 Life cycle of Toxoplasma gondii.

Fig. 16.2 Oocysts of Toxoplasma gondii. (a) Unsporulated oocyst in a fecal flotation. Note the oocyst wall (arrow). Bar = 10 μm for a and b. (b) Flotation demonstrating a sporulated oocyst containing two sporocysts (arrowhead) each with four sporozoites (s). (c) Transmission electron micrograph of a sporulated oocyst. Four sporozoites are in each sporocyst. Arrows indicate junctions in the sporocyst wall, which collapse during excystation. The posteriorly located nucleus (N) of a sporozoite is visible. Bar = 1 μm.

Fig. 16.3 Tachyzoites, schizonts, and sexual stages in hematoxylin and eosin stained tissue sections. (a) Liver of a mouse containing many groups of tachyzoites (arrows). One group of tachyzoites (arrowhead) is in an endothelial cell which contains densely packed tachyzoites. Bar = 10 μm. (b) Section of ileum from a cat demonstrating schizonts (arrows) inside enterocytes. (c) Section of ileum from a cat demonstrating macrogamonts (arrows) inside enterocytes. Bar = 10 μm.

Fig. 16.4 Tissue cysts of Toxoplasma gondii containing bradyzoites in brain tissue. (a) Seven tissue cysts containing bradyzoites are present in this unstained brain smear from a mouse. Bar = 20 μm. (b) Transmission electron micrograph of a tissue cyst of the VEG strain in a neuron (N) from a mouse. A tissue cyst wall (arrow) surrounds hundreds of bradyzoites. Bar = 1 μm.

Fig. 16.5 Young tissue cyst demonstrating bradyzoites (B) and tachyzoites (T) which are enclosed by the tissue cyst wall (arrow).

Chapter 17

Fig. 17.1 Trypanosome morphology. This illustration shows fundamental features of a trypanosome trypomastigote. The flagellum exits the cell at the flagellar pocket and is attached lengthwise along the cell body. The nucleus contains the nuclear genome while the kinetoplast contains the mitochondrial genome.

Fig. 17.2Trypanosoma brucei lifecycle. The lifecycle starts when a tsetse fly injects metacyclic trypomastigotes into the host bloodstream. The parasite transforms in a long slender trypomastigote that multiplies via binary fission. The long slender trypomastigote then differentiates into a non-proliferating short stumpy trypomastigote form. When these are ingested by the insect vector they differentiate into the procyclic trypomastigote form that colonizes in the midgut of the tsetse fly. After migration to the salivary glands they differentiate into epimastigotes, and finally to the metacyclic trypomastigote form.

Fig. 17.3 Simulation of antigenic variation during T. brucei infection. When mammals are infected with T. brucei, subsequent parasitemic waves are formed by new variable antigenic types and host antibody responses.

Fig. 17.4 VSG switching can occur by four mechanisms. Telomere exchange, duplicative gene conversion, and partial gene conversion are recombination events responsible for switch in VSG. The genetic material is exchanged but transcription occurs from the original promotor. In contrast, ES switch results in the switch of all ES-associated genes and occurs from a different promotor. Figure inspired by Gloria Rudenko.

Fig. 17.5 Distribution of HAT on the African continent. T. b. gambiense, depicted in grey, occurs primarily in West and Central Africa and is responsible for 98% of HAT infections. T. b. rhodesiense infection, depicted in black, occurs in East Africa. Based on Report of a WHO meeting on elimination of African trypanosomiasis 2012.

Fig. 17.6 Reported HAT cases 1945–2012. Based on Report of a WHO Meeting on Elimination of African Trypanosomiasis 2012.

Fig. 17.7 Overview of immunopathology during murine T. brucei infection. Upon injection of the parasite in the bloodstream, pathogen-associated molecular patterns such as VSG and GIP lead to macrophage (MΦ) activation. These MΦ are further activated by IFN-γ, produced by a yet unidentified source. Activated macrophages produce TNF, NO, IL-1, and IL-6 amongst others, inducing a proinflammatory environment essential for parasite killing but also responsible for disease-associated pathology. IL-10, supposedly produced by lymphocytes and macrophages, partly dampens this pro-inflammatory environment.

