University of Khartoum

Graduate College

Medical and Health Studies Board

INDUCED ERYTHROCYTE SUICIDAL DEATH AND ITS EFFECTS ON THE PARASITEMIA AND SURVIVAL OF PLASMODIUM BERGHEI

INFECTED MICE: ROLE OF AMPHOTERICIN B AND GUM ARABIC

By

Adil Ballal Mohammed BSc, MSc, U of K

A thesis submitted for the degree of Ph. D. in Human Physiology

Supervisor

Dr. Amal Mahmoud Saeed, MBBS, Ph. D.

Associated Professor of Human Physiology

Department of Physiology, Faculty of Medicine,

University of Khartoum

2010

 

 

 

 

Dedication

 

 

To

my wife Hanadi,

my son Mohammed,

my daughters Maryam and Mayada

 

I  

Table of contents: Page No. Acknowledgement.............................................................................................................................III Abbreviations....................................................................................................................................IV Abstract............................................................................................................................................VII Arabic abstract...................................................................................................................................IX List of figures....................................................................................................................................XI

Chapter One 1. Introduction.....................................................................................................................................1 1. 1. Malaria.........................................................................................................................................1 1. 1. 1. History.....................................................................................................................................1 1. 1. 2. Epidemiology of malaria.........................................................................................................2 1. 1. 3. Clinical manifestations............................................................................................................3 1. 1. 4. The parasites............................................................................................................................4 1. 1. 5. Life cycle of malaria parasites.................................................................................................4 1. 1. 5. 1. Tissue schizogony (pre-erythrocytic schizogony)...............................................................5 1. 1. 5. 2. Erythrocytic schizogony......................................................................................................6 1. 1. 5. 3. Sexual phase in the mosquito (Sporogony).........................................................................7 1. 1. 6. Malaria immunity....................................................................................................................7 1. 1. 6. 1. Innate immunity...................................................................................................................8 1. 1. 6. 2. Acquired immunity..............................................................................................................8 1. 1. 6. 3. Strain specific immunity......................................................................................................8 1. 1. 7. Malaria diagnosis.....................................................................................................................8 1. 1. 8. Malaria chemotherapy.............................................................................................................9 1. 1. 8. 1. 4, 8-aminoquinoline drugs...................................................................................................9 1. 1. 8. 2. Antifolate drugs.................................................................................................................10 1. 1. 8. 3. Quinine..............................................................................................................................10 1. 1. 8. 4. Artemisinin and its derivatives..........................................................................................11 1. 1. 8. 5. Combination therapy to combat the spread of drug resistance..........................................11 1. 2. Apoptosis...................................................................................................................................12 1. 2. 1. Morphology of Apoptosis.....................................................................................................12 1. 2. 2. Classification of cell death: apoptosis and necrosis..............................................................13 1. 2. 3. Mechanisms of apoptosis.......................................................................................................14 1. 2. 4. Biochemical features of apoptosis.........................................................................................15 1. 2. 4. 1. Extrinsic pathway of apoptosis..........................................................................................16 1. 2. 4. 2. Perforin/granzyme pathway of apoptosis..........................................................................17 1. 2. 4. 3. Intrinsic pathway of apoptosis...........................................................................................18 1. 2. 4. 4. Execution pathway of apoptosis........................................................................................19 1. 2. 5. Physiologic apoptosis............................................................................................................19 1. 2. 6. Pathologic apoptosis..............................................................................................................20 1. 2. 7. Inhibition of apoptosis...........................................................................................................20 1. 2. 8. Assays for apoptosis..............................................................................................................21 1. 2. 8. 1. Cytomorphological alterations..........................................................................................21 1. 2. 8. 2. DNA fragmentation...........................................................................................................22 1. 2. 8. 3. Membrane alterations........................................................................................................22 1. 2. 8. 4. Detection of apoptosis in whole mounts............................................................................22 1. 2. 8. 5. Mitochondrial assays.........................................................................................................23 1. 3. Erythrocyte suicidal death (Erypyosis)......................................................................................23 1. 3. 1. Normal erythrocyte membrane and cation transport.............................................................23 1. 3. 1. 1. Na+ / K+ pump in non-infected erythrocytes......................................................................25 1. 3. 1. 2. Nonselective cation channels in non-infected erythrocytes...............................................26 1. 3. 1. 3. Ca2+ activated K+ channels (Gardos).................................................................................27 1. 3. 2. Mechanisms of erythrocyte cell death or eryptosis...............................................................28 1. 3. 2. 1. Role of prostaglandins in stimulation of erythrocyte cation channels...............................28

II  

Page No. 1. 3. 2. 2. Role of Ca2+ sensitive K+ channels (Gardos channels) in eryptosis................................28 1. 3. 2. 3. Role of protein kinase C in eryptosis.................................................................................29 1. 3. 2. 4. Role of oxidative stress and caspase activation in eryptosis.............................................29 1. 3. 2. 5. Role of platelet-activating factor (PAF) and stimulation of

sphingomyelinase in eryptosis.............................................................................................30 1. 3. 3. Eryptosis inhibitors................................................................................................................30 1. 3. 4. Clinical conditions associated with eryptosis........................................................................31 1. 3. 5. Eryptosis in malaria...............................................................................................................32 1. 3. 5. 1. Stimulation of membrane transport in P. falciparum infected

erythrocytes and activation of ion channels.........................................................................32 1. 3. 5. 2. Induction of eryptosis by P. falciparum infection.............................................................33 1. 3. 5. 3. Role of eryptosis in parasitized erythrocytes.....................................................................33 1. 4. Rodent malaria parasites as models for human malaria............................................................34 1. 5. Pharmacological agents.............................................................................................................35 1. 5. 1. Gum Arabic...........................................................................................................................35 1. 5. 2. AmphotericinB......................................................................................................................36 1. 5. 2. 1. Mode of action...................................................................................................................37 1. 5. 2. 2. Side effects........................................................................................................................37 1. 6. Rational of the study..................................................................................................................37 1. 7. The objectives of the study........................................................................................................39 1. 7 1. General objective....................................................................................................................39 1. 7. 2. Specific objectives.................................................................................................................39

Chapter Two 2. Materials and methods...................................................................................................................40 2. 1. Animals......................................................................................................................................40 2. 2. Parasites.....................................................................................................................................40 2. 3. Infection of mice........................................................................................................................40 2. 4. Gum Arabic treatment...............................................................................................................40 2. 5. Amphotericin B administration.................................................................................................41 2. 6. Preparation of human erythrocytes............................................................................................41 2.7. Investigations..............................................................................................................................41 2.7.1. In vitro culture of Plasmodium falcifarum infected human erythrocytes................................41 2.7. 1. 1. Freezing and sawing of parasites........................................................................................42 2.7. 1. 2. Isosmotic sorbitol synchronization of P. falciparum infected human erythrocytes...........43 2.7. 1. 3. In vitro P. falciparum growth assay...................................................................................43 2.7. 2. Intraerythrocytic DNA amplification of P. falciparum..........................................................44 2.7. 3. Annexin binding experiments (Determination of phosphatidylserine exposure)...................44 2.7. 4. Determination of intracellular Ca

2+ influx in the erythrocytes...............................................45

2.7. 5. Measurement of hemolysis.....................................................................................................46 2.7. 6. In vivo proliferation of P. berghei ANKA..............................................................................46 2.7. 7. Measurement of the in vivo clearance of fluorescence-labelled erythrocytes........................46 2.7. 8. Measurement of the fluorescence-labelled erythrocytes in the spleens of mice....................47 2.7. 9. Data analysis and statistics.....................................................................................................47

Chapter Three 3. Results...........................................................................................................................................48 3. 1. Influence of amphotericin B on the course of malaria..............................................................48 3. 2. Influence of butyrate and gum Arabic on the course of malaria...............................................50

Chapter Four 4. Discussion......................................................................................................................................71 4. 1. Influence of amphotericin B on the course of malaria..............................................................72 4. 2. Influence of butyrate and gum Arabic on the course of malaria...............................................75 References.........................................................................................................................................81 Appendices........................................................................................................................................95 Appendices......................................................................................................................................103

III  

Acknowledgements:

I would like to express my deep and sincere gratitude to my supervisor, Dr. Amal Mahmoud Saeed, Department of Physiology, Faculty of Medicine, University of Khartoum for providing me with unique opportunity to work with her. I do appreciate her assistance, patience, continuous support and valuable suggestions. Her wide knowledge and her logical way of thinking have been of great value for me. Her understanding, encouraging and personal guidance have provided a good basis for the present work.

