Advances in Malaria Research E-Book

• Herausgeber: John Wiley & Sons
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 1558

Veröffentlichungsjahr: 2016

Beschreibung

* Thoroughly reviews our current understanding of malarial biology * Explores the subject with insights from post-genomic technologies * Looks broadly at the disease, vectors of infection, and treatment and prevention strategies * A timely publication with chapters written by global researchers leaders

Leseprobe

Table of Contents

Cover

Title Page

List of contributors

Foreword

Preface

CHAPTER 1: Introduction

History

The life cycle of

Plasmodium

A significant milestone in malaria research: Adaptation of

Plasmodium

to laboratory culture

The advent of present‐day technologies and their applications in malaria research

Bibliography

CHAPTER 2: Exoerythrocytic development of

Plasmodium

parasites

The sporozoite’s journey from the skin to the liver

Sporozoite invasion

Parasite development

Protein export from the parasite into the host cell

Parasite egress

The role of innate immune responses during merosome formation

Acknowledgments

Bibliography

CHAPTER 3: Molecular basis of erythrocyte invasion by

Plasmodium

merozoites

The structure of the merozoite

The steps of erythrocyte invasion

Redundancy and ligand–receptor interactions that mediate parasite adhesion during erythrocyte invasion

Signaling events during erythrocyte invasion by malaria parasites

Summary and conclusions

Bibliography

CHAPTER 4: The biology of malaria transmission

Purpose

History

The current research agenda

Biology

Population dynamics

Transmission‐blocking interventions

Bibliography

CHAPTER 5: Comparative and functional genomics of malaria parasites

An Introduction to

Plasmodium

genomes

Genome structure of malaria parasites

From genome sequences to gene function

Summary

Acknowledgments

Bibliography

CHAPTER 6: Gene regulation

Introduction

Modes of gene regulation

Drug targeting

Perspectives

Bibliography

CHAPTER 7: Molecular genetic approaches to malaria research

Transfection methods

Genetic approaches for deriving gene function

Conditional knockdown of protein function

Protein reporters

Conclusions

Bibliography

CHAPTER 8: Transcriptomics and proteomics

Transcriptional profiling throughout the parasite life cycle

Transcriptional regulation

Transcriptional variation

Biological insights

Proteomics

Translational regulation

Conclusion

Bibliography

CHAPTER 9: The biochemistry of

Plasmodium falciparum

MPMP

Carbohydrates

Lipid metabolism

Amino acids

Nucleotide metabolism

Cofactors

Redox metabolism

Mitochondrial functions

Hemoglobin digestion and hemozoin production

Some reflections for the future

Bibliography

CHAPTER 10: Signaling in malaria parasites

Protein phosphorylation in

Plasmodium

Calcium‐mediated signaling in

Plasmodium

Phosphoinositide signaling and trafficking in malaria parasites

Cyclic nucleotide signaling in the malaria parasite

Future challenges

Bibliography

CHAPTER 11: Membrane transport proteins as therapeutic targets in malaria

Host erythrocyte membrane: A shared ion and nutrient channel

Parasitophorous vacuolar membrane: Protein export and solute uptake

Parasite plasma membrane: Similar to other eukaryotic cells, but different

Digestive vacuole: A specialized lysosome‐equivalent in the parasite

Mitochondrial inner membrane: An unusual ATP synthase with uncertain function

Conclusions

Acknowledgments

Bibliography

CHAPTER 12: The proteolytic repertoire of malaria parasites

Aspartic proteases

Cysteine proteases

Metalloproteases

Serine proteases

Threonine proteases

Roles of proteases in parasite development

Summary and conclusions

Acknowledgments

Bibliography

CHAPTER 13: Development of medicines for the control and elimination of malaria

Targets for the development of future medicines for malaria

The process of drug development

Advances in drug development made in the 21st century

The global pipeline of new medicines for treating malaria

Medicines in the broader context of malaria eradication

Conclusion

Acknowledgments

Bibliography

CHAPTER 14: Antimalarial drug resistance

Background

Causes of drug resistance

Detection of drug resistance

Managing drug resistance

Conclusion

Disclosures

Bibliography

Further reading

CHAPTER 15: Epidemiology of

Plasmodium falciparum

malaria

Burden of disease

Geographical distribution of the disease

Chain of transmission and infection cycle

Malaria endemicity and malaria transmission

Malaria elimination and eradication

Bibliography

CHAPTER 16: Malaria pathogenesis

Malaria illness

Host susceptibility

Parasite virulence

Conclusions

Bibliography

CHAPTER 17: Host genetics

What evidence is there that the risk of malaria is genetically determined?

Identifying the genes involved

Why is genetic resistance important?

Genetic polymorphisms of the red blood cell

Hemoglobinopathies

G6PD deficiency

Non–red blood cell polymorphisms

Concluding remarks

Bibliography

CHAPTER 18: The immune response in mild and severe malaria

The picture of the acquisition of resistance to uncomplicated and severe malaria: Framing the questions

Searching for host genes that confer immune resistance to severe malaria

SLE susceptibility and resistance to severe malaria

The relationship between the pathogen environment and susceptibility to severe malaria

Summary

Acknowledgments

Bibliography

CHAPTER 19: Progress in development of malaria vaccines

Immunity to malaria

Life cycle of malaria parasites and points of intervention with vaccines

Pre‐erythrocytic stage vaccines

Blood‐stage vaccines

Transmission‐blocking vaccines

Live attenuated vaccines for malaria

Conclusion

Bibliography

CHAPTER 20:

Plasmodium vivax

Burden of

Plasmodium vivax

infection and disease

Severe

Plasmodium vivax

malaria

Molecular basis of severe disease

Concluding remarks and outstanding research questions

Bibliography

Index

End User License Agreement

List of Tables

Chapter 03

Table 3.1 Characteristic features of Invasion related proteins of

P. falciparum.

Chapter 05

Table 5.1 Genome data of some representative

Plasmodium

parasites.

Chapter 06

Table 6.1 Functional chromatin compartmentalization.

Table 6.2 Validated

Plasmodium

DNA‐binding proteins regulating transcription and/or chromatin organization.

Table 6.3 Predicted and verified histone mark writers in

Plasmodium falciparum.

Table 6.4 Predicted and verified histone mark readers in

P. falciparum.

Chapter 12

Table 12.1  Proteolytic repertoire of malaria parasites.

Chapter 13

Table 13.1 Combining different types of blood‐stage activities to produce a new medicine, with a summary of their advantages and disadvantages.

Table 13.2 New malaria medicines in development and their targets.

Chapter 14

Table 14.1 Time difference between the date of introduction and the first report of clinical resistance for selected drugs

Table 14.2 Studies associating increased

pfmdr1

copy number in

P. falciparum

malaria with clinical outcomes

Table 14.3 Antimalarial drugs and associated molecular markers of resistance by species

Chapter 15

Table 15.1 Estimated number of malaria cases and proportion to

P. falciparum

cases by WHO region (2010).

Table 15.2 Estimated number of malaria deaths and proportion of deaths in children younger than 5 years by WHO region (2010).

Table 15.3 Classification of malaria endemicity levels

Chapter 16

Table 16.1 Adjunctive therapies studied or considered for treatment of SM, and their impact on mortality.

Table 16.2 SM and adhesion to specific endothelial receptors.

Chapter 17

Table 17.1 Approaches to providing evidence for protective association between specific genetic factors and malaria.

Table 17.2 Examples of candidate malaria‐association genes relating to non–red blood cell products.

List of Illustrations

Chapter 01

Figure 1.1 Life cycle of the malaria parasite.

Chapter 02

Figure 2.1 Comparison of the two

Plasmodium

invasive forms in the mammalian host. On the left, a sporozoite is depicted, and on the right, a merozoite is shown. For the sporozoite, the composition of the membrane and underlying features, including the molecules responsible for gliding motility, are shown as a magnified image.

Figure 2.2 Exoerythrocytic development of

Plasmodium

parasites.

A,

On the left, a time scale of the events is shown. The cartoon shows the entire development from transmigration of the sporozoites to budding of merosomes. On the right, three TEM images are shown.

B,

Cross section of an invaded sporozoite.

C,

Developing schizont.

D,

Merosome containing many merozoites.