Fig. 17.8 HAT treatment options. Pentamidine (Lomidine™, Aventis) is used for the treatment of early-stage T. b. gambiense infection. Eflornithine or DFMO (Ornidyl®, Aventis) is often used in combination with Nifurtimox for the treatment of late-stage T. b. gambiense. Suramin (Germanin®, Bayer) is used for early-stage T. b. rhodesiense infection and the highly toxic drug Melarsoprol (Arsobal®, Aventis) is used for the treatment of late-stage T. b. rhodesiense infection.

Chapter 19

Fig. 19.1

Fig. 19.2

Fig. 19.3

Chapter 20

Fig. 20.1Cryptosporidium spp. life cycle.

Fig. 20.2 Transmission electron micrograph showing an intracellular Cryptosporidium trophozoite inside their parasitophorous vacuole.

Fig. 20.3 Ileocecal neoplastic lesions in a dexamethasone-treated SCID mouse infected with C. parvum. Numerous parasites (arrows) inside the glands. Bar = 20 mm (hematoxylin and eosin). This photomicrograph was kindly provided by Dr. Colette Creusy from the Service d'Anatomie et de Cytologie Pathologiques, Groupe Hospitalier de l'Université Catholique de Lille, France.

Fig. 20.4 Microscopic observation of oocyts of Cryptosporidium. (a) Differential interference contrast image of oocysts recovered from stools using immunomagnetic separation (IMS). (b) Oocyts labeled by immunofluorescence (fluorescein) using a monoclonal antibody-based assay after recovering from stools using IMS. (c) Auramine staining of Cryptosporidium sp. oocysts. This photomicrograph was kindly provided by Dr. Emilie Frealle et Dr Yohann Le Govic from Parasitologie-Mycologie, Centre Hospitalier Régional et Universitaire de Lille, Univ Lille Nord de France. (d) Modified Ziehl–Neelsen staining of Cryptosporidium sp. oocysts in a fecal smear.

Chapter 21

Fig. 21.1 Life cycle of Plasmodium falciparum. P. falciparum has two developmental stages: the sexual stage in the Anopheles mosquito vector and the asexual stage in the human host. The infection in humans begins when sporozoites are injected during a blood meal, where they subsequently travel to the liver and replicate in hepatocytes. The sporozoites differentiate into merozoites, which are released into the blood circulation where they invade and replicate within host erythrocytes. Some of the blood-stage parasites may develop into gametocytes that could be taken up by an Anopheles mosquito during feeding. In the mosquito, the gametocytes form male and female gametes, which then fuse to form zygotes. These differentiate into ookinetes in the mosquito midgut, mature into oocysts, and eventually form sporozoites that migrate to the mosquito salivary glands. Injection of sporozoites to a human host during a blood meal completes the life cycle of P. falciparum. Reprinted with permission from Macmillan Publishers Ltd: NATURE GENETICS (Paslov, G., Protective hemoglobinopathies and Plasmodium falciparum transmission, 42(4): 284–5), Copyright (2010).

Fig. 21.2 Mechanism of erythrocyte invasion by P. falciparum merozoites. Erythrocyte invasion is a four-step process: (a) adhesion of a merozoite to a host erythrocyte; (b) reorientation of the merozoite to allow its apex to contact the erythrocyte and the release of organelles to form a tight junction at the area of contact; (c) and (d) ingress of the merozoite into a host erythrocyte using an actin-myosin motor, while shedding its surface coat; (e) closure of the tight junction to fully encase the merozoite within a parasitophorous vacuole. Reprinted from Cell, 124 (4), Cowman, A. F. and Crabb, B. S., Invasion of red blood cells by malaria parasites, 755–66, Copyright (2006), with permission from Elsevier.

Fig. 21.3 Cerebral malaria. Histological section of a brain of a patient who died of cerebral malaria showing the accumulation of P. falciparum-infected erythrocytes within a cerebral microcapillary. (An infected erythrocyte is indicated by the black arrow.) Reprinted from Acta Tropica, 121 (3), Wernsdorfer, W. H., Global challenges of changing epidemiological patterns of malaria, 158–65, Copyright (2012), with permission from Elsevier.