I wish to thanks Professor Florian Lang, Department of Physiology, Eberhard-Karls-University of Tübingen, Germany, for his usual unlimited support, encouragement and for offering me a space at his laboratories and providing a healthy scientific environment and facilities to accomplish the practical part of this study.

I wish to express my gratitude and sincere thanks to Professor Sayda H. El Safi, former Dean of the Faculty of Medical Laboratory Sciences, University of Khartoum, for her unlimited support and interest. I warmly thank Dr. Ahmed M Abd Alhaleem, vice Dean of Faculty of Medical Laboratory Science, for his valuable advice and friendly help.

Deep gratitude is due to Dr. Atif Hassan Khirelsied, Faculty of Medicine and Health Sciences, International University of Africa, for his great help and valuable suggestions.

I am very indebted to all the members of the Unit of Basic Medical Sciences, Faculty of Medical Laboratory Sciences, University of Khartoum for their co-operation .advices and support during the course of this study. Special and deep thanks to Dr. Thoraya M. El Hassan and Mr. Omar Osman.

I am very grateful to Dr. Michael Föller, Mr. Diwakar Bobbala, Miss. Adriana Estremera, Mss. Ioana Alesutan and for the whole entire staff of the Institute of Physiology, Eberhard-Karls-University of Tübingen, Germany, for moral support and help.

This work would have never been done without the unlimited assistance and support of friends and colleagues, Dr. Amir Abu Shouk, Dr. Nadia Omar, Mrs. Magbola Mamoun, Dr. A/El Hafiz A. Bashear, Dr. Tarig Hakim, Mr. Awad Gaber, Mr. Hisham Hamid, Mr. M Osman El Hassan, Mr. Salah M Idrees and Mr. El Sideeg H. El Rasoul. Special sincere gratitude and thanks are extended to Mr. Yasser El Tayb El Mahdi for his unlimited help and interest. I extend my deep thanks to all of the members of the Department of Physiology, Faculty of Medicine, University of Khartoum.

Finally I want to thank my family. The encouragement and support from my beloved wife Hanadi and our always positive and joyful children Mohammed, Maryam and mayada is a powerful source of inspiration and energy. A special thought is devoted to my mother and my brothers for a never-ending support. I owe my loving thanks to my stepmother Ebititham and her daughters Hana and Shadia.

IV  

Abbreviations

AIDS Acquired immune deficiency syndrome

AIF apoptosis-inducing factor

AMA1 apical membrane antigen 1

AO acridine orange

Apo3 L Apoptosis 3 ligand

ATP Adenosine triphosphate

Bcl-2 B-cell lymphoma 2

[Ca2+]i (free) cytosolic ionic calcium concentration

CAD caspase-activated Dnase

CD 95, Apo-1 cluster of differentiation 95, Fas receptor

CFSF carboxyfluorescein diacetate, succinimidyl ester

CFTR cystic fibrosis transmembrane conductance regulator

COX cyclooxygenase

CT combination therapy

Df dilution factor

DIDS 4,4’ - diisothiocyantostilbene – 2,2’ – disulfonic acid

DISC death-inducing signalling complex

DMSO dimethylsulfoxid

DR death receptors

EDTA ethylenediaminetetraacetic acid, [Ethylenedinitrilo]tetraacetic acid

EIA enzyme immunoassay

ELISA enzyme-linked immunosorbent assay

FACS fluorescence assisted cell sorting

FADD Fas-associated death domain

Fas-L fatty acid synthase ligand

Fas-R fatty acid synthase receptors

FITC fluorescein isothiocyanate

FP IX ferri/ferroprotoporphyrin IX, free heme

G6PDH glucose-6-phosphate dehydrogenase

GA gum Arabic

GAPDH glycerinaldehyd-3-phosphat-dehydrogenase

V  

Gardos channel calcium activated potassium channel

GSH Gluthathione

GSSG oxidized gluthathione

Hb Hemoglobin

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HIV human immunodeficiency virus

IAP inhibitors of apoptosis proteins

ICAD Inhibitor of Caspase Activated DNAse

ICAM intracellular adhesion molecule

IFA immunofluorescence assay

IFN interferon

IL Interleukin

L-NAME L-NG-Nitroarginine methyl ester (hydrochloride)

NO Nitric Oxide

LSCM Laser scanning confocal microscopy

MACS magnetic assisted cell sorting

MAEBL membrane antigen erythrocyte binding protein, paralogue of both

AMA1 and DBL-EBP

MOM mitochondrial outer membrane

MPT Mitochondrial permeability transition

MSP merozoite surface protein

NBS Nile blue sulphate

NF-κ B nuclear factor kappaB

NHE sodium/hydrogen exchanger

NMDG+ N-methyl-D-glucamine

NPP New Permeability Pathway

NPPB 5-Nitro-2- (3-phenylpropylamino) benzoic acid

NR Neutral red

NSC nonselective cation

NuMA nuclear mitotic apparatus

P. (pf, Pf) Plasmodium (Plasmodium falciparum)

PAF platelet activation factor

PARP poly [(Adenosine-5'-triphosphate)-Ribose] polymerase

PBS phosphate buffered saline

VI  

PC phosphatidylcholine

PCD programmed cell death

PCR polymerase chain reaction

PE phosphatidylethanolamine

PfEMP Plasmodium falciparum erythrocyte membrane protein

PG prostaglandin

PK protein kinase

PL phospholipase

PS phosphatidylserine

RBC red blood cell

RDTs Rapid Diagnostic Tests

RIP receptor-interacting protein

RPMI Roswell Park Memorial Institute

RT room temperature

SM sphingomyelin

Smac/DIABLO Second Mitochondria-derived Activator of Caspases/Direct IAP

Binding Protein with Low PI

TH T- helper cells

TNF tumor necrosis factor

TNFR tumor necrosis factor receptor

TRADD TNFReceptor-associated death domain

TRAP-PfSSP2 thrombospondin-related anonymous protein, P. falciparum sporozoite

surface protein type 2

TRP transient receptor potential

TSP thrombospondin

VII  

Abstract:

Background: Malaria is one of the most devastating diseases with lethal outcome of more

than one million humans per year. This study intended for a novel drug discovery in order

to prevent the Plasmodium related resistance focusing on identifying and targeting host

factors essential for pathogen entry, survival and proliferation. The innovative methods

involve induction of suicidal death in infected erythrocytes (eryptosis) and recognition by

the spleen macrophages to get rid of the pathogen and prevent the further course of the

disease. Eryptosis is characterized by cell shrinkage, membrane blebbing and cell

membrane phospholipid scrambling with phosphatidylserine exposure at the cell surface.

Phosphatidylserine-exposing erythrocytes are identified by macrophages which engulf and

degrade the eryptotic cells.

Objectives: The study has been performed to explore whether the administration of

amphotericin B or gum Arabic may modify the course of malaria and survival of

Plasmodium berghei -infected mice.

Materials and methods: Human erythrocytes were infected in vitro with Plasmodium

falciparum (strain BinH) in the absence and presence of amphotericin B or butyrate,

parasitaemia determined utilizing Syto16, cytosolic calcium estimated from fluo3

fluorescence intensity, phosphatidylserine exposure estimated from annexin V-binding and

cell volume from forward scatter in fluorescence activated cell sorting analysis. Mice were

infected with Plasmodium berghei ANKA by injecting parasitized murine erythrocytes (1

× 106) intraperitoneally. Where indicated amphotericin B (1.4 mg/kg b.w.) was

administered subcutaneously from the eighth day of infection for three successive days.