Figure 2.3

P. berghei

schizont develops close to the host cell Golgi. A schematic representation

(A),

a TEM image

(B),

and an IFA image

(C)

are shown. HCC: host cell cytoplasm; HCN: host cell nucleus, HG: host Golgi; PC: parasite cytoplasm; PN: parasite nucleus; PV: parasitophorous vacuole. For the IFA, HepG2 cells were infected with

P. berghei

sporozoites and fixed 16 hpi. Staining of the host‐cell Golgi was with an anti‐BIP antibody

(red)

and the parasite was stained with an anti‐PbICP antiserum

(green).

DNA was visualized with DAPI.

Figure 2.4 Organelle development in

P. berghei

liver schizonts.

A,

Scheme of a schizont with many nuclei

(blue),

one mitochondrion

(red),

and one apicoplast

(green). B,

TEM image showing a cross section of a schizont with mitochondria and apicoplast.

C

and

D,

Live imaging of early schizont

(C)

and late schizont

(D)

expressing a red fluorescent mitochondrion marker protein and a green fluorescent apicoplast marker protein.

Figure 2.5 Vesicle transport in the parasite and delivery into the host cell cytoplasm.

Upper panels,

The schemes are an overview

(A)

and a magnified area of this overview

(B)

highlighting the vesicle fusion with the parasite plasma membrane and the release of the content into the PV. In the

lower panels,

two TEM images are presented.

C

shows the vesicles accumulating at the plasma membrane.

D

shows the vesicles delivered to the host cell. HCC: host cell cytoplasm; PC: parasite cytoplasm; PN: parasite nucleus, PV: parasitophorous vacuole; asterisks mark parasite‐derived vesicles.

Figure 2.6 Merosome formation

in vivo

and

in vitro:

The scheme depicts the events surrounding merosome formation

in vivo. A,

Upon PVM rupture, the host cell detaches from the neighboring cells; vesicles (merosomes) bud off and are constantly filled with infectious merozoites.

B,

Intravital image of mCherry‐expressing parasites infecting an LC3‐GFP‐expressing transgenic mouse.

C,

TEM of a detached cell with a budding merosome.

D,

REM image of a merosome filled with many mature merozoites.

Chapter 03

Figure 3.1 Asexual blood‐stage life cycle of

Plasmodium.

Different stages of

P. falciparum

development in the erythrocyte are depicted. The merozoites attach the erythrocyte, apically reorient, form the junction, and invade erythrocytes. Within the infected erythrocyte, the parasite is present in the parasitophorous vacuole (PV), where it develops through the ring (0–24 hours), trophozoite (24–36 hours), and schizont stages (40–48 hours). During maturation, the erythrocyte surface gets remodeled with the formation of membrane‐bound structures (cleft and loop structures, Maurer’s clefts) in the erythrocyte cytoplasm and knobs on the erythrocyte membrane displaying the surface variant protein PfEMP1 (

P. falciparum

erythrocyte membrane protein 1). At around 48 hours, the infected erythrocyte undergoes egress, and 16 to 32 daughter merozoites are released into the blood stream, where they further invade nascent erythrocytes.

Figure 3.2 Three‐dimensional structure of the

P. falciparum

merozoite depicting its architecture and subcellular organelles. The merozoite is a pear‐shaped structure with an apical pole that has underlying polar rings. It possesses an outer surface coat that is fibrillar in nature and comprises a number of merozoite surface proteins. Internally, it is characterized by the presence of the apical organelles (rhoptry, microneme) that are a key feature of the apicomplexan organisms and are known to harbor most of the parasite proteins involved in host cell invasion (e.g., EBA, RH, AMA‐1). Dense granules are released immediately after invasion and harbor components of the protein translocation machinery that is inserted into the parasitophorous vacuole membrane. The merozoite possesses all the other subcellular organelles characteristically found in eukaryotes, such as the nucleus, mitochondria, endoplasmic reticulum, Golgi, microtubules, and ribosomes. The key structures that are unique to this eukaryotic pathogen are the presence of the apicoplast (a relict of the chloroplast), the subpellicular microtubules, and the inner membrane complex.

Figure 3.3 Steps of erythrocyte invasion by

P. falciparum

merozoites and potential strategies to block invasion. The different steps constituting the merozoite invasion process in erythrocytes are depicted. Merozoites undergo egress from an infected erythrocyte and are released into the blood stream. The calcium‐dependent protein kinase 5 (CDPK5) plays an important role in egress. Thereafter, the merozoite attaches to the erythrocyte membrane, apically reorients, and forms a junction. A number of DBL and RH proteins mediate attachment by binding with their respective erythrocyte receptors. AMA‐1 binds with RON2, which traverses the host red cell surface and associates with RON4/5 that are localized in the erythrocyte cytosol.

Figure 3.4 Structural features of the MSP family of proteins are depicted in the schematic diagram. All MSPs possess a signal peptide. MSP‐1/‐2/‐4/‐5/‐8/‐10 are GPI‐anchored, and all of them except MSP‐2 comprise EGF domains. MSP‐3 possesses alanine heptad repeats, a glutamate‐rich region, and a leucine‐zipper region. The closely related MSP‐6 possesses a glutamate‐rich region and a coiled‐coil region at its C‐terminal end. Pf92 is a GPI‐linked member of the 6‐cysteine protein family that is localized on the merozoite surface. Two DBL domain–containing MSPs have been identified: MSPDBL‐1 and MSPDBL‐2.

Figure 3.5

Processing of the MSP‐1/6/7 Complex.

Part I. Proteolytic processing of the MSP1/6/7 complex. Biosynthesis of all three MSPs begins at the start of schizogony (nuclear division). As indicated, the complex is a stoichiometric assembly of the various components, except that the MSP‐6 precursor binds as a tetramer (Kauth 2006). Primary processing converts it to MSP‐6

36

. The MSP‐7 precursor undergoes an early processing step during secretory transport (Pachebat 2007) to produce MSP‐7

33

, the form that binds to the MSP‐1 precursor. Primary processing converts MSP‐7

33

to both MSP‐7

22

and MSP‐7

19

. The various MSP‐1 processing products are numbered for clarity. Secondary processing, which occurs following egress and probably at the point of invasion, involves further cleavage within MSP‐1

42

to produce MSP‐1

33

and MSP‐1

19

. The latter is carried into the newly invaded erythrocyte on the merozoite surface (to form a so‐called ring‐stage parasite), whereas MSP‐1

33

is shed along with the remaining MSP‐1/‐6/‐7 complex. Secondary processing is mediated by a membrane‐bound protease called PfSUB2, whereas the protease(s) responsible for primary processing has been hitherto unknown (indicated by a question mark). Part II. Primary structure and processing of

P. falciparum

3D7 MSP‐1, MSP‐6, MSP‐7 and AMA‐1. SS, signal sequence; GA, GPI anchor; PS, pro‐sequence; TM, transmembrane domain. (A) Outline of the MSP‐1 precursor. The grey arrows indicate the sites of primary processing of the precursor protein into its major subunits MSP‐1

83

, MSP‐1

30

, MSP‐1

38

, and MSP‐1

42

(Stafford 1994, Koussis 2009). A secondary proteolytic cleavage mediated by PfSUB2 (black arrow) occurs during invasion, cleaving MSP‐1

42

into MSP‐1

33

and MSP‐1

19

. (B) AMA‐1 is synthesized as an 83 kDa precursor protein containing a C‐terminal transmembrane domain (TM). After targeting to the micronemes the N‐terminal pro‐sequence (PS) is removed, resulting in AMA‐1

66

, which appears at the merozoite surface at the time of schizont rupture. During invasion AMA‐1

66

is proteolytically cleaved by PfSUB2 (black arrow) resulting in release of AMA‐1

48/4

4

(Howell 2003). MSP‐6 (C) and MSP‐7 (D) are peripheral merozoite surface proteins, membrane‐bound through non‐covalent associations with MSP‐1. MSP‐6 is processed into MSP‐6

36

. MSP‐7 is initially cleaved into MSP‐7

33

(Pachebat 2007). Around the time of merozoite release from the newly ruptured schizont, MSP‐7

33

is further cleaved into MSP‐7

22

and MSP‐7

19

(Pachebat 2001, 2007).