Fig. 21.4 Placental malaria and low birth weight. Placental histology section showing the sequestration of P. falciparum-infected erythrocytes (one is indicated by the arrow) in the placental intervillous spaces (left diagram) * indicates partially damaged syncytiotrophoblast, the epithelial cell layer covering placental villous tree and separating maternal and foetal blood. A child with low birth weight born to an African woman with placental malaria (right diagram). Reprinted from The Lancet Infectious Diseases, 7 (2), Desai, M., ter Kuile, F.O., Nosten, F., McGready, R., Asamoa, K., Brabin, B. and Newman, R.D., Epidemiology and burden of malaria in pregnancy, 93–104, Copyright (2007), with permission from Elsevier.

Fig. 21.5 Life cycle of P. vivax. P. vivax life cycle is largely similar to that of P. falciparum. Uniquely, some of the sporozoites in the liver form hypnozoites that can persist to cause latent P. vivax infections. Another feature unique to P. vivax is that the merozoites preferentially invade reticulocytes. Reprinted from The Lancet Infectious Diseases, 9 (9), Mueller, I., Galinski, M.R., Baird, J.K., Carlton, J.M., Kochar, D.K., Alonso, P.L. and del Portillo, H.A., Key gaps in the knowledge of Plasmodium vivax, a neglected human malaria parasite, 555–66, Copyright (2009), with permission from Elsevier.

Fig. 21.6 Current strategies employed in malaria prevention and control.

Chapter 22

Fig. 22.1 Schematic three-dimensional representation of T. vaginalis showing an external view (left) and the internal cell structures (right). The undulating membrane, which is formed by a fold in the plasma membrane, is observed in contact with the recurrent flagellum (RF). AF, anterior flagella; Ax, axostyle; bb, basal bodies; C, costa; G, Golgi complex; H, hydrogenosomes; Pe, pelta; N, nucleus; Nu, nucleolus; PF, parabasal filament; Sg, sigmoidal filaments; V, vacuole.

Fig. 22.2 Routine preparations of T. vaginalis for scanning (a) and transmission electron microscopy (b). Panel (a) shows a general view, with the anterior flagella (AF), the recurrent flagellum (RF) forming the undulating membrane, and the axostyle (Ax) forming the posterior tip of the cell. Panel (b) shows the cell interior. A single anterior nucleus (N), the axostyle (Ax), the flagella (F), the Golgi complex (G), hydrogenosomes (H), the endoplasmic reticulum (ER), and an endocytic vacuole (EV) containing ingested material can be seen. Scale bars: (a) 1 μm; (b) 500 nm.

Fig. 22.3 Metabolic pyruvate metabolism in T. vaginalis hydrogenosomes. The process of pyruvate metabolism starts in the cytosol, where glycolysis generates intermediate products, such as pyruvate and malate, that enter the hydrogenosome. Pyruvate is oxidatively decarboxylated, and the electrons are transferred to protons, with the formation of H2. In T. vaginalis, oxidative pyruvate decarboxylation is catalyzed by a different enzyme than in other eukaryotic cells; this enzyme is pyruvate:ferredoxin oxidoreductase, an enzyme that is found in a number of bacteria and other microorganisms.

Fig. 22.4T. vaginalis in the process of cellular division. The spindle (S) is extranuclear and originates from the attractophore (A), which is a microtubule-organizing center in trichomonads. The attractophore is localized under the basal bodies (BB), next to the origin of the parabasal filaments (PB). ER, endoplasmic reticulum; L, lysosome; H, hydrogenosomes; N, nucleus. Scale bar: 500 nm.

Fig. 22.5 SEM of T. vaginalis (T) adhered to vaginal epithelial cells. (a) The epithelial monolayer exhibits signs of injury (arrows), such as retraction from neighboring cells. (b)T. vaginalis, which is normally pear-shaped, flattens itself after attaching to vaginal epithelial cells, maximizing the surface area between the parasite and the host cell. Doderlein's lactobacilli (B) can also be seen. AF, anterior flagella; RF, recurrent flagellum. Scale bars: (a) 10 μm; (b) 4 μm.