10% gum Arabic dissolved in tap water started from the day of infection for the other

group of mice. Preparations of gum Arabic were refreshed every day during the treatment.

Results: Amphotericin B during in vitro infection of human erythrocytes with P.

falciparum, increased phosphatidylserine exposure, decreased forward scatter and

significantly increases cytosolic calcium activity. Amphotericin B did not significantly

alter intraerythrocytic DNA/RNA amplification only at 1 µM but significantly (≥0.01 µM)

decreased in vitro parasitemia. Erythrocytes from amphotericin B treated mice were more

rapidly cleared from circulating blood than non-treated erythrocytes. Moreover,

parasitemia in P. berghei infected mice was significantly decreased (from 50.61% to

39.36% of circulating erythrocytes 20 days after infection) and mouse survival

VIII  

significantly enhanced (from 0% to 80% 27 days after infection) upon administration of

amphotericin B (1.4 mg/kg b. w) subcutaneously from the eighth day of infection.

In the other in vitro part of this study butyrate which is one of the important gum Arabic

fermentation products, significantly (≥0.5 mM) decreased the proliferation of P.

falciparum. On the other hand it influences intraerythrocytic DNA/RNA amplification only

at 10 mM concentration. Butyrate was significantly increased phosphatidylserine exposure,

decreased forward scatter and significantly increases cytosolic calcium activity. Most

importantly, gum Arabic treatment (10% in drinking water from the day of infection)

resulted in a significant decrease of parasitemia (from 43.6 ± 4.39 and 58.08) and increased

the survival of P. berghei-infected mice (70% of the gum Arabic-treated animals survived

the infection for more than 26 days after infection).

Conclusions: In conclusion amphotericin B and gum Arabic stimulate the erythrocytic

machinery responsible for the eryptosis following infection with Plasmodium. The

acceleration of eryptosis precedes the full intraerythrocytic maturation of the pathogen,

thus prevents the further lethal course of the disease and fosters host survival during

malaria. The revelations defend that the stimulation of eryptosis in infected erythrocytes is

a host dependent mechanism to combat against infection. The experimental results

indicated that the stimulation of eryptosis in infected erythrocytes is not only a host

dependent defense mechanism that can be used as a novel approach to prevent the chances

of resistance in plasmodia.

IX  

ملخص األطروحة

ة هذه الدراس. في وفاة أآثر من مليون انسان سنويا و تتسبب المالريا هي واحدة من أآثر األمراض المدمرة: خلفية

المخصصة الآتشاف أدوية جديدة من أجل منع مسببات المرض المقاومة لألدوية ذات الصلة مع الترآيز على تحديد

هذه األساليب المبتكرة . واستهداف عوامل أساسية في جسم العائل تتعلق بدخول مسببات المرض وبقائها وانتشارها

و تعرف تعرف الخاليا البالعة في الطحال عليها  ء المصابةتنطوي على تحريض الوفاة االنتحارية لكريات الدم الحمرا

 الوفاة االنتحارية لكريات الدم الحمراء وتتميز. من أجل التخلص من مسببات المرض ومنع المزيد من مسار المرض

 يلسيرينجزيئات الفوسفاتيد و بعثرة الدهون الفوسفورية لغشاء الخلية مع عرض بانكماش الخلية ، وانتفاخ غشاء الخلية

على سطح الخلية مما يؤدي إلى سهولة التعرف عليها بواسطة الخاليا البالعة و التي بدورها تقوم بإلتهام خاليا الدم

.الحمراء المصابة و تكسيرها

تهدف هذه الدراسة الستكشاف ما اذا آان إعطاء دواء االمفوتريسين ب أو الصمغ العربي قد يعدل مسار : األهداف

.اء المتصورة البرجيه في الفئران المصابةالمالريا وبق

) المتصورة المنجلية(تمت إصابة الكريات الحمراء البشرية في المختبر بالمتصورة المنجلية : المواد و األساليب

في غياب و وجود االمفوتريسين ب أو الزبدات ، تم تحديد مدى إنتشار المتصورات في الدم بإستخدام ) BinHساللة (

و تم تقدير ترآيز الكالسيوم في السيتوبالزم عن طريق قياس شدة اإلستضاءة الفلورية ، أما عرض , 16سايتو

الفوسفاتيديلسيرين فقد تم تقديره عن طريق إرتباط بروتين األنكسين الخامس و حجم الخلية عن طريق البعثرة األمامية

الفيئران بالمتصورة البرجيه أنكا عن طريق حقن تمت إصابة . في فرز التحليلي للخاليا المنشط باإلضاءة الفلورية

1.4(في الفيئران المخصصة للدراسة االمفوتريسين تم حقنه ). 106* 1(التجويف البريتوني بخاليا حمراء مصابة

و في المجموعة األخرى . تحت الجلد من اليوم الثامن من العدوى لمدة ثالثة أيام متتالية) آغم من وزن الجسم/ ملغم

٪ من الصمغ العربي المذاب في مياه الحنفية اعتبارا من اليوم األول من التجربة 10الفيئران المصابة تم تقديم من

.مع تجديد مستحضر الصمغ العربي آل يوم خالل فترة العالج

زيادة في في الكريات الحمراء البشرية في المختبر المصابة بالمتصورة المنجلية أدى االمفوتريسين ب إلى: النتائج

عرض الفوسفاتيديلسيرين على سطح الخاليا و نقصان في التبعثر األمامي و زيادة معتبرة في نشاط الكالسيوم داخل

لم يؤدي االمفوتريسين ب إلى تغيير معتبر في نسبة األحماض النووية دنا و رنا في داخل آريات الدم . السيتوبالزم

. إنتشار الطفيل داخل الكريات الحمراء الحمراء إال أنه أدى إلى نقصان معتبر في

الكريات الحمراء من الفئران المعالجة باالمفوتريسين ب تمت إزالتها من الدورة الدموية بسرعة أآبر مقارنة بالكريات

و إضافة إلى ذلك انخفض عدد الكريات المصابة . الحمراء في الفئران التي لم تتم معالجتها باالمفوتريسين ب

٪ من الكريات الحمراء المنتشرة 39.36٪ الى 50.61من (برجيه في الفئران المصابة بشكل ملحوظ بالمتصورة

يوما بعد 27٪ 80٪ إلى 0من (آما زادت نسبة بقاء الفئران على قيد الحياة بدرجة آبيرة ) يوما بعد اإلصابة 20

 . ثامن من العدوىفي حالة معالجتها باالمفوتريسين ب تحت الجلد من اليوم ال) اإلصابة

X  

0.5≥ (في جانب آخر من الدراسة المختبرية أدت الزبدات التي هي واحدة من اهم منتجات تخمير الصمغ العربي

من ناحية أخرى فإنها أثرت على الحمض النووي داخل . إلى إنخفاض ملحوظ في تكاثر المتصورة المنجلية) ملم

آما أدت الزبدات إلى زيادة . مليمول 10فقط في ترآيز / الريبي الكريات المصابة و ذلك بتضخيم الحمض النووي

آبيرة في عرض الفوسفاتيديلسيرين و انخفاض التبعثر إلى األمام وإلى زيادة آبيرة في نشاط الكالسيوم في

في ) ٪ في مياه الشرب اعتبارا من اليوم للعدوى 10(األهم من ذلك ، أدت المعالجة بالصمغ العربي . السيتوبالزم

وزيادة في نسبة بقاء الفئران المصابة ) 58.08و 4.39± 43.6من (تناقص آبير في عدد الخاليا المصابة

).يوما بعد اإلصابة 21٪ ٪ 100الى 0من (بالمتصورة برجيه على قيد الحياة

تخلص هذه الدراسة إلى أن االمفوتريسين ب و الصمغ العربي يقومان بتحفيز اآلليات المسؤولة عن إنتحار : الخالصة

و هذا اإلزدهار في إنتحار الخاليا الحمراء يسبق نضوج مسببات . لحمراء بعد العدوى مع المنجليةآريات الدم ا