Figure 3.6 Electron micrograph of merozoite invasion. Transmission electron microscopy image of a merozoite

(Mz)

contacting an erythrocyte

(E)

during the process of invasion. The apical organelles in the merozoite (rhoptry, microneme, and dense granules) are shown. The erythrocyte membrane is thickened (15 nm) at the attachment site marked by the arrow, which represents the junction (magnification × 54,000). The formation of the junction commits the merozoite to invade the specific erythrocyte.

Figure 3.7 Structure of apical membrane antigen 1. Polymorphic amino acids shown on the apical membrane antigen 1 (AMA1) crystal structure. Polymorphisms are based on sequence data from

P. falciparum

infections acquired at a vaccine testing site in Mali, West Africa.

A,

Polymorphic residues are numbered and highlighted.

Yellow

and

blue

residues are dimorphic,

orange

residues are trimorphic, and

red

residues have four to six possible amino acids. Residues highlighted in

green

and

blue

make up the hydrophobic pocket hypothesized to be a binding site between AMA1 and the rest of the erythrocyte invasion machinery, with

blue

indicating polymorphic residues within the pocket.

B,

Conserved residues in AMA1 domains I, II, and III are highlighted, respectively, in

light pink, light blue,

and

light orange.

Polymorphic residues in domain I are highlighted in

dark brown

(c1),

red

(c1 and c1L),

purple

(c2),

dark pink

(c3), and

light brown

(not incorporated in a cluster). Polymorphic residues in domains II and III are highlighted, respectively, in

dark blue

and

dark orange. Light gray

residues are not part of any of the three major domains, and

dark gray

residues are polymorphisms within the interdomain region.

Figure 3.8 Invasion motor complex of

P. falciparum

. A schematic diagram of the invasion motor complex depicting a motility model of the malaria parasite during erythrocyte invasion. The sub‐pellicular microtubules (top) are connected to the inner membrane complex (IMC). Myosin (MyoA) is anchored in the IMC through the MTIP‐GAP45‐GAP50 complex and walks on an actin treadmill that is formed by short actin filaments, which are polymerized at one end and depolymerized at the other. Actin is linked to the surface adhesins such as TRAP through a glycolytic enzyme, aldolase. The surface adhesins binds with the erythrocyte surface receptors (e.g., glycophorins, semaphorin‐7A) to form an anchor with the substrate host erythrocyte. Myosin moves across the actin treadmill toward the end of actin polymerization. In the process, force is generated that is transmitted via the adhesins to the substrate to propel the parasite forward. The surface adhesins are eventually cleaved by the rhomboid proteases, which results in disengagement of the ligand–receptor interactions between the parasite and host erythrocyte substrate.

Figure 3.9

EBA/PfRH Ligand‐receptor interactions involved in erythrocyte invasion by

P. falciparum

. Schematic representation of the interactions between the

P. falciparum

EBA and PfRH ligand proteins, with their corresponding erythrocyte receptors that mediate erythrocyte invasion. These molecular interactions are responsible for imparting redundancy to the erythrocyte invasion process. The erythrocyte receptors for some parasite ligands still remain unknown and are thus referred in literature by the letters “E”, “Y”, and “Z”. “?” refers to the unknown ligands and receptors that have yet not been discovered. PfRH5 is tethered by Ripr and CyRPA denoted as “R/C”.

Figure 3.10

The structural features of the EBA and PfRH family of proteins are depicted

. The erythrocyte‐binding domains of the ligand proteins are shown, which among the EBA proteins are known as Region II. The

lines

in PfRH2 and PfRH4 also represent regions that have been reported to bind erythrocytes. The pattern‐coded structural features of the proteins are shown in the key. PfRipr and RRMAP (CyRPA) form a multiprotein complex with PfRH5 that is the unique PfRH protein lacking a transmembrane domain. PfRipr comprises 10 EGF domains, and RRMAP (CyRPA) is GPI‐anchored.

Figure 3.11 Three‐dimensional structure of PfRH5 and basigin.

A,

Crystal structure of the PfRH5 protein.

B,

The structure of PfRH5 (depicted in

yellow

) bound to basigin (depicted in

blue

).

Figure 3.12 The PfRH5‐Ripr‐CyRPA multiprotein complex is essential for erythrocyte invasion. Whereas the PfRH5–basigin interaction is essential for

P. falciparum

erythrocyte invasion, PfRH5 lacks a transmembrane domain or a GPI‐anchor. It has been elucidated that PfRH5 exists on the surface of the invading merozoite as part of a multiprotein complex consisting of Ripr (RH5 interacting protein) and CyRPA (cysteine‐rich protective antigen). Ripr is a 125‐kDa protein that also lacks a transmembrane domain and is proteolytically processed into two equal fragments of ~65 kDa. CyRPA is the only GPI‐anchored protein among the three proteins and is thus the RH5‐Ripr membrane anchoring protein.

Chapter 04

Figure 4.1

In situ

hybridization to illustrate translational repression of a female specific messenger RNA.

1a

and

2a,

All parasites hybridize with a probe to the small subunit rRNA probe. r, ring stage asexual parasite; t, trophozoite; fg, female gametocyte; mg, male gametocyte.

1b,

Only the female gametocyte hybridizes with a Pbs28‐specific probe, revealing a punctate cytoplasmic distribution of the stored messenger RNA.

Figure 4.2 Sequential development of

Plasmodium

in the mosquito vector. 1 and 2, Male and female gametocyte ingested with blood meal (0–30 min); 3 and 4, male and female gametes (15–60 min); 5, zygote (15 min– ~9 h); 6, Ookinete burrows through the midgut epithelium (9–36 h); 7, Oocyst develops under the basal lamina of midgut, (days 1–25); 8, Rupture of oocyst, releasing sporozoites into the hemocele (days 9–21); 9, Salivary gland sporozoites (about day 10 until death of mosquito).

Figure 4.3 Scanning electron micrograph of

Plasmodium yoelii nigeriensis

microgametocyte undergoing the process of exflagellation, releasing the sometimes nucleated (N) male gametes.

Figure 4.4 Transmission electron micrograph of

Plasmodium yoelii nigeriensis

zygote illustrating a synaptonemal complex diagnostic of the leptotene and diplotene stages of meiosis.

Figure 4.5 Reconstruction of the apical complex of the

Plasmodium berghei

ookinete. p, plasma membrane; c, collar; ar, apical ring (the organizing center for the microtubular cytoskeleton); mt, subpellicular microtubules, pr, polar ring; imc, inner membrane complex (see Figure 4.6).

Figure 4.6 3D reconstruction of ookinete pellicle based upon cryofracture images of

Plasmodium gallinaceum.

EL1 and 2 external layers on the cell surface; PM‐ES, plasma membrane external surface; PM‐EF, plasma membrane external face; PM‐PF, plasmalemma protoplasmic face; OAM‐PF, outer alveolar membrane protoplasmic face; OAM‐EF outer alveolar membrane external face; mts, microtubules; su, suture between adjacent edges of the alveolar vacuole membrane. Note the 8‐fold symmetry of pores in the inner membrane complex.

Figure 4.7 Summary of known immune mechanisms used by the mosquito to attack malaria parasites. Ookinetes (a) can be killed by expulsion of the invaded midgut cell, by LRIM/APL1C/TEP‐mediated lysis (c) and phenol oxidase‐mediated melanization (d) in the basal labyrinth. Oocysts can be killed by NO (e). Melanization of oocysts to form Ross’s black spores is not illustrated.

Figure 4.8 Image of dissected midgut of

Anopheles stephensi

infected 5 days previously with a GFP‐expressing clone of

Plasmodium berghei.

Note that oocysts on one side of the gut appear small (in focus) and on the other side large (out of focus). Oocyst numbers in such images are readily counted by simple algorithms.

Figure 4.9 Transmission electron micrograph of 6‐ to 7‐day

Plasmodium falciparum

oocyst in

Anopheles gambiae.

Nuclear profiles contain ‘nucleoli’ (n) and synchronous spindles (s) originating in characteristic invaginations of the nuclear envelope (i). Mitochondrial (m) and apicoplast (here termed spherical bodies, sb) profiles are seen in the cytoplasm. The electron‐dense cyst wall is distinct from the paler‐staining basal lamina of the midgut wall.

Figure 4.10 Transmission electron micrograph of 10‐day

Plasmodium falciparum

oocysts in

Anopheles gambiae.