Fig. 22.6 Scanning electron microscopy of a 1-h interaction between a virulent T. vaginalis (T) strain and a confluent monolayer of oviduct epithelial cells (E). The epithelial cells are being pulled up by T. vaginalis, which causes mechanical damage to the host cells. (b) The parasite can be seen phagocytozing a yeast cell (Y) via sinking engulfment. AF, anterior flagella; RF, recurrent flagellum. Scale bar: 2 μm.

Chapter 23

Fig. 23.1 Map of distribution for Loa loa in Africa.

Fig. 23.2 Life cycle of Loa loa.

Fig. 23.3Chrysop silacea, one vector of Loa loa.

Fig. 23.4 Microfilariae isolated from hypermicrofilaremic patient (microfilariae > 30,000/mL), cause of encephalitis in loiasis.

Fig. 23.5 Spectra of infection in loiasis showing the prevalence of different infections status.

Fig. 23.6 Influence of Loa loa on immune response against other antigens. Mitogen Concanavalin A (ConA) and Mycobacterium tuberculosis PPD immune response in a high (High T) or low transmission (Low T) area for L. loa.

Fig. 23.7 Potential mechanism for blocking allergic reactions by IgG4 and IgE in loiasis. Cells may be activated by immunoreceptor tyrosine-based motif (ITAM) by crosslinking of IgE/IgE (via L. loa antigens) fixed on Fcϵ RI receptor from the cells surface and induce allergy. Or, inhibition of cells' activation by immunoreceptor tyrosine-based inhibitory motif (ITIM) followed crosslinkage between IgE/IgG4 (via L. loa antigens) and fixation on FcϵRI (IgE) and FcγRII by IgG4, resulting in the absence of allergic reaction.

Fig. 23.8 Immunoprecipitation of ¹²⁵I-labeled Loa loa adult antigen with endemic sera. Immune complexes formed by L. loa antigen and loiasis sera were separated on 12% SDS-PAGE, followed by autoradiography. Lanes 1 and 2: high- microfilaraemic individuals; lanes 3–7: low- microfilaraemic individuals; lanes 8–12: amicrofilaraemic individuals; lane 13: negative human control. Molecular weight standards are shown in kDa.

Fig. 23.9 Effect of N-glycanase on Loa loa and Brugia pahangi antigens. Both are ¹²⁵I surface-labeled extracts treated with 10 U/mL (lanes 3 and 4) and 100 U/mL (lanes 1 and 2) of N-glycanase, run side by side with non-treated Loa loa antigen (lane 5) and Brugia pahangi (lane 6). Arrows on both sides indicate the shift in molecular weight of the 30–31 kDa antigen.

Fig. 23.10 Concomitant evolution of immune marker (IgG4) and appearance of microfilaria. Concomitant evolution of immune marker (IgG4) and microfilaria appearance in the blood of individuals exposed to L. loa, according to their age.

Fig. 23.11Loa loa microfilaria on direct examination by microscopy.

Chapter 24

Fig. 24.1 Toxocariasis. (a) Life cycle of T. canis supplied by the Centers for Disease Control and Prevention (Blaine Mathison). (b) Infective T. cati egg recovered from a child's sandbox. Courtesy of Kevin Kazacos. (c) Eosinophilic granuloma in a child's liver, with larva (arrow). Courtesy of Kevin Kazacos, contributed by M.D. Little, PhD, Tulane University. (d) Intense eosinophilic inflammation in the liver of a child with VLM, with associated larva (arrow). Courtesy of Kevin Kazacos, contributed by M.D. Little, PhD, Tulane University. (e) Larva migrating in dog retina, with larval sections. Courtesy of Kevin Kazacos, contributed by R.R. Dubielzig, DVM, PhD, University of Wisconsin.

Fig. 24.2 Toxocariasis. (a) Enucleated human eye with Toxocara endophthalmitis, showing a large retrolental mass with a funnel-shaped retinal detachment (AFIP Neg. 53-18355). (b) Toxocaral larval granuloma in a dog's eye, showing larval sections. Courtesy of Kevin Kazacos, contributed by R.R. Dubielzig, DVM, PhD, University of Wisconsin.