يدافع هذا اإلآتشاف عن أن . لمضيف أثناء المالرياالمرض، ويمنع بالتالي المسار القاتل من هذا المرض وتعزز بقاء ا

النتائج التجريبية بجانب أنها تبرر . تنشيط إنتحارالكريات الحمراء المصابة هي آلية يعتمدها المضيف لمكافحة العدوى

ومة في إنتحار الكريات الحمراء آآلية للحد من إنتشار المرض فإنها ايضًا تعزز نهجا جديدا للحيلولة دون فرص للمقا

 .المتصورات

XI  

List of figures:

Figure. 1. 1. Life cycle of the malaria parasite P. falciparum............................................................5

Figure.1. 2: Schematic representation of apoptotic events..............................................................17

Figure.3. 3: In vitro growth of Plasmodium falciparum in human erythrocytes as a function of

amphotericin B concentration...............................................................................................53

Figure.3. 4: DNA amplification of Plasmodium falciparum in human infected

erythrocytes as a function of amphotericin B concentration................................................54

Figure.3. 5: Effect of amphotericin B on erythrocyte membrane integrity......................................55

Figure.3. 6: Effect of amphotericin B and ionomycin cytosolic fluo-3 fluorescence

intensity in non-infected erythrocytes..................................................................................56

Figure.3. 7: Effect of amphotericin B on cytosolic Ca2+ concentration in

non-infected erythrocytes.....................................................................................................57

Figure.3. 8 A & B: Annexin binding of non-infected and Plasmodium falciparum

infected erythrocytes from control and amphotericin B treated samples.............................58

Figure.3. 9 A & B: Effects of amphotericin B on forward scatter of Plasmodium

falciparum infected and noninfected erythrocytes................................................................59

Figure.3. 10: Effect of amphotericin B treatment on parasitemia

of Plasmodium berghei infected mice..................................................................................60

Figure.3. 11: Effect of amphotericin B on parasitemia of P. berghei infected mice........................61

Figure.3. 12 A & B: In vivo clearance of fluorescence labelled erythrocytes from

circulating blood...................................................................................................................62

Figure.3. 13: Effect of amphotericin B on survival of P. berghei infected mice.............................63

Figure.3. 14: In vitro growth of P. falciparum in human erythrocytes

as a function of butyrate concentration.................................................................................64

Figure.3. 15: Intraerythrocytic DNA amplification of P. falciparum in human

erythrocytes as a function of butyrate concentration............................................................65

Figure.3. 16: Effect of butyrate on cytosolic Ca2+ activity in non-infected

erythrocytes..........................................................................................................................66

Figure.3. 17: Effects of butyrate on phosphatidylserine exposure of infected

and non-infected erythrocytes..............................................................................................67

Figure.3. 18 A & B: Effects of butyrate on forward scatter of infected

and non-infected erythrocytes....................................................................................................68

Figure.3. 19: Parasitemia and survival of Plasmodium berghei infected mice................................69

Figure.3. 20: Effect of gum Arabic treatment on survival of

Plasmodium berghei infected mice......................................................................................70

1 

 

1. Introduction

1. 1. Malaria

Malaria is one of the most prevalent parasitic infections in the world and certainly the most

detrimental. Each year, over one million people die from the disease, with the vast majority

of the deaths in children under five years old in Sub-Saharan Africa (1). In addition to the

overwhelming death toll, over 213 million malarial “attacks” lead to more than 800 million

days of illness in Africa annually (2, 3) The spread of drug-resistant malaria parasites

threatens to compound the problem even more. The situation is so dire that economists

have determined that malaria is a definitive cause of poverty in many afflicted regions (4).

Furthermore, in malaria-endemic regions, the effect of the disease is also manifested by its

lasting influence on human genetics, resulting in the preservation of potentially harmful

variants of human genes (i.e., sickle cell trait), largely because of their advantage in

heterozygotes protected from severe, complicated, and fatal malaria (1).

1. 1. 1. History

The malaria-like intermittent fevers and other symptoms such as enlarged spleen, periods

of high fever, headaches, shaking chills, and weakness were already known more than

3000 years ago in Chinese, Assyrian, Indian, and Egyptian manuscripts. Hippocrates (460-

370 B.C.) in Greece is credited with having accurately described the clinical symptoms in

his medical writings (5). Malaria was widespread in Europe during the middle ages, but

malaria tropica was endemic only in the warmer South Europe. The name is derived from

the Italian word mala aria (Latin: malus = bad, aeris = air) because the disease often

appeared near swamps with their typical odor (6) It was thought that “miasma”, which

literally means “bad air”, caused the disease (7). The “miasma” theory was proved wrong

in the 19th century. In 1880 Charles Louis and Alphonse Laveran identified the parasite in

its blood stage in the act of exflagellation under the microscope (8). In 1897 Sir Ronald

Ross, whose teacher was Patrick Manson, who discovered the mosquito transmission of

filariasis, demonstrated the transmission of malaria via mosquitoes (9). However, it was

only in 1948 that Short and Garnham described the liver stage of primate and human

malaria Plasmodium vivax (P. vivax) (5). Finally, in 1966 the liver stage of P. falciparum

was discovered (10).

2 

 

1. 1. 2. Epidemiology of malaria

The "many epidemiologies" of malaria have been characterized by degrees of endemicity

(11). Malaria is described as endemic when there is a constant incidence of cases over a

period of many successive years. At the other extreme malaria transmission may be

epidemic when there is a periodic or occasional increase in the incidence of cases. A more

general classification into stable and unstable malaria has been introduced. Stable malaria

refers to high transmission without any marked fluctuations over years, although seasonal

fluctuations may exist. Unstable malaria describes transmission that varies from year to

year with frequent epidemics. The former situation is characterized by high degrees of

collective immunity whereas the latter is not. These terms describe extremes of a wide

range of situations (12).

Malaria is transmitted primarily by the bite of infected female Anopheline mosquitoes. The

source of infection can be either a sick person or an otherwise asymptomatic carrier of the

parasite. Malaria can also be transmitted congenitally or by inoculation of infected blood.

Anophelines feed at night and their breeding sites are primarily in rural areas. The greatest

risk of malaria infection is therefore from dusk to dawn in rural areas. In many malaria-

endemic areas, there is little or no risk in urban areas. However, urban transmission is

common in some parts of the world, especially Africa (13, 14).

The incidence of severe clinical manifestations varies seasonally within endemic areas and

it is not possible at present to predict which asymptomatic individual will develop severe

disease (14). The epidemiological profile and clinical pattern of severe malaria in Africa

has been shown to vary according to the intensity of exposure in persons living in rural

areas with different levels of transmission (15). The degree of endemicity varies between

countries and even between different areas in the same country. In Sub-Saharan Africa

cerebral malaria and severe malarial anaemia are the leading causes of mortality (16).

The impact of global climate change poses an obvious threat to human health. The insect-

vectors of Plasmodium spp. thrive in warm climates of tropical regions. Global warming

leading to increased temperature in temperate areas, could provide a habitat suitable for the

increased distribution of Anopheline vectors. Whether the potential increase in vector

populations will lead to a concomitant increase in malaria transmission is not clear (17).

3 

 

Malaria is unstable in the semi-arid savannah of central and northern Sudan and the great

majority of infective bites take place immediately prior to the seasonal peak in number of

malaria cases in September and October (18). Malaria morbidity and mortality, with

special reference to central Sudan, affects all age groups (19). In Sudan it has been found

that, despite variation between districts, urbanization tends to lead to reduced human

malaria transmission (20).

1. 1. 3. Clinical manifestations

The most characteristic symptom of malaria is fever. Other common symptoms include

chills, headache, myalgia, nausea, and vomiting. Diarrhea, abdominal pain, and cough are

occasionally seen. As the disease progresses, some patients may develop the classic

malaria paroxysm with bouts of illness alternating with symptom free periods. The malaria

paroxysm comprises three successive stages. The first is a 15 to 60 minute cold stage

characterized by shivering and a feeling of cold. Next, the 2 to 6 hour hot stage, in which

there is fever, sometimes reaching 41°C, flushed, dry skin, and often headache, nausea, and

vomiting. Finally, there is the 2 to 4 hour sweating stage during which the fever drops

rapidly and the patient sweats. In all types of malaria the periodic febrile response is

caused by rupture of mature schizonts. In P. vivax and P. ovale malaria, a brood of

schizonts matures every 48 hr, so the periodicity of fever is tertian ("tertian malaria"),

whereas in P. malariae disease, fever occurs every 72 hours ("quartan malaria").