Sporoblast formation begins (top left oocyst) by expansion of the lumen of the endoplasmic reticulum; thereafter the mitotic nuclei are located immediately beneath the sporoblast plasma membrane, and the sporozoites form by outgrowth from the cell surface.

Figure 4.11 Sequential steps in sporozoite formation in the oocyst.

A,

Vacuole of inner membrane, and vesicle precursors of rhoptries (Rh) and micronemes laid down under plasma membrane adjacent (linked?) to a pole of final nuclear mitotic division.

B,

Assembly and elongation of the IMC and microtubular cytoskeleton draws spindle pole into sporozoite bud. Mitochondrion (Mit) and apicoplast (A) move together into bud.

C,

Maturation of secretory vesicles, differentiation between rhoptry and micronemes detectable.

Figure 4.12 3D reconstruction of anterior pole from cryoelectron tomographs of sporozoite (cf. ookinetes, Fig. 4.3a). Purple, plasma membrane; green, microtubules; yellow, inner membrane complex; pink, rhoptry; blue, micronemes; brown, polar rings (MTOC).

Figure 4.13 Scanning electron micrograph of

Plasmodium yoelii nigeriensis

sporozoites immediately prior to cytokinesis from the sporoblast body of the oocyst.

Figure 4.14 Scanning electron micrograph of

Plasmodium yoelii nigeriensis

sporozoites emerging from catastrophically ruptured oocysts on the midgut of

Anopheles stephensi

.

Figure 4.15 Transmission electron micrograph of

Plasmodium yoelii nigeriensis

sporozoites in the salivary glands of

Anopheles stephensi

. Note sporozoites lying free in the cytoplasm and the acinus of the cells (s, sm) and tightly packed in the, here partially chitinised, lumen (l) of the duct of the gland from where they will be ejected into the skin of the vertebrate host.

Figure 4.16 Graph illustrating the two key population bottlenecks through which malaria parasites pass as they are transmitted from the vertebrate to the mosquito, and from the mosquito back into the vertebrate host. Note parasite abundance is plotted on a log scale.

Figure 4.17 Diagram illustrating the density dependence observed in successive developmental transitions as

Plasmodium berghei

progresses through the mosquito

Anopheles stephensi.

1, gametocytes; 2 and 3, gametes; 4, ookinete; 5, oocyst; 6, salivary gland sporozoites.

Figure 4.18 The theoretical relationships between the reductions that could be achieved by a transmission blocking intervention in oocyst (yellow), ookinete (blue), or macrogametocyte (green) numbers and the corresponding reductions that would ensue in salivary gland sporozoite number. Note how this relationship is markedly affected by the initial challenge infection (described as macrogametocyte number, top left corner of each box A–H) offered to the mosquito.

Figure 4.19 The activities of the most widely used current antimalarial compounds throughout the life cycle of

Plasmodium.

Stars denote components of current artemisinin combination therapies. Data obtained using

P. falciparum, P. yoelii,

or

P. berghei.

Chapter 05

Figure 5.1 Cladogram showing relationships among

Plasmodium

parasite species based on information from Perkins 2002, Martinsen 2008, and Liu 2010.

Figure 5.2 Synteny map of the

Plasmodium vivax

genomes as compared to

(A)

a closely related species,

Plasmodium knowlesi,

and

(B)

a more distantly related species,

Plasmodium falciparum

. Each block of color on the periphery of the circles represents a chromosome. Pk,

Plasmodium knowlesi

; Pv,

Plasmodium vivax;

Pf

, Plasmodium falciparum

. These figures show a larger number of rearrangements that have occurred over a greater amount of evolutionary time. They also show that the rearrangements do not occur as single genes, but as large “synteny blocks.”

Figure 5.3 Evolutionary relatedness of genes of the

pir

families. Each wedge represents a group of paralogues from each different subfamily of

pir

genes. Note that the primate parasite genes group together, the

virs

(from

P. vivax

) and

kirs

(from

P. knowlesi

) group separately from the rodent parasite genes; however, they do not form a single group by species. Similarly, some of the rodent

pirs

group by species relatedness (a group of

yirs

from

P. yoelii

and

birs

from

P. berghei

), shown in Figure 5.1. Other rodent

pirs

group all together. This shows that the defining characteristics of the rodent and primate

pir

paralogues originated after the split of these two larger groups (see Figure 5.1), whereas within the groups many paralogues retain older characteristics from before the species split.

Chapter 06

Figure 6.1 Multiple layers of gene regulation exist in

P. falciparum

.

Figure 6.2 A) Schematic representation of histone marks and histone variants linked with silent and active

var

gene loci. A similar situation applies to other clonally variant gene families. The spatial organization of the

var

gene family in the nucleus is also illustrated. B) Model for the nuclear organization of

P. falciparum

genomic DNA at the ring stage indicating the major nuclear compartments: nucleolus, Pol I and Pol II/III transcription sites, telomeric clusters, and chromosome organization.

Figure 6.3 A pie chart summary of putative transcription factors in the genome of

P. falciparum

as determined by bioinformatic analyses.

Chapter 07

Figure 7.1

A,

Diagram showing how the Bxb1 mediated integration system works.

B,

The

attB

recombination site is introduced into the wild‐type genomic locus of a target gene via single‐crossover homologous recombination.

Figure 7.2 Diagram outlining how the

piggyBac

transposon system works. Parasites are fed erythrocytes electroporated with the pHTH and pXL‐BacII‐DHFR plamids, which are then taken up by the parasites. Expression of the

piggyBac

transposase (scissor symbol) leads to excision of the long terminal repeat (LTR)‐flanked DHFR cassette and insertion into random TTAA sites within

Plasmodium

chromosomes. Insertions can occur within or flanking protein ‐coding sequences. After obtaining a library of parasite clones, the

piggyBac

insertions and their flanking genes responsible for interesting phenotypes can by identified by PCR.

Figure 7.3 Diagram showing how conditional Cre/

loxP

DNA deletion and protein destabilization systems work.

A,

The gene of interest (GOI) is replaced via double site recombination with another version of itself encoding the same amino acids but with a different codon usage. This codon‐optimized gene of interest (CO‐GOI) is flanked with

loxP

recombination sites and is contained on a gene cassette also containing DHFR selectable marker. The CO‐GOI is epitope tagged (HA) so its expression can be verified in place of the original gene. These parasites are then transfected with a Cre recombinase conditionally expressed to ensure deletion of the CO‐GOI only occurs when desired. In the example shown here, Cre is under the control of an anhydrotetracycline (ATc) regulated promoter. In the presence of ATc, the TetR‐TATi2 transactivator protein cannot bind to the

tet07

repeats (vertical bars) preceding the minimal promoter (MinP), leading to weak transcription of Cre. Once ATc is washed out of the parasites, TetR‐TATi2 binds to

tet07

recruiting transcription factors, which results in strong transcription of Cre. The recombinase can then excise the CO‐GOI, producing a phenotype diagnostic of gene function.

B,

To conditionally regulate degradation of a protein encoded by a gene of interest, the GOI is tagged with a DNA encoding an epitope‐tagged destabilization domain 24 mutant (HA‐DD24). The HA‐DD24 DNA is fused to a gene‐targeting flank and replaces the 3′ end of the GOI via single site recombination. During the transfection and integration period, Shld‐1 compound is added to stabilize the GOI‐HA‐DD24 tagged fusion protein. Once the desired parasites expressing the fusion protein have been cloned, Shld1 is washed out and the parasites are examined for a phenotype due to degradation of the GOI‐HA‐DD24.

Chapter 08

Figure 8.1 Overview of transcription profiles of the ApiAp2 genes. The figure shows a list of AP2 domains of 20 ApiAP2 proteins, which are predicted to bind with DNA motifs in

P. falciparum

(Campbell 2010). The name of the AP2 domains, corresponding PlasmoDB gene ID, primary motif recognized, and IDC transcriptional profile are listed in the first, second, third, and fourth columns, respectively. D1, D2, and D3 refer to the domain number on the protein, and DLD refers to two domains (domain linker domain). The transcriptional profiles shown are taken from a previous study (Foth 2011), where microarrays were carried out at 24 time points across the 48‐hour IDC.