Fig. 24.3 Baylisascariasis. (a) Life cycle of B. procyonis supplied by the Centers for Disease Control and Prevention (Blaine Mathison). (b) B. procyonis adults from the intestine of a raccoon in Indiana. (c) B. procyonis larva from brain, showing anterior end, lips, and esophagus. Courtesy of Kevin Kazacos. (d) Lung from a child with fatal B. procyonis infection in Illinois (Fox et al., 1985) showing numerous larval granulomas. Courtesy of N.S. Gould, MD, Chicago, IL). (e) Section of granuloma from the lung demonstrating larval cross sections. Courtesy of Kevin Kazacos, contributed by N.S. Gould, MD, Chicago, IL, and adaptd with permission from Kazacos et al. (2013b). Copyright © 2014, Elsevier.

Fig. 24.4 Baylisascariasis. (a) T2-weighted MRI of the brain of a child with B. procyonis encephalitis (Park et al., 2000), showing marked T2 hyperintensity throughout the central white matter (from Park et al., 2000, with permission). (b) Brain of a child with fatal B. procyonis infection in Illinois (Fox et al., 1985), showing severe periventricular necrosis (Courtesy of N.S. Gould, MD, Chicago, IL). (c) Gliotic granuloma in the brain of a child with fatal B. procyonis infection in Pennsylvania (Huff et al., 1984) (Courtesy of Kevin Kazacos, contributed by D.S. Huff, MD, Philadelphia, PA). (d) Cross section of a larva in the brain of a child with fatal infection in Illinois (Fox et al., 1985). Note prominent, single lateral alae, large laterally compressed intestine, and smaller, paired triangular excretory columns (Courtesy of N. S. Gould, MD, Chicago, IL, and adapted with permission from Kazacos et al. (2013b)).

Fig. 24.5 Baylisascariasis. (a) Funduscopic photograph of the right eye of a teenage girl with diffuse unilateral subacute neuroretinitis due to B. procyonis in New York City, showing coiled subretinal nematode in inferior macula (arrow), arteriole attenuation, pigment epithelium mottling, and optic nerve pallor. (b) Magnified view of living, uncoiled, motile larva in the right eye one week after photocoagulation and (c) two weeks later (Courtesy of N. A. Saffra, MD, New York, NY, and adapted with permission from Kazacos et al. (2013b).

Fig. 24.6 Gnathostomiasis. (a) Life cycle of the parasite adapted from figure supplied by the Centers for Disease Control and Prevention (Blaine Mathison). (b) Creeping eruption due to Gnathostoma (from Chai et al. (2003), with permission of the journal). (c) Intraocular gnathostomiasis (from Pillai (2012), with permission of the journal).

Fig. 24.7 Cutaneous larva migrans. (a) Life cycle supplied by the Centers for Disease Control and Prevention (Blaine Mathison). (b) Foot of a patient who vacationed in Jamaica and returned with CLM (Courtesy of Christina M. Coyle and Herbert B. Tanowitz). (c) Lesions on the buttocks of a child and associated excoriations. (d) Foot of a child with CLM (Figures (c) and (d) from Herman Zaiman's A Pictorial Presentation of Parasites, with permission from the American Society of Topical Medicine and Hygiene).

Chapter 25

Fig. 25.1 Global distribution map of six important human schistosome species. Artwork by Greg Galin.

Fig. 25.2 General life cycle diagram for schistosomes infecting man. Artwork by Greg Galin.

Fig. 25.3 Egg size and morphology for six important human schistosome species. Artwork by Greg Galin.

Fig. 25.4 Isolated adult female Schistosoma haematobium worm.

Fig. 25.5  Deposition of microinjected Schistosoma haematobium eggs in a mouse bladder wall seven days post-infection. Note the presence of miracidia inside the eggs.

Fig. 25.6  Microultrasounds depicting the growth of granuloma in a mouse bladder wall following microinjection with a bolus of Schistosoma haematobium eggs. (a) Day 1 post-injection. (b) Day 6 post-injection. (c) Day 21 post-injection. (d) Day 36 post-injection.