The fever in P. falciparum malaria may occur every 48 hr, but is usually irregular, showing

no distinct periodicity. These classic fever patterns are usually not seen early in the course

of malaria, and therefore the absence of periodic, synchronized fevers does not rule out a

diagnosis of malaria (21).

A variety of laboratory abnormalities may be seen in a case of uncomplicated malaria.

These include normochromic, normocytic anemia, thrombocytopenia, leukocytosis or

leukopenia, hypoglycemia, hyponatremia, elevated liver and renal function tests,

proteinuria. Patients with complicated malaria may occasionally show evidence of massive

intravascular hemolysis with hemoglobinemia and hemoglobinuria.

If the diagnosis of malaria is missed or delayed, especially with P. falciparum infection,

potentially fatal complicated malaria may develop. The most frequent and serious

complications of malaria are cerebral malaria and severe anemia. Other complications

4 

 

include: hyperparasitemia (more than 3 to 5% percent of the erythrocytes parasitized);

severe hypoglycemia; lactic acidosis; shock; pulmonary, cardiac, hepatic, or renal

dysfunction; seizures; spontaneous bleeding; or massive diarrhea or vomiting. These

manifestations are usually associated with poor prognosis (21).

1. 1. 4. The parasites

There is approximately 400 species of Anopheles throughout the world; about 60 are

malaria vectors under natural conditions, 30 of which are of major importance. Malaria is

transmitted by the bite of an infected female Anopheles mosquito. Malaria parasites are

eukaryotic single-celled microorganisms that belong to the genus Plasmodium. More than

100 species of Plasmodium can infect various animal species such as reptiles, birds and

various mammals, but only four species of parasite can infect humans under natural

conditions: P falciparum, P.vivax, P. ovale and P. malariae. These four species differ in

their morphology, antigenicity, pathogenicity, drug sensitivity and geographical

distribution. P.falciparum is species responsible for the severe complications of malaria

and is the principal cause of malaria deaths in young children in Africa (22, 23). The

widely worldwide distributed P. vivax is rarely fatal, while P.malariae has a relatively low

frequency. On the other hand, the least common P. ovale is restricted to West Africa.

Although both P. falciparum and P.vivax can cause anaemia, mild anaemia is more

common in P.vivax infections whereas severe anemia is usually caused by P. falciparum..

P. ovale and P. vivax have dormant liver stages named hypnozoites that may remain in this

organ for weeks to many years before the onset of a new round of pre-erythrocytic

schizogony, resulting in relapses of malaria infection. In some cases P. malariae can

produce long-lasting blood-stage infections, which, if left untreated, can persist

asymptomatically in the human host for periods extending into several decades (24).

1. 1. 5. Life cycle of malaria parasites

The life cycle of malaria parasites is extremely complex and requires specialized protein

expression for survival in both the invertebrate and vertebrate hosts. These proteins are

required for the invasion of a variety of cell types and for the evasion of host immune

responses. Once injected into the human host, P. falciparum and P. malariae sporozoites

trigger immediate schizogony, whereas P. ovale and P. vivax sporozoites may either

trigger immediate schizogony or lead to delayed schizogony as they pass through the

5 

 

hypnozoite stage. The life cycle of the malaria parasite can be divided into several stages

as shown in figure 1. 1 below, starting with sporozoite entry into the bloodstream (21).

Fig. 1. 1. Life cycle of the malaria parasite P. falciparum (24).

1. 1. 5. 1. Tissue schizogony (pre-erythrocytic schizogony)

Infective sporozoites from the salivary gland of the Anopheles mosquito are injected into

the human host along with anticoagulant-containing saliva to ensure an even-flowing blood

meal. It was thought that sporozoites move rapidly away from the site of injection, but a

recent study using the rodent parasite (25) species Plasmodium yoelii as a model system

indicates that, at least in this case, the majority of infective sporozoites remain at the

injection site for hours, with only slow release into the circulation (26). Once in the human

bloodstream, P. falciparum sporozoites reach the liver and penetrate the liver cells

(hepatocytes) where they remain for 9–16 days and undergo asexual replication known as

exo-erythrocytic schizogony. The mechanism of targeting and invading the hepatocytes is

not yet well understood, but studies have shown that sporozoite migration through several

hepatocytes in the mammalian host is essential for completion of the life cycle. Each

sporozoite gives rise to tens of thousands of merozoites inside the hepatocytes, upon

6 

 

release these merozoites invades red cells to undergo successive waves of schizogeny

releasing more merozoites to reinvade new red blood cells (RBCs). The time taken to

complete the liver stage of the parasite life cycle, called the prepatent period, varies,

depending on the infecting species; (8–25 days for P. falciparum, 8–27 days for P. vivax,

9–17 days for P. ovale and 15–30 days for P. malariae) (27).

1. 1. 5. 2. Erythrocytic schizogony

Merozoites enter erythrocytes by a complex invasion process, which can be divided into

four phases: (a) initial recognition and reversible attachment of the merozoite to the

erythrocyte membrane; (b) reorientation and junction formation between the apical end of

the merozoite (irreversible attachment) and the release of substances from the rhoptry and

microneme organelles, leading to formation of the parasitophorous vacuole; (c) movement

of the junction and invagination of the erythrocyte membrane around the merozoite

accompanied by removal of the merozoite's surface coat; and finally (d) resealing of the

parasitophorous vacuole and erythrocyte membranes after completion of merozoite

invasion. Asexual division starts inside the erythrocyte and the parasites develop through

different stages therein. The early trophozoite is often referred to as the 'ring stage',

because of its characteristic morphology. Trophozoite enlargement is accompanied by

highly active metabolism, which includes glycolysis of large amounts of imported glucose,

the ingestion of host cytoplasm and the proteolysis of hemoglobin into constituent amino

acids. Malaria parasites cannot degrade the by-product heme which is potentially toxic to

the parasite. Therefore, during hemoglobin degradation, most of the liberated heme is

polymerized into hemozoin (malaria pigment), a crystalline substance that is stored within

the food vacuoles (28).

The end of this trophic stage is marked by multiple rounds of nuclear division without

cytokinesis resulting in the formation of schizonts. Each mature schizont contains around

20 merozoites which are released upon lysis of the RBC to reinvade further uninfected

RBCs. This release coincides with the sharp increases in body temperature during the

progression of the disease. This repetitive intraerythrocytic cycle of invasion–

multiplication–release–reinvasion continues, taking about 48 h in P. falciparum, P. ovale

and P. vivax infections and 72 h in P. malariae infection. It usually occurs quite

synchronously and the merozoites are released at approximately the same time of the day.

The contents of the infected RBC that are released upon its lysis stimulate the production

7 

 

of tumour necrosis factor (TNF) and other cytokines, which are responsible for the

characteristic clinical manifestations of the disease (24).

A small proportion of the merozoites in the RBCs eventually differentiate to produce

micro- and macrogametocytes (male and female, respectively), which have no further

activity within the human host. These gametocytes are essential for transmitting the

infection to new hosts after undergoing an sporogenic cycle in female Anopheles

mosquitoes. Normally, variable numbers of cycles of asexual erythrocytic schizogony

occur before any gametocytes are produced. In P. falciparum, erythrocytic schizogony

takes 48 h and gametocytogenesis takes 10–12 days. Gametocytes appear on the fifth day

of primary attack in P. vivax and P. ovale infections, and thereafter become more

numerous; they appear at anything from 5 to 23 days after a primary attack by P. malariae

(24).