Figure 8.2

Overview of the expression, protein, and histone modification profiles during

P. falciparum

IDC.

The figure depicts abundance profiles of 4464 transcripts (corresponding to 4464 genes), 409 protein isoforms (corresponding to 149 genes), and 7260 acetylated histone H4 lysine 8 (H4K8ac) profiles (corresponding to 3108 genes) across the IDC in

P. falciparum

. The results are based on oligonucleotide microarray for mRNA or histone‐modification profiles and 2D‐DIGE for protein profiles and show the prevalence of single peak profiles across the IDC in each case. The heat maps depicting mRNA (on the left), protein (in the middle) and H4K8ac (on the right) abundance profiles comprise 24, 24, and 6 time points, respectively, covering 0 to 48 hours after invasion (hpi). Note that the rows in each heat map representing transcripts, proteins, and H4K8ac

do not

correspond to one another. Each row represents the transcript‐abundance profile for each gene, protein‐abundance profile for each protein isoform, and H4K8ac‐abundance profile for each genetic locus as seen in left, middle, and right panels, respectively. The abundance profiles were sorted according to their Fourier phase, and the color scale represents the lowest smoothed profiles calculated from centered curves of relative log

2

occupancy ratios. Data for this figure were derived from 2 previous studies (Gupta 2013; Foth 2011).

Chapter 09

Figure 9.1 Glycolysis. The classical glycolytic pathway is shown with its different connections to other pathways. Also shown are the transport systems through which glucose is imported and lactate is exported and the pathways of glycerol uptake and production.

Figure 9.2 Pentose phosphate pathway (PPP). The pathway is a major producer of NADPH and of 5‐phospho‐

D

‐ribose 1‐pyrophosphoric acid (PRPP). The PPP can function in different modes depending on the needs of the cell. As shown in the inset, in mode 1, both ribose‐5‐phosphate and NADPH are needed. All the ribulose 5‐phosphate is isomerized to ribose 5‐phosphate, which is used for the synthesis of PRPP. In mode 2, more ribose‐5‐phosphate is needed than NADPH. Ribose 5‐phosphate is synthesized by the non‐oxidative arm using fructose‐6‐phosphate and glyceraldehydes‐3‐phosphate supplied by glycolysis. In mode 3, the cell needs NADPH and ATP but not ribose‐5‐phosphate. Ribulose‐5‐phosphate is converted to fructose‐6‐phosphate and glyceraldehydes‐3‐phosphate, which are channeled into glycolysis. Notice that no gene for encoding transaldolase could be found in the genome, indicated by the “no entry” icon.

Figure 9.3 Pyruvate metabolism. The compartmentation of pyruvate is emphasized where all reactions appearing in the apicoplast are shown on a shaded background. While the apicoplast produces acetyl‐CoA from phosphoenolpyruvate, the mitochondrion uses α‐ketoglutarate for this purpose (see Figure 9.34). Acetyl‐CoA is used both for protein acetylation (mostly histones) and for the synthesis of amino sugars (see Figure 9.5).

Figure 9.4 Glyoxalase and mannose metabolism.

A,

the glyoxalase pathway detoxifies methylglyoxal, which is formed during high rates of glycolysis. The upward path from methylglyoxal is supported by the presence of encoding genes but not by experimental evidence.

B,

Mannose is supplied either from glucose or from the extracellular space. Its products are used eventually for the synthesis of GPI anchors (Figure 9.12) after combining with dolichol (Figure 9.11). Although products can also be used for glycan synthesis, this process is debatable (see text).

Figure 9.5 Aminosugar metabolism. Amino sugars are synthesized as precursors for the generation of GPI anchors and for

N

‐ and

O

‐glycosylations of proteins. The “no entry” sign indicates that a gene encoding this enzyme could not be found in the genome by sequence homology.

Figure 9.6 Fatty acid synthesis in the apicoplast. All this intricate pathway is located in the apicoplast, receiving inputs from different other pathways. The fatty acid type II synthase complex is shown on a shaded background. Also shown are some of the known inhibitors (cerulenin, fenoxaprop clonidafop, thiolactomycin, triclosan) of particular enzymes.

Figure 9.7 Phosphatidylcholine (PC) metabolism. PC is the major constituent of parasite membranes. The two alternative pathways for PC generation are shown – follow the black and grey lines. Also shown are some of the PC degradation products and their recycling.

Figure 9.8 Phosphatidylethanolamine (PE) and phosphatidylserine (PS) metabolism. PE can be synthesized directly from ethanolamine or via the decarboxylation of PS. The intermediate CDP‐diacylglycerol is also the source for the synthesis of cardiolipin, a typical mitochondrial membrane phospholipid.

Figure 9.9 Inositol phosphate metabolism. Inositol can be phosphorylated to various levels, and each product can serve a specific role. Products serve in the synthesis of GPI anchors, in the endocytosis of host cell cytosol, and probably in other, as yet unraveled, signaling pathways.

Figure 9.10 Sphigomyelin (SM) and ceramide metabolism. Sphingomyelin can be obtained from the host cell membrane, but the parasite has the full complement of enzymes that could synthesize SM

de novo

as well as for its conversion ceramide. SM is instrumental in the genesis of the Golgi apparatus and the tubulovesicular membrane (TVM).

Figure 9.11 Isoprenoids metabolism. This pathway derives its substrates from glycolysis and provides substrates for the synthesis of terpenoids (Figure 9.12). It resembles the plant biosynthetic pathway and occurs in the apicoplast. Of note is the use of ferredoxin for redox cycling of NADP/H.

Figure 9.12 Terpenoids metabolism. Using dimethylallyl‐PP and isopentenyl‐PP, the pathway produces farnesyl, geranyl, and geranylgeranyl moieties for the posttranslational modification of proteins. Geranylgeranyl can be further elongated by adding prenyl (3‐methyl‐but‐2‐en‐1‐yl) moieties.

Figure 9.13Dolichol metabolism. Dolichol cycles in the endoplasmic reticulum (ER), where the dolichol phosphate pool provides for

N

‐glycosylation,

O

‐mannosylation and GPI‐anchor biosynthesis. The substrates for dolichol biosynthesis are produced in the apicoplast and must be transferred to the ER.

Figure 9.14 Biosynthesis of GPI anchors. GPI anchors serve to anchor proteins to membranes. Their structure, shown in the right upper corner, reveals that it is composed of inositol phosphate, sugars, and ethanolamine. The different lipid chains and their prevalence are also shown. Notice the first biosynthetic reactions occur at the exoface of the ER, and afterward GlcN‐PI‐Palm flips to endoface, where the path is completed by building the sugar tree.

Figure 9.15 Methionine and polyamine metabolism. Methionine is one of the central amino acids in parasite metabolism. It provides methyl groups for the methylation of DNA, RNA, phospholipids, and proteins, mostly histones, and thus it is essential for regulation of gene expression. Polyamines are cationic compounds that play roles in the cell cycle, cell division, and differentiation.

Figure 9.16 Arginine:proline:asparagine and aspartate metabolism.

A,

Arginine and proline provide ornithine for polyamine biosynthesis. The conversion of arginine to ornithine also reduces the levels of arginine, which is a substrate for NO synthesis, a noxious compound for the parasite.

B,

Aspartate and asparagine obtained from Hb digestion and uptake for the extracellular medium can interconvert. Aspartate is a major substrate for the generation of

N

‐carbamoyl‐

L

‐aspartate, which in turn is a major substrate for pyrimidine synthesis and of adenylo‐succinate, an intermediate in purine metabolism.

Figure 9.17 Glutamate and glutamine metabolism. While being obvious substrates for protein biosynthesis, glutamate serves as a substrate for the synthesis of glutathione and is also converted to α‐ketoglutarate by three alternative paths, to be funneled to the TCA cycle. Glutamine, on the other hand, is a substrate for the synthesis of glucosamine‐6‐P and carbamoyl‐P, which in themselves are substrates for aminosugar and pyrimidine biosyntheses, respectively.

Figure 9.18 Glycine and serine metabolism.The glycine cleavage complex (GCV) is a source of the one carbon donor 5,10‐methylene‐tetrahydrofolate (CH

2

H

4

folate), necessary for nucleotide synthesis (see Figure 9.31). However, a gene necessary for encoding glycine dehydrogenase could not be found in the genome. Glycine is also a substrate for the synthesis of 5‐aminolevulinate. which is used for porphyrin biosynthesis. Serine is a substrate for phosphatidylserine and for sphingomyelinbiosyntheses.