Fig. 25.7  Extensive squamous metaplasia resulting from chronic human infection with Schistosoma haematobium.

Fig. 25.8  Well-differentiated squamous cell carcinoma following chronic human infection with Schistosoma haematobium.

Fig. 25.9  Calcification of Schistosoma haematobium eggs within a mouse bladder wall 35 days after microinjection.

Fig. 25.10  Moderately differentiated squamous cell carcinoma with a large number of dead or degenerated Schistosoma haematobium eggs in a human bladder.

Chapter 26

Fig. 26.1Leishmania infantum amastigotes infesting canine macrophage cells.

Fig. 26.2 Dog infected by Leishmania infantum displaying clinical signs (e.g., periocular alopecia) of infection.

Fig. 26.3 Microfilaria of Onchocerca lupi collected at skin snipping.

Fig. 26.4 Conjunctivitis and ocular discharge in a dog with heavy ocular infection by Thelazia callipaeda.

Fig. 26.5Phortica variegata feeding on lacrimal secretion of a human.

List of Contributors

Shaw M. Akula

Department of Microbiology and Immunology, Brody School of Medicine at East Carolina University, Greenville, NC, USA

Zeal T. Akula

Department of Microbiology and Immunology, Brody School of Medicine at East Carolina University, Greenville, NC, USA

Fatih Anfasa

Department of Viroscience, Erasmus Medical Center, Rotterdam, The Netherlands

Department of Internal Medicine, Faculty of Medicine, Universitas Indonesia, Jakarta, Indonesia

Juana Angel

Instituto de Genética Humana, Facultad de Medicina, Pontificia Universidad Javeriana, Bogotá, Colombia

Olympia Apostolopoulou

Department of Critical Care Medicine, Medical School, University of Athens, Chaidari-Athens, Greece

George Arabatzis

Mycology Research Laboratory, Department of Microbiology, Medical School, National and Kapodistrian University, Athens, Greece

Alicia I. Arechavala

Mycology Unit, Francisco J. Muniz Hospital, Buenos Aires City, Argentine Republic

Ricardo Ataíde

Burnet Institute, Center for Biomedical Research, Melbourne, Victoria Australia

Lucilla Baldassarri

Department of Infectious, Parasitic and Immune-Mediated Diseases, Istituto Superiore di Sanità, Rome, Italy

Monique Barel

Université Paris Descartes, Sorbonne Paris Cité, Bâtiment Leriche, Paris, France

INSERM, U1151, Unité de Pathogénie des Infections Systémiques, Paris, France

Alfonso Barreto

Departamento de Microbiología, Facultad de Ciencias, Pontificia Universidad Javeriana, Bogotá, Colombia

Daniela Basso

Department of Medicine-DIMED, University of Padova, Padova, Italy

Sadia Benamrouz

Biologie et Diversité des Pathogènes Eucaryotes Emergents (BDEEP), Centre d'Infection et d'Immunité de Lille (CIIL), Institut Pasteur de Lille, INSERM U1019, CNRS UMR 8402, Université de Lille, France

Ecologie et Biodiversité, Faculté Libre des Sciences et Technologies de Lille, Université Catholique de Lille, France

Marlene Benchimol

Universidade do Grande Rio, UNIGRANRIO, Rio de Janeiro, Brazil

Centro Nacional de Biologia Estrutural e Bioimagem (CENABIO), Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil

Instituto Nacional de Metrologia, Qualidade e Tecnologia – Inmetro, Duque de Caxias, Rio de Janeiro, Brazil

Mark Eric Benbow

Department of Entomology and Department of Osteopathic Medical Specialties, Michigan State University, MI, USA

Carlos Fernández Benítez

Unidad de Gestión Clínica Centro de Salud de Laviana, Asturias, Spain

Alberto Berardi

Neonatal Intensive Care Unit, Polyclinic University Hospital, Modena, Italy

Philippe Boeuf

Burnet Institute, Center for Biomedical Research, Melbourne, Victoria Australia

José Antonio Boga

Servicio de Microbiología, Hospital Universitario Central de Asturias, Oviedo, Spain