1. 1. 5. 3. Sexual phase in the mosquito (Sporogony)

A mosquito taking a blood meal on an infected individual may ingest these gametocytes

into its midgut, where macrogametocytes form macrogametes and exflagellation of

microgametocytes produces microgametes. These gametes fuse, undergo fertilization and

form a zygote. This transforms into an ookinete, which penetrates the wall of a cell in the

midgut and develops into an oocyst. In a recent study (29), it has been shown that gamete

surface antigen Pfs230 mediates human RBC binding to exflagellating male parasites to

form clusters termed exflagellation centers, from which individual motile microgametes

are released. This protein thus plays an important role in subsequent oocyst development,

which is a critical step in malaria transmission. Sporogony within the oocyst produces

many sporozoites and when the oocyst ruptures, they migrate to the salivary glands for

onward transmission into another vertebrate host. This infective “sporozoite” form of the

parasite reaches the salivary glands after 10–18 days and thereafter the mosquito becomes

infective for 1–2 months (30).

1. 1. 6. Malaria immunity

There are different types of protective immunity against malaria, one type is the clinical

immunity which reduces the intensity of clinical symptoms and the risk of death from

malaria, the other type is the antiparasitic immunity which directly reduces the number of

parasites in the infected individual (31). In the case of P .falciparum malaria, it is possible

8 

 

that a certain degree of immunity to some aspects of severe disease may be attained after

only one or two infections (32). However, effective antiparasitic immunity is only attained

after several frequent infections (33).

1. 1. 6. 1. Innate immunity

Some individuals are either naturally resistant to malaria infection or less likely to develop

a severe form of the disease. Innate resistance to infection is generally only a partial

resistance, and may be linked to the fact that malaria parasites find it harder to invade

certain types of human erythrocytes. Also, it could be due to the fact that certain host

erythrocytes have a reduced ability to sustain the growth of parasites (33).

1. 1. 6. 2. Acquired immunity

In endemic areas, most adults have a degree of immunity that controls parasite replication

but not sterile immunity which eliminates parasite from blood. Acquired immunity to

malaria develops after repeated exposure to the parasite (34).

1. 1. 6. 3. Strain specific immunity

The long time required to develop protective immunity against malaria may be explained

by poor immunogenicity of the parasite and/or that immune responses are strain specific. A

large number of infections would then be required to encounter the whole local repertoire

of different antigens. Early studies of induced malaria indicated that malaria immunity is

strain specific (35, 36).

1. 1. 7. Malaria diagnosis

The standard method for detection of Plasmodium in human blood is by microscopical

examination of Romanovsky-stained thick and thin blood films. This technique can, when

used in optimal conditions by a competent microscopist, detect a parasitaemia as low as

0.001% (10-40 parasites per µl of blood), but it is a time-consuming technique for the

detection of scanty parasites and often difficult to use accurately to identify mixed

infections. Several alternative diagnostic methods have been developed in order to reduce

the time spent examining slides or to enable fairly competent personnel to achieve equally

reliable results. These include:

Antigen detection: Various test kits are available to detect antigens derived from malaria

parasites. Such immunochromatographic tests most often use a dipstick or cassette format,

9 

 

and provide results in 2-15 minutes. These "Rapid Diagnostic Tests" (RDTs) offer a useful

alternative to microscopy in situations where reliable microscopic diagnosis is not

available. Malaria RDTs are currently used in some clinical settings and programs.

However, before malaria RDTs can be widely adopted, several issues remain to be

addressed, including improving their accuracy; lowering their cost; and ensuring their

adequate performance under adverse field conditions (37).

Molecular diagnosis: Parasite nucleic acids are detected using polymerase chain reaction

(PCR). This technique is more accurate than microscopy. However, it is expensive, and

requires a specialized laboratory even though technical advances will likely result in field-

operated PCR machines (37).

Serology: Serology detects antibodies against malaria parasites, using either indirect

immunofluorescence (IFA) or enzyme-linked immunosorbent assay (ELISA). Serology

does not detect current infection but rather measures past experience (37).

1. 1. 8. Malaria chemotherapy

Chemotherapy is important in alleviating the suffering and reducing the mortality caused

by the disease. The action of antimalarial drugs ties in their effect on the metabolic process

or the vital structure of the parasite which lead to the disruption of its activity and then to

parasite death. However the efficacy of the drug depends also on the species and strain of

the parasite and its sensitivity towards a given drug, as well as the state of host immunity

(38).

Patients successfully treated with antimalarial drugs may thus be healthy but may remain

infective for up to two months until the P. falciparum gametocytes die off naturally, or

until another drug such as primaquine is given to eliminate the mature gametocytes (39).

1. 1. 8. 1. 4, 8-aminoquinoline drugs

Such as chloroquine, amodiaquine, and primaquine. Chloroquine is a 4-aminoquinoline

that has marked and rapid schizonticidal activity against all infections of P. malariae and

P. ovale and against chloroquine-sensitive infections of P. falciparum and P. vivax. It is

also gametocytocidal against P. vivax, P, malariae and P. ovale as well as immature

gametocytes of P. falciparum. It is not active against hepatic stages, and should therefore

be used with primaquine to effect radical cure of P. vivax and P. ovale. It was the most

important antimalarial for more than 40 years because of its activity against the four

10 

 

plasmodia that infect human (39). In addition, chloroquine is known to be safe for children

and pregnant women (38, 40). However, the efficacy of chloroquine has been severely

compromised by the spread of chloroquine-resistant strains of P. falciparum, which are

now prevalent in Southeast Asia, South and Central America, and sub-Saharan Africa (41)

where drug-resistance led to increased morbidity and mortality of malaria and forced the

use of alternative drugs (42).

1. 1. 8. 2. Antifolate drugs

The only useful combinations of antifolate drugs for the treatment of malaria are

synergistic mixtures that act against the parasite-specific enzymes, dihydropteroate

synthetase and dihydrofolate reductase. Available combinations include the sulfa and

pyrimethamine combinations sulfadoxine-pyrimethamine and sulfalene-pyrimethamine,

the former being more widely available (43).

Antifolate drugs such as pyrimethamine and proguanil are slow blood schizonticides but

they have the advantage that they are effective in smaller doses, safe and free from side

effects. The ability of sulfa (sulfonamides and sulfones) to show potentiating effect with

proguanil or pyrimethamine (44) suggested that combination of these two groups of drugs

would not only be more effective antimalarial agent, but would also delay, if not avoid, the

development of resistance. The most commonly used combination is that of sulfonamide,

sulfadoxine, and pyrimethamine (fansidar), which has been particularly effective in areas

such as Africa where chloroquine resistance is now wide spreading (45). However,

resistance to the pyrimethamine sulfadoxine combination became widespread and the

successive failure of these two drugs in Africa was described in 1998 as malaria disaster

(45).

1. 1. 8. 3. Quinine

It is normally effective against P.falciparum infections that are resistant to chloroquine and

sulfa drug-pyrimethamine combinations. Decreasing sensitivity to quinine has been

detected in areas of South-East Asia where it has been extensively used for malaria

therapy. This has occurred particularly when therapy was given in an unsupervised and

ambulatory setting with regimens longer than 3 days. In these settings, patient adherence to

therapy is low, leading to incomplete treatment; this may have led to the selection of

resistant parasites (46).

11 

 

1. 1. 8. 4. Artemisinin and its derivatives

Artemisinin is poorly soluble in oils or water but the parent compound has yielded

dihydroartemisinin, the oil-soluble derivatives artemether and arteether, and the more

water-soluble derivatives sodium artesunate and artetinic acid. These derivatives have

more potent blood schizonticidal activity than the parent compound and are the most

rapidly effective antimalarial drugs known. They are used for the treatment of severe and

uncomplicated malaria (47). They are not hypnozoiticidal but gametocytocidal activity has

been observed. These compounds are not recommended for use in the treatment of malaria

due to P. vivax, P. malariae or P. ovale since other effective antimalarial drugs are

available for this purpose (39).