Figure 9.19 eucine:isoleucine and valine metabolism. All three amino acids are possible subjected to degradation. However, the degradation pathways were constructed based on biochemical and genetic knowledge, but none of the degradation products has ever been recorded.

Figure 9.20 Phenylalanine‐tyrosine and lysine metabolism.

A,

Phenylalanine and tyrosine can possibly be degraded by the concerted actions of aspartate transaminase, aromatic‐amino‐acid transaminase, and phenylpyruvate‐tautomerase. These enzymes are not specific for the substrates shown here, but they have a rather wide range of substrates, allowing the drawing of this chart. None of the degradation products has ever been reported. B, Lysine degradation is mediated by rather specific enzymes for which encoding genes have been found in the genome. The possible roles of saccharopine and 5‐oxopentanoate are not known and were not identified yet in the parasite.

Figure 9.21 Selenocysteine metabolism. The role of this 21st amino acid in parasite proteins has received little attention, but all the elements that are involved in its metabolism are present.

Figure 9.22 Purine biosynthesis. The parasite is a purine auxotroph, salvaging host cell purines, preferring hypoxanthine over other nucleobases and nucleosides. This preference is met by the extensive purine salvage processes that take place in the host cell, shown in light grey background. As shown, the starting points for purine biosynthesis may be at different points on the pathway with different substrates, provided all substrates can be imported by means of the nucleotide (Pfnt) carrier. At the end of the paths, deoxynucleosides are used for DNA synthesis and nucleosides for the various types of RNA. Also shown in the inset at the right upper corner are the various paths of nucleoside catabolism.

Figure 9.23 Pyrimidine biosynthesis. Pyrimidine nucleoside synthesis is allegedly

de novo,

starting from initial substrates provided by different metabolic pathways, while other substrates from yet other pathways are added along the way. The oxidation of

L

‐dihydroorotate to orotic acid is coupled to the electron transport chain of the mitochondrion. As in purine metabolism, the reduction of nucleosides to deoxynucleosides is mediated by ribonucleotide reductase, which is then recycled by thioredoxin reductase.

Figure 9.24 CoA biosynthesis. The synthesis of coenzyme A is totally dependent on the import of pantothenate from the extracellular space. This step, as well as some other enzymatic activities along the path, has been investigated to reveal some biochemical details.

Figure 9.25 Nicotinate:nicotinamide and riboflavin metabolism.

A,

NAD and NADP, produced by the nicotinate and nicotinamide pathway, are essential co‐factors for many redox reactions. Nicotinate and/or nicotinamide must be obtained from the extracellular space. Nicotinate is the first substrate of the pathway.

B,

Riboflavin is obtained from the host cell or from the extracellular medium. In one step it is converted to FMN, and a second enzymatic step produces FAD. Both FMN and FAD are co‐factors in redox reactions.

Figure 9.26 Thiamine (vitamin B

1

) metabolism. As shown, thiamine pyrophosphate (TPP), which is the active form of vitamin B

1

, is an indispensable co‐factor for many essential enzymes (right, lower corner). Although thiamine can be obtained from the extracellular space an in one step, the parasite has the full complement of enzymes that are needed for the synthesis of TPP. The biosynthesis of thiamine occurs by the combination of the pyrimidine (right) branch and the thiazole (left) branch. In the absence of thiamine‐phosphate kinase, thiamine‐P is first dephosphorylated to thiamine and then rephosphorylated to TPP.

Figure 9.27 Pyridoxal phosphate (Vitamin B

6

). Pyridoxal 5‐phosphate (PLP) is the active cofactor, known to serve ~150 enzymes (may be less in the parasite). Some examples in the parasite are shown in the lower right. The biosynthetic pathway uses intermediates from glycolysis, PPP, and glutamate. Pdx1/Pdx2 is a multimeric glutamine amidotransferase. Pyridoxal kinase accepts other B

6

vitamers (pyridoxal, pyridoxine, and pyridoxamine), which can be salvaged from the host erythrocyte.

Figure 9.28 Ubiquinone biosynthesis. Ubiquinones are electron carriers in the mitochondrial electron transport. The quinoid nucleus of ubiquinone is composed of 4‐hydroxybenzoate derived from chorismate and all‐

trans

geranyl‐geranyl‐PP, and then undergoes a series of modifications such as hydroxylations,

O

‐methylations, methylations, and decarboxylations. Note that genes encoding for two essential enzymes in the biosynthetic chain could not be found in the genome.

Figure 9.29 Porphyrin metabolism. The parasite cannot use the heme of the host cell Hb and synthesizes it, dividing the chore between the two juxtaposed organelles of the parasite, starting in the mitochondrion, then in the apicoplast; after a short sojourn in the cytosol, the path goes back to the mitochondrion for the final insertion of iron by ferrochelatase. Protoheme then serves as a prosthetic group in various cytochromes. Notice that aminolevulinatedehydratase (ALAD) has been shown to be imported from the host cell. Some heme is derived from hemoglobin digestion.

Figure 9.30 Shikimate biosynthesis. The products of the shikimate pathway serve for ubiquinone and folate synthesis, yet it poses one of the outstanding enigmas in the biochemistry of the parasite: for

four enzymes in a row,

no encoding genes could be found in the genome, and neither could an alternative pathway be identified. Yet the necessary intermediates that serve the depending pathways are there.

Figure 9.31 Folate biosynthesis. The folate pathway generates tetrahydrofolate (THF) essential cofactors for single carbon‐transfer reactions in the synthesis of nucleic acids. The pathway, which is absent in humans, has been the target of antimalarial chemotherapy. Some drugs are shown next to their targets.

Figure 9.32 Redox metabolism. An intricate network of antioxidant defense is put in place in order to defend the parasite from intrinsic and extrinsic oxidative challenges. The system consists of glutathione, thioredoxin, and peroxiredoxin and their redox cycling, mostly through NADPH, which is recycled mostly by the PPP.

Figure 9.33 Mitochondrial electron flow (ETC). Because oxidative phosphorylation could not be demonstrated in the parasite, the role of the ETC is considerably reduced, and its main function is the removal of electrons from dihydroorotic acid during its oxidation to orotate. However, new functions seem to emerge explaining the roles of the various dehydrogenases that are linked to the TEC (see text).

Figure 9.34 Mitochondrial TCA cycle. All genes encoding TCA enzymes and the ancillary transporters are found in the genome. Metabolomic studies using stable isotope‐labeled substrates in which the number and position of labeled atoms are determined have revealed that essentially a typical TCA cycle is operating, although it uses glutamate rather than pyruvate as its main substrate. Results also showed that the BCKDH complex is able to convert pyruvate to acetyl‐CoA (see Figure 9.35). The major products are acetyl‐CoA used for various acetylation reactions, succinyl‐CoA used in the synthesis of porphyrin and NADH, which fuels the mitochondrial electron transport. The malate shuttle (inset) is able to transfer reducing equivalents from the cytosol to the mitochondrial matrix.

Figure 9.35 Acetyl‐CoA production in the mitochondrion. The mitochondrion‐located branched‐chain α‐keto acid dehydrogenase complex (BCKDH) converts pyruvate to acetyl‐CoA. It uses the same E3 dihydrolipoamide dehydrogenase as the TCA cycle but has particular E1 and E2 subunits.

Figure 9.36 Hemoglobin digestion and ferriprotoporphyrin IX polymerization. Polyphosphorylated phosphoinositides (PIPs) of parasite origin and their specific kinase are exported into the host cell cytosol and participate in its endocytosis. Hb delivered by endocytic vesicles to the FV releases its ferriprotoporphyrin IX (FPIX) and the globin is degraded by sequential action of aspartate proteases, cysteine proteinases, and dipeptidylaminopeptidase. The parasites does not have carboxypeptidase activity, and amino peptidase, distributed between the FV and the cytosol, terminate the digestion to individual amino acids. FPIX released from Hb during vacuolar digestion is a noxious compound that is detoxified by polymerizing it to hemozoin, assisted by histidine‐rich proteins and lipids. A small portion of FPIX evades polymerization and is destroyed in the cytosol by GSH.