Michael S. Bronze

Department of Internal Medicine, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA

Andreas Burkovski

Department Biologie, Friedrich-Alexander-Universität Erlangen-Nürnberg, Erlangen, Germany

Gabriela Certad

Biologie et Diversité des Pathogènes Eucaryotes Emergents (BDEEP), Centre d'Infection et d'Immunité de Lille (CIIL), Institut Pasteur de Lille, INSERM U1019, CNRS UMR 8402, Université de Lille, France

Alain Charbit

Université Paris Descartes, Sorbonne Paris Cité, Bâtiment Leriche, Paris, France

INSERM, U1151, Unité de Pathogénie des Infections Systémiques, Paris, France

Kaw Bing Chua

Temasek Life Sciences Laboratory, National University of Singapore, Singapore

Jennifer Cnops

Laboratory for Cellular and Molecular Immunology, Vrije Universiteit Brussel, Brussels, Belgium

Department of Structural Biology, VIB, Brussels, Belgium

Alexandra Correia

Instituto de Biologia Molecular e Celular, Universidade do Porto, Porto, Portugal

Instituto de Ciências Biomédicas de Abel Salazar, Universidade do Porto, Porto, Portugal

Christina M. Coyle

Departments of Medicine, Jacobi Medical Center and the Montefiore Medical Center, Albert Einstein College of Medicine, Bronx, NY, USA

Roberta Creti

Department of Infectious, Parasitic and Immune-Mediated Diseases, Istituto Superiore di Sanità, Rome, Italy

Filipe Dantas-Torres

Aggeu Magalhães Research Centre, Oswaldo Cruz Foundation, Recife, Pernambuco, Brazil

Frank R. DeLeo

Laboratory of Human Bacterial Pathogenesis, Rocky Mountain Laboratories, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT, USA

Wanderley de Souza

Centro Nacional de Biologia Estrutural e Bioimagem (CENABIO), Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil

Instituto Nacional de Metrologia, Qualidade e Tecnologia – Inmetro, Duque de Caxias, Rio de Janeiro, Brazil

Laboratório de Ultraestrutura Celular Hertha Meyer, Universidade Federal do Rio de Janeiro, Brazil

Elizabeth S. Didier

Division of Microbiology, Tulane National Primate Research Center, Covington, LA, USA

Department of Tropical Medicine, School of Public Health and Tropical Medicine, Tulane University, New Orleans, LA, USA

George Dimopoulos

Department of Critical Care Medicine, Medical School, University of Athens, Chaidari-Athens, Greece

Esteban Domingo

Centro de Biología Molecular Severo Ochoa (CSIC-UAM), Consejo Superior de Investigaciones Científicas (CSIC), Campus de Cantoblanco, Madrid, Spain

John Doorbar

Department of Pathology, University of Cambridge, Cambridge, United Kingdom

Laurent Dortet

INSERM U914 Emerging Resistance to Antibiotics, Le Kremlin-Bicêtre, Paris, France

Douglas A. Drevets

Department of Internal Medicine, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA

Department of Veterans Affairs Medical Center, Oklahoma City, OK, USA

Taylor Eddens

Richard King Mellon Foundation Institute for Pediatric Research, Children's Hospital of Pittsburgh of UPMC, Pittsburgh, PA, USA

Peter Q. Eichacker

Critical Care Medicine Department, Clinical Center, National Institutes of Health, Bethesda, MD, USA

Brenda L. Fredericksen

Maryland Pathogen Research Institute, University of Maryland, College Park, MD, USA

Department of Cell Biology and Molecular Genetics, University of Maryland College Park, College Park, MD, USA

Chi-Ling Fu

Department of Urology, Stanford University School of Medicine, Stanford, CA, USA

Joaquim Gascón

Global Health Institute (ISGlobal), Hospital Clínic-Universitat de Barcelona, Barcelona, Spain

Thomas W. Geisbert

Department of Microbiology and Immunology, The University of Texas Medical Branch, Galveston, TX, USA

Giovanni Gherardi

Centro Integrato di Ricerche, Laboratory of Microbiology, University Campus Biomedico, Rome, Italy