1. 1. 8. 5. Combination therapy to combat the spread of drug resistance

Due to the emergence and rapid spread of resistance to almost all available antimalarial

drugs, much of interest has been focused upon the use of drugs in combination. The

underlying science behind the therapeutic effect of the combinations that include an

artemisinin derivative is that the artemisinin rapidly kills most of the parasites and then

those that remain are killed by the companion drug (39).

Combination therapy (CT) is defined as "the simultaneous use of two or more antimalarial

drugs with different biochemical targets in the parasite or in tissue hosting the parasite,

which are synergistic, additive or complementary in their effect". Using this definition, two

or more drugs which have the same biochemical target in the parasite such as sulfadoxine,

and pyrimethamine, chlorproguanil-dapsone, atovaquone-proguanil are not considered CT.

Similarly, an antimalarial which is co-administered with a non-antimalarial drug which

enhances its action is also not classified as CT. Suffice to say that CT therefore can be

either fixed combinations, where all components are coformulated in a single

tablet/capsule or free combinations where the components are in separate tablets/capsules

but are coadministered (39).

In sub-Saharan Africa, the first two combinations considered in a larger scale were

pyrimethamine/sulfadoxine plus chloroquine and pyrimethamine/sulfadoxine plus

amodiaquine (39). Recently interest has been directed towards the use of drugs in

combination with artesunate in purpose of delaying the emergence of resistance (48). The

12 

 

antifolate product proguanil has been formulated together with a new type of inhibitor,

atovaquone, to yield malar-one, recently licensed for clinical use (49).

1. 2. Apoptosis

Apoptosis is the process of programmed cell death (PCD), that may occurs normally

during development and aging as a homeostatic mechanism to maintain cell populations in

tissues. Apoptosis also occurs as a defence mechanism such as in immune reactions or

when cells are damaged by disease or noxious agents. Although there are a wide variety of

stimuli and conditions, both physiological and pathological, that can trigger apoptosis, not

all cells will necessarily die in response to the same stimulus. Irradiation or drugs used for

cancer chemotherapy results in DNA damage in some cells, which can lead to apoptotic

death through a p53-dependent pathway. Some hormones, such as corticosteroids, may

lead to apoptotic death in some cells (e.g., thymocytes) although other cells are unaffected

or even stimulated. Apoptosis is a coordinated and often energy-dependent process that

involves the activation of a group of cysteine proteases called “caspases” and a complex

cascade of events that link the initiating stimuli to the final demise of the cell (50).

1. 2. 1. Morphology of apoptosis

Light and electron microscopy have identified the various morphological changes that

occur during apoptosis (51). During the early process of apoptosis, cell shrinkage and

pyknosis are visible by light microscopy (52). With cell shrinkage, the cells are smaller in

size, the cytoplasm is dense and the organelles are more tightly packed. Pyknosis is the

result of chromatin condensation and this is the most characteristic feature of apoptosis. On

histological examination with hematoxylin and eosin stain, the apoptotic cell appears as a

round or oval mass with dark eosinophilic cytoplasm and dense purple nuclear chromatin

fragments. Early during the chromatin condensation phase, the electron-dense nuclear

material characteristically aggregates peripherally under the nuclear membrane although

there can also be uniformly dense nuclei. Blebbing which is an extensive irregular bulging

of the plasma membrane detached from the cytoskeleton occurs followed by karyorrhexis

and separation of cell fragments into apoptotic bodies during a process called “budding.”

Apoptotic bodies consist of cytoplasm with tightly packed organelles with or without a

nuclear fragment. The organelle integrity is still maintained and all of this is enclosed

within an intact plasma membrane. These bodies are subsequently phagocytosed by

13 

 

macrophages, parenchymal cells, or neoplastic cells and degraded within phagolysosomes.

Macrophages that engulf and digest apoptotic cells are called “tingible body macrophages”

and are frequently found within the reactive germinal centers of lymphoid follicles or

occasionally within the thymic cortex. The tingible bodies are the bits of nuclear debris

from the apoptotic cells. There is essentially no inflammatory reaction associated neither

with the process of apoptosis nor with the removal of apoptotic cells because: (1) apoptotic

cells do not release their cellular constituents into the surrounding interstitial tissue; (2)

they are quickly phagocytosed by surrounding cells thus likely preventing secondary

necrosis; and, (3) the engulfing cells do not produce anti-inflammatory cytokines (53, 54).

1. 2. 2. Classification of cell death: apoptosis and necrosis

Mechanisms of cell death varies with cell type, developmental stage, metabolism and the

kind of death signal. Apoptosis is a major form of programmed cell death, although there

are still other types of programmed cell death, especially in pathological situations

(autophagic death, cytoplasmic death, parapoptosis etc.). These modes of cell death are,

however, much less studied (55, 56).

The alternative to apoptotic cell death is necrosis, which is considered to be a toxic process

where the cell is a passive victim and follows an energy-independent mode of death. But

since necrosis refers to the degradative processes that occur after cell death, it is considered

to be an inappropriate term to describe a mechanism of cell death. Oncosis is therefore

used to describe a process that leads to necrosis with karyolysis and cell swelling whereas

apoptosis describes the process that involves cell shrinkage, pyknosis, and karyorrhexis.

Necrosis is an uncontrolled and passive process that usually affects large fields of cells

whereas apoptosis is controlled and energy-dependent and can affect individual or clusters

of cells. Necrotic cell injury is mediated by two main mechanisms; interference with the

energy supply of the cell and direct damage to cell membranes (57).

Although the mechanisms and morphologies of apoptosis and necrosis differ, there is

overlap between these two processes. Evidence indicates that necrosis and apoptosis

represent morphologic expressions of a shared biochemical network described as the

“apoptosis-necrosis continuum” (58). For example, two factors that convert an ongoing

apoptotic process into a necrotic process include a decrease in the availability of caspases

and intracellular ATP (58, 59). It is not always easy to distinguish apoptosis from necrosis

14 

 

by conventional histological examinations. Both processes can occur simultaneously

depending on factors such as the intensity and duration of the stimulus, the extent of ATP

depletion and the availability of caspases (58).

Some of the major morphological changes that occur with necrosis include cell swelling;

formation of cytoplasmic vacuoles; distended endoplasmic reticulum; formation of

cytoplasmic blebs; condensed, swollen or ruptured mitochondria; disaggregation and

detachment of ribosomes; disrupted organelle membranes; swollen and ruptured

lysosomes; and eventually disruption of the cell membrane (52, 57, 60). This loss of cell

membrane integrity results in the release of the cytoplasmic contents into the surrounding

tissue, sending chemotatic signals with eventual recruitment of inflammatory cells.

Because apoptotic cells do not release their cellular constituents into the surrounding

interstitial tissue and are quickly phagocytosed by macrophages or adjacent normal cells,

there is essentially no inflammatory reaction (53, 54). It is also important to note that

pyknosis and karyorrhexis are not exclusive to apoptosis and can be a part of the spectrum

of cytomorphological changes that occurs with necrosis (58).

1. 2. 3. Mechanisms of apoptosis

The mechanisms of apoptosis are highly complex and sophisticated, involving an energy-

dependent cascade of molecular events (Figure 1. 2). To date, research indicates that there

are two main apoptotic pathways: the extrinsic or death receptor pathway and the intrinsic

or mitochondrial pathway. However, there is now evidence that the two pathways are

linked and that molecules in one pathway can influence the other (61). There is an

additional pathway that involves T-cell mediated cytotoxicity and perforin-granzyme-

dependent killing of the cell. The perforin/granzyme pathway can induce apoptosis via

either granzyme B or granzyme A. The extrinsic, intrinsic, and granzyme B pathways

converge on the same terminal, or execution pathway. This pathway is initiated by the

cleavage of caspase-3 and results in DNA fragmentation, degradation of cytoskeletal and

nuclear proteins, cross-linking of proteins, formation of apoptotic bodies, expression of

ligands for phagocytic cell receptors and finally uptake by phagocytic cells. The granzyme

A pathway activates a parallel, caspase-independent cell death pathway via single stranded

DNA damage (62).