Chapter 10

Figure 10.1 Calcium homeostasis and signaling in

Plasmodium.

Calcium is stored in the endoplasmic reticulum (ER), acidocalciosomes, or mitochondria (not indicated here) in the parasite. SERCA‐like calcium ATPase may facilitate the entry of calcium into the ER. The activation of unidentified receptors (??) by extracellular signals like xanthurenic acid may cause the activation of phospholipase C PfPLC, which yields IP3 and DAG. IP3 facilitates the calcium release from the ER, which may also be mediated by the cADP–ribose–RyR–like pathway. The IP3R and RyR have not been identified in the parasite. Ca

2+

regulates a wide variety of parasitic processes via targets like CDPKs, which play diverse roles in calcium signaling.

Figure 10.2 Phosphatidylinositol‐3‐phosphate (PI3P) in

Plasmodium falciparum

.

Plasmodium

codes for a single PI3‐kinase, PfPI3K, which can generate PI3P (

smaller grey circles

) as well as other 3′‐phosphorylated PIPs. PfPI3K (

larger white circles

) is present in vesicular compartments at PM/PVM and food vacuole. It is also exported to the host RBCs and it may generate 3′‐PIPs at these locations. PI3P was localized at the cytoplasmic face of the food vacuole (FV) and apicoplast (AP) membrane and the endoplasmic reticulum (ER) lumen. It may be involved in the trafficking of proteins to these organelles. PfPI3K may regulate the endocytosis of hemoglobin (

black circles

) to the food vacuole. PI3P may interact with the PEXEL motif (

rhombus

) containing proteins (

pentagons

) in the ER, which facilitates their export to the host RBC after cleavage by plasmepsin V. PfPI3K exported to the host erythrocyte is active; however, its role in the host is unclear.

Figure 10.3 Cyclic nucleotide signaling in

Plasmodium falciparum.

Adenylate (AC) and guanylate cyclases (GC) generate cAMP and cGMP, respectively, and phosphodiesterases (PDE) hydrolyze these cyclic nucleotides. Binding of cAMP to PfPKA‐R dissociates it from PfPKAc, resulting in its activation. PfPKG is activated upon binding of cGMP to its N‐terminus. The identity of substrates for PfPKA and PfPKG will provide insights into their role in the parasite. Some of the parasitic processes in which these kinases may be involved are indicated.

Chapter 11

Figure 11.1

A,

Schematic showing the multiple membranes within an infected erythrocyte and locations of specific transport proteins. PSAC is on the host erythrocyte membrane (RBC). PTEX and the PVM channel reside on the parasitophorous vacuolar membrane (PVM). A glucose transporter (HT) and an aquaglyceroporin (AQP) are shown on the parasite plasma membrane (PPM). PfCRT and PfMDR1 are shown on the digestive vacuolar membrane (DV); question marks reflect uncertainties in the properties of these transporters. Hemozoin is shown as crystalline material within the DV. ATP synthase (ATP Syn) localizes to the inner membrane of the mitochondrion. N indicates the nucleus.

B,

Individual panels showing relevant features of selected transporters. PSAC activity is determined by CLAG3, a protein that is integral to the host membrane; ribbon diagram shows a possible transmembrane topology of CLAG3. A highly variable domain is exposed at the host membrane, based on protease susceptibility studies (Nguitragool 2011). PTEX localizes to the PVM membrane, is a high‐molecular‐weight complex of several parasite proteins, and may function as a protein translocon; Crystal structure of AQP from

P. falciparum

, showing four‐fold symmetry and four parallel routes for water transport; ATP synthase is a multi‐subunit motor on the inner mitochondrial membrane; in most organisms, this enzyme uses the downhill uptake of H

+

into the matrix to synthesize ATP. In a few organisms, H

+

is exported using the energy of ATP hydrolysis; the precise function in malaria parasites is uncertain.

Chapter 12

Figure 12.1 Domain organization of plasmepsins. Shown on the top is the general architecture of

P. falciparum

plasmepsins, with prodomain (Pro) and mature protease domain (Protease). The hydrophobic transmembrane region (TM) in the prodomain and active site motif 1 and motif 2 (star) in the protease domain are indicated. The names of plasmepsins sharing this architecture, the sizes, and the sequences of active site motifs are shown below the schematic. The sizes are scaled (≈150 amino acid residues/inch) and the amino acid numbers are based on the protease sequences on the PlasmoDB.

Figure 12.2 Domain organization of cysteine proteases. Shown is a comparison of general architectures of different

P. falciparum

cysteine proteases with papain. The prodomains (Pro) and mature protease domain (Mat) are indicated with hydrophobic transmembrane regions (black bars), the active site amino acid residues (C, H, and N), processing site (arrow), and the number of amino acid residues or boundary of the indicated region (bracket). Falcipain‐2A is a representative of the FP2/3 subfamily proteases of malaria parasites, which differ from all other known papain‐like proteases in having the refolding (Ref) and hemoglobin‐binding domains (Hb). DPAP1 and other DPAPs, as in human cathepsin C, contain a putative exclusion domain (white box) that confers exopeptidase dipeptidyl activity to cathepsin C‐like proteases. The

Plasmodium

calpain is a giant protease with long amino‐terminal and C‐terminal extensions (broken bar), the protease region (DIIa and DIIb) is interrupted by an insert, and the Ca

2+

‐binding region (DIII) is indicated. The active site residues (C, H, and N) common to all papain‐family proteases are indicated. The sizes are scaled (≈150 amino acid residues/inch) and the amino acid numbers are based on the protease sequences on the PlasmoDB.

Figure 12.3 Schematics of the

P. falciparum

metallopeptidases. The indicated metallopeptidases are shown with the total number of amino acid residues in brackets. Active site residues are shown below the indicated protease, underlined residues are putative metal binding ligands, and residues in bold font are catalytic residues. PfA‐M1 has a hydrophobic transmembrane region in the N‐terminus (black bar), and PfMet1b has a Zn‐finger motif (triangle). The sizes are scaled (≈150 amino acid residues/inch) and the amino acid numbers are based on the protease sequences on the PlasmoDB.

Figure 12.4 Schematic representation of the subtilisin domain organization. Shown are the zymogen forms with hydrophobic domains (black bar), prodomains (Pro), protease domains (Protease), cytoplasmic tail (Cyt), and the residues forming the active site. The PfSUB1 zymogen is processed into the protease domain (p54) and then to the p47 form. The PfSUB2 zymogen is processed to the p75 form. The prodomain and protease domain boundaries of PfSUB3 have been predicted based on sequence analysis on the pfam database. The sizes are scaled (≈150 amino acid residues/inch) and the amino acid numbers are based on the subtilisin sequences on the PlasmoDB.

Figure 12.5 Schematic of the domain organization of

P. falciparum

rhomboids. The schematic of PfROM1 shows 7 transmembrane domains (black rectangles), with the active site serine (S) and histidine (H) residues. The table shows key features (TM, number of transmembrane domains; the catalytic serine (GxSx) and histidine (HxxGxxxG) residue motifs; and the length of the protein) of the indicated rhomboids. The catalytic residues are in bold font. Note that ROM8 has Glu in place of Gly in the GxSx motif, and ROM10 does not seem to have the catalytic HxxGxxxG motif, suggesting that these two are not active proteases. The sequences were analyzed for the presence of motifs on the MEROPS and Pfam databases, and transmembrane domains were predicted using the TMPred and TMHMM on the SACS (http://www.sacs.ucsf.edu/). The sizes are scaled (≈150 amino acid residues/inch) and the amino acid numbers are based on the protease sequences on the PlasmoDB.

Figure 12.6 Schematic of hemoglobin degradation. Shown is a cartoon of an erythrocyte infected with the trophozoite stage, which voraciously degrades hemoglobin in the food vacuole. The parasite takes up the erythrocyte cytosol through a cytostome‐like organelle (1) wherefrom vesicles bud off (2) and transport hemoglobin to the food vacuole. Hemoglobin appears to be first degraded (3) by falcipains and plasmepsins (FP/PM) that generate oligopeptides, which are further degraded (4) by oligopeptidases (OP: falcilysin, PfA‐M1, and DPAP1) into smaller peptides. The smaller peptides have been proposed to be transported into the parasite cytosol (5), where aminopeptidases (AP: PfLAP and PfAP) degrade these into free amino acids, which are used by the parasite. Hemoglobin degradation (3) results release of free heme, which is polymerized into a nontoxic polymer, known as hemozoin (Hz).