Namraj Goire

Microbiology Department, PathWest Laboratory Medicine WA, Queen Elizabeth II Medical Centre, Nedlands, Western Australia, Australia

Queensland Paediatric Infectious Diseases Laboratory, Queensland Children's Medical Research Institute, Royal Children's Hospital, Brisbane, Queensland, Australia

Thaddeus G. Golos

Department of Comparative Biosciences, Wisconsin National Primate Research Center, Madison, WI, USA

Christopher R. Gourley

School of Molecular Biosciences, College of Veterinary Medicine, Washington State University, Pullman, WA, USA

Nathalie Grall

Laboratoire de Bactériologie, Hôpital Bichat-Claude Bernard, APHP, Paris, France

EA 3964 - Université Paris-Diderot, Paris, France

Luis Otero Guerra

Servicio de Microbiología, Hospital de Cabueñes, Gijón, Asturias, Spain

Belinda Hall

Department of Microbial and Cellular Sciences and School of Biosciences and Medicine, University of Surrey, Guildford, UK

John J. Halperin

Department of Neurosciences, Overlook Medical Center, Summit, NJ, USA

Sidney Kimmel Medical College of Thomas Jefferson University, Philadelphia, PA, USA

Olfat Hammam

Department of Urology, Stanford University School of Medicine, Stanford, CA, USA

Alistair Harrison

The Center for Microbial Pathogenesis, The Research Institute at Nationwide Children's Hospital, Columbus, OH, USA

Elizabeth L. Hartland

Department of Microbiology and Immunology, University of Melbourne, Victoria, Australia

Caitlin Hicks

Johns Hopkins Hospital, Department of Surgery, Baltimore, MD, USA

Michael Hsieh

Division of Urology, Children's National Health System, Washington, DC, USA

Departments of Urology and Pediatrics, The George Washington University, Washington, DC, USA

The Biomedical Research Institute, Rockville, MD, USA

Katherine L. Hussmann

Department of Cell Biology and Molecular Genetics, University of Maryland College Park, College Park, MD, USA

Tetsuro Ikegami

Department of Pathology, The University of Texas Medical Branch at Galveston, Galveston, TX, USA

J. Igor Iruretagoyena

Department of Obstetrics and Gynecology, University of Wisconsin-Madison, Madison, WI, USA

Cong Jin

National Institute for Viral Disease Control and Prevention, Chinese Center for Disease Control and Prevention, Beijing, China

Heather Williamson Jordan

Department of Biological Sciences, Mississippi State University, MS, USA

Aniket Kaloti

Richard King Mellon Foundation Institute for Pediatric Research, Children's Hospital of Pittsburgh of UPMC, Pittsburgh, PA, USA

Sophia Kathariou

Department of Food, Bioprocessing and Nutrition Sciences, North Carolina State University, Raleigh, NC, USA

Kevin R. Kazacos

Department of Comparative Pathobiology, Purdue University College of Veterinary Medicine, West Lafayette, IN, USA

Jay K. Kolls

Richard King Mellon Foundation Institute for Pediatric Research, Children's Hospital of Pittsburgh of UPMC, Pittsburgh, PA, USA

Michael E. Konkel

School of Molecular Biosciences, College of Veterinary Medicine, Washington State University, Pullman, WA, USA

Stefan Kunz

Institute of Microbiology, University Hospital Center and University of Lausanne, Lausanne, Switzerland

Monica M. Lahra

Neisseria Reference Laboratory and WHO Collaborating Centre for STD, Microbiology Department, South Eastern Area Laboratory Services, Prince of Wales Hospital, Sydney, New South Wales, Australia

Thien-Linh Le

Department of Urology, Stanford University School of Medicine, Stanford, CA, USA

Dexin Li

National Institute for Viral Disease Control and Prevention, Chinese Center for Disease Control and Prevention, Beijing, China

Caroline Lin Lin Chua

Taylor's University Lakeside Campus, School of Biosciences, Subang Jaya, Selangor, Malaysia

David S. Lindsay

Center for Molecular Medicine and Infectious Diseases, Department of Biological Sciences and Pathobiology, Virginia-Maryland Regional College of