Chapter 13

Figure 13.1 The growth and development of medicines for malaria control and elimination.

Figure 13.2 The research and development process for new antimalarial medicines, giving relative timelines for each phase. The times given are indicative, but historical timescales have often been significantly slower for a number of reasons, including irregular funding.

Figure 13.3 The global malaria portfolio at the end of 2012. inhrs.: inhibitors.

Chapter 14

Figure 14.1 Numbers of reported cases of

P. falciparum

malaria and

P. vivax

malaria over the 50 years 1961 to 2011.

Figure 14.2 Decrease in parasite density following treatment.

Figure 14.3 The figure shows

pfmdr1

copy numbers at several sites in Cambodia: Pailin, Kampoong Seila, Chumkiri, Memut, and Rattanakiri.

Chapter 15

Figure 15.1 Areas at risk for

Plasmodium falciparum

transmission during 2010 (Malaria Atlas Project 2010).

Figure 15.2

Plasmodium falciparum

distribution in Africa by endemicity level for 2010 (Gething 2011).

Figure 15.3

Plasmodium falciparum

life cycle.

Figure 15.4 Malaria program milestones toward malaria elimination.

API

, annual parasitic incidence

SPR

, slide positivity rate.

Chapter 16

Figure 16.1 Sequestration of

P. falciparum

‐infected red blood cells is thought to play a key role in parasite virulence and occurs in several vascular beds in the human host. The histologic appearance and level of sequestration vary between vascular beds and between infections. The level of sequestered parasites in the cerebral vasculature correlates with cerebral malaria risk in several studies of adults and children. In addition to brain, lungs, heart, and gut, sequestered parasites appear in vessels of skin, adipose, kidneys, and other organs.

Figure 16.2 Pathological processes implicated in the pathogenesis of malarial disease are associated with specific phases of parasite development. Specific factors that reduce the risk of severe malaria are thought to affect these pathological processes or to prevent the initial infection that can result in severe malaria. These factors often form the conceptual basis for adjunctive therapies to reduce mortality after severe malaria develops or for vaccines to prevent the development of severe malaria.

Chapter 18

Figure 18.1

The

P. falciparum

life cycle.

The

P. falciparum

life cycle in humans includes the asymptomatic liver stage; the blood stage which causes disease; and the sexual gametocyte blood stage which infects mosquitoes that transmit the parasite. Infection begins when

Anopheles

mosquitoes inject sporozoites into the skin where they may persist for days

(a).

Sporozoites migrate from the skin via the blood to the liver and invade a small number of hepatocytes

(b).

Some sporozoites migrate to the draining lymph nodes where sporozoites antigens can be presented by dendritic (DCs) to CD8

+

T cells that then migrate to the liver (

b

′). Each sporozoite gives rise to tens of thousands of asexual parasites called merozoites

(c).

Approximately one week after hepatocyte invasion merozoites exit the liver into the bloodstream and begin a 48 h cycle

(d)

of RBC invasion, replication, RBC rupture, and merozoites release

(e).

Symptoms only occur during the blood‐stage and can begin as early as 3 days after the release of merozoites from the liver. Once inside RBCs the parasite exports proteins such as PfEMP1s to the RBC surface. PfEMP1s mediate binding of iRBCs to the microvascular endothelium of various tissues

(f)

sequestering the parasites from clearance in the spleen and promoting the inflammation and circulatory obstruction associated with clinical syndromes. In severe disease, such as cerebral malaria, the iRBC sequester in the brain and are associated with microhemorrhages and brain damage

(g).

PfEMP1‐mediated rosetting (binding of iRBCs to RBCs) may also contribute to disease

(h).

A small number of blood‐stage parasites differentiate into sexual gametocytes

(i)

which are taken up by mosquitoes

(j)

where they differentiate into gametes that fuse to form zygotes, and then develop into motile ookinetes

(k)

. The ookinetes cross the midgut wall and form oocysts

(l)

that develop into sporozoites that enter mosquito salivary glands to complete the life cycle

(a).

Figure 18.2

Points of control of severe disease.

Theoretically, risk of severe malaria could be reduced at several points in the progression of the disease. Transmission can be controlled by mosquito control and bed nets. Infection and the liver stage of the disease do not appear to be points of control in that resistance to liver stage disease is not acquired naturally although future vaccines may produce protection against this stage. The blood stage of the disease can be controlled by anti‐malarials and by mutations in a number of genes expressed in the RBC that block invasion or inhibit parasite growth. Once a blood stage infection is established the innate and adaptive immune responses are essential to prevent disease or limit the disease to mild disease.

Figure 18.3

Selecting for and controlling genetic susceptibility to autoimmunity in malaria endemic areas.

We propose that genetic susceptibility to the autoimmune disease, SLE, may have been selected for in Africa as it protects from severe disease.

Top.

In the US, SLE

s

genes would initiate inflammation that would trigger an anti‐inflammatory response. As the anti‐inflammatory response wanes the uncontrolled SLE

s

‐mediated inflammation would rise again. Ultimately, the balance would tip toward inflammation and autoimmune disease.

Bottom.

In Africa SLE

s

genes have the same potential to cause inflammatory autoimmunity. However, this potential is attenuated by environmental pathogens,

P. falciparum

infections themselves or perhaps viral infections such as EBV resulting in an anti‐inflammatory environment that protects against both severe disease and autoimmunity.

Figure 18.4

Pro‐ and anti‐inflammatory battles in malaria.

Blood stage infections induce inflammatory responses producing the symptoms of mild malaria. To control the inflammatory response and avoid damage, the immune system mounts an anti‐inflammatory response. Malaria symptoms during the anti‐inflammatory response may be attenuated producing asymptomatic infections. As the anti‐inflammatory response wanes new

P. falciparum

infections produce inflammation and disease. We propose that the anti‐inflammatory response although protective may hinder the acquisition of MBC and LLPCs.

Chapter 19

Figure 19.1

Immune mechanisms can target malaria parasites at different stages in the life cycle.

Antibodies (Abs) elicited against sporozoite surface antigens (Ags) may target sporozoites and block hepatocyte invasion. Cellular immune responses including cytotoxic CD8 T‐cell responses may clear infected hepatocytes and prevent emergence of merozoites. Antibodies against merozoite Ags may block erythrocyte invasion by blocking receptor–ligand interactions that mediate invasion or by antibody‐dependent cellular inhibition (ADCI). Antibodies against variant surface antigens expressed on the surface of

P. falciparum

–infected trophozoites and schizonts may block cytoadherence and mediate clearance by alternative mechanisms such as antibody‐dependent opsonization. Antibodies against sexual or transmission stages will block transmission from infected to uninfected individuals.

Chapter 20

Figure 20.1 Evidence for rapid acquisition of immunity to

P. vivax

in PNG children.

A

, Incidence of malaria in a cohort of children 1 to 4 years (Lin 2010a).

B

, Time to first PCR‐positive infection and clinical episode in children 5 to 14 years (Michon 2007).

Figure 20.2 Thorax x‐rays and computed tomography of a 40‐year‐old patient with diagnosis of

P. vivax

monoinfection by TBS and PCR, in which other chronic and acute comorbidities were ruled out, developing ARDS after chloroquine was started. Phenomena behind this common clinical complication, such as cytoahesion, still need to be clarified.

Figure 20.3 Subtelomeric multigene families of

Plasmodium vivax

and

P. falciparum

. Protein similarity relations among

P. vivax

and

P. falciparum

were obtained when running a new clustering procedure (Lopez 2013) over the subtelomeric families of

P. vivax

(Pv or v): VIR, PvPIRA‐D‐H, Pv‐fam, v‐Pf‐fam, vRHOPTRY, v‐PST‐A, and

P. falciparum

(Pf or f): VAR, RIFIN, STEVOR, Pfmc‐2TM, SURFIN, fRHOPTRY, fHAD (Carlton 2008). The position of the clusters with respect to the axes is just a qualitative representation; axes do not represent any metric. Graph figures were obtained using BioLayout (Crooks 2004).

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