15 

 

1. 2. 4. Biochemical features of apoptosis

Apoptotic cells exhibit several biochemical changes such as protein cleavage, protein

cross-linking, DNA breakdown, and phagocytic recognition that together result in the

distinctive structural pathology. Cysteine-aspartic proteases (Caspases) are widely

expressed in an inactive proenzyme form in most cells and once activated can often

activate other procaspases, allowing initiation of a protease cascade. Some procaspases can

also aggregate and autoactivate. This proteolytic cascade, in which one caspase can

activate other caspases, amplifies the apoptotic signalling pathway and thus leads to rapid

cell death (63).

Caspases have proteolytic activity and are able to cleave proteins at aspartic acid residues,

although different caspases have different specificities involving recognition of

neighbouring amino acids. Once caspases are initially activated, there seems to be an

irreversible commitment towards cell death. Fourteen major caspases have been identified

and were broadly categorized into initiators (caspase-2,-8,-9,-10), effectors or executioners

(caspase-3,-6,-7) and inflammatory caspases (caspase-1,-4,-5) (64, 65). The other caspases

that have been identified include caspase-11 which regulates apoptosis and cytokine

secretion during septic shock, caspase-12 which mediates endoplasmic-specific apoptosis

and cytotoxicity by amyloid-β, caspase-13 which is suggested to be a bovine gene and

caspase-14 which is highly expressed in embryonic tissues but not in adult tissues (66-68).

Extensive protein cross-linking is another characteristic of apoptotic cells and is achieved

through the expression and activation of tissue transglutaminase (69). DNA breakdown by

Ca2+-and Mg2+-dependent endonucleases also occurs, resulting in DNA fragments of 180

to 200 base pairs. A characteristic “DNA ladder” can be visualized by agarose gel

electrophoresis with an ethidium bromide stain and ultraviolet illumination (70).

Another biochemical feature is the expression of cell surface markers that result in the

early phagocytic recognition of apoptotic cells by adjacent cells, permitting quick

phagocytosis with minimal compromise to the surrounding tissue. This is achieved by the

movement of the normal inward-facing phosphatidylserine of the cell’s lipid bilayer to

expression on the outer layers of the plasma membrane (71). Although externalization of

phosphatidylserine is a well-known recognition ligand for phagocytes on the surface of the

apoptotic cell, other proteins are also being exposed on the cell surface during apoptotic

16 

 

cell clearance. These include Annexin I and calreticulin. Calreticulin is a protein that binds

to an LDL-receptor related protein on the engulfing cell and is suggested to cooperate with

phosphatidylserine as a recognition signal (72). Annexin V is a recombinant

phosphatidylserine-binding protein that interacts strongly and specifically with

phosphatidylserine residues and can be used for the detection of apoptosis (73).

1. 2. 4. 1. Extrinsic pathway of apoptosis

The extrinsic signalling pathways that initiate apoptosis involve trans-membrane receptor-

mediated interactions. These involve death-receptors that are members of the tumour

necrosis factor (TNF) receptor gene superfamily (74). Members of the TNF receptor

family share similar cyteine-rich extracellular domains and have a cytoplasmic domain of

about 80 amino acids called the “death domain” (75). This death domain plays a critical

role in transmitting the death signal from the cell surface to the intracellular signalling

pathways. To date, the best-characterized ligands and corresponding death receptors

include fatty acid synthase ligand/ fatty acid synthase receptors (FasL/FasR), tumor

necrosis factor-α/ tumor necrosis factor receptor (TNF-α/TNFR1), apoptosis 3 ligand/death

receptors 3 (Apo3L/DR3), and apoptosis 2 ligand/death receptors 4 (Apo2L/DR4) (76-78).

The sequences of events that define the extrinsic phase of apoptosis are best characterized

with the FasL/FasR and TNF-α/TNFR1 models. In these models, there is clustering of

receptors and binding with the homologous trimeric ligand. Upon ligand binding,

cytplasmic adapter proteins are recruited which exhibit corresponding death domains that

bind with the receptors. The binding of Fas ligand to Fas receptor results in the binding of

the adapter protein, Fas-associated death domain (FADD) and the binding of TNF ligand to

TNF receptor results in the binding of the adapter protein TNFReceptor-associated death

domain (TRADD) with recruitment of FADD and receptor-interacting protein (RIP) (79).

FADD then associates with procaspase-8 via dimerization of the death effector domain. At

this point, a death-inducing signalling complex (DISC) is formed, resulting in the auto-

catalytic activation of procaspase-8 (80). Once caspase-8 is activated, the execution phase

of apoptosis is triggered (76).

17 

 

Fig. 1. 2: Schematic representation of apoptotic events. The two main pathways of apoptosis are extrinsic and intrinsic as well as a perforin/granzyme pathway. Each requires specific triggering signals to begin an energy-dependent cascade of molecular events. Each pathway activates its own initiator caspase (8, 9, 10) which in turn will activate the executioner caspase-3. However, granzyme A works in a caspase-independent fashion. The execution pathway results in characteristic cytomorphological features including cell shrinkage, chromatin condensation, formation of cytoplasmic blebs and apoptotic bodies and finally phagocytosis of the apoptotic bodies by adjacent parenchymal cells, neoplastic cells or macrophages (69).

1. 2. 4. 2. Perforin/granzyme pathway of apoptosis

T-cell are able to exert their cytotoxic effects on tumour cells and virus-infected cells via a

novel pathway that involves secretion of the trans-membrane pore-forming molecule

perforin with a subsequent exophytic release of cytoplasmic granules through the pore and

into the target cell (79). The serine proteases granzyme A and granzyme B are the most

important component within the granules (80). Granzyme B will cleave proteins at

aspartate residues and will therefore activate procaspase-10 and can cleave factors like

ICAD (Inhibitor of Caspase Activated DNAse) (81). Granzyme B also can utilize the

mitochondrial pathway for amplification of the death signal by specific cleavage of Bid

and induction of cytochrome c release (82).

18 

 

Granzyme A is also important in cytotoxic T cell induced apoptosis and it activates caspase

independent pathways. Once in the cell, granzyme A activates DNA nicking through

DNAse NM23-H1, a tumor suppressor gene product (83). This DNAse has an important

role in immune surveillance to prevent cancer through the induction of tumor cell

apoptosis. The nucleosome assembly protein SET normally inhibits the NM23-H1 gene.

Granzyme A protease cleaves the SET complex thus releasing inhibition of NM23-H1,

resulting in apoptotic DNA degradation. In addition to inhibiting NM23-H1, the SET

complex has important functions in chromatin structure and DNA repair. The proteins that

make up this complex (SET, Ape1, pp32, and HMG2) seem to work together to protect

chromatin and DNA structure. Therefore, inactivation of this complex by granzyme A is

likely to contribute to apoptosis by blocking the maintenance of DNA and chromatin

structure integrity (84).

1. 2. 4. 3. Intrinsic pathway of apoptosis

The intrinsic signalling pathway of apoptosis involves a diverse array of non-receptor-

mediated stimuli that produce intracellular signals which act directly on targets within the

cell and are mitochondrial-initiated events. The stimuli that initiate the intrinsic pathway

produce intracellular signals that may act in either a positive or negative fashion. Negative

signals involve the absence of certain growth factors, hormones and cytokines that can lead

to failure of suppression of death programs, thereby triggering apoptosis. In other words,

there is the withdrawal of factors, loss of apoptotic suppression, and subsequent activation

of apoptosis. Other stimuli that act in a positive fashion include, but are not limited to,

radiation, toxins, hypoxia, hyperthermia, viral infections, and free radicals (87).

All apoptotic stimuli cause changes in the inner mitochondrial membrane and result in an

opening of the mitochondrial permeability transition (MPT) pore, loss of the mitochondrial

trans-membrane potential and release of two main groups of normally sequestered pro-

apoptotic proteins from the intermembrane space into the cytosol (87). The first group

consists of cytochrome c, Second Mitochondria-derived Activator of Caspases/Direct IAP

Binding Protein with Low PI (Smac/DIABLO), and the serine protease HtrA2/Omi (88).

These proteins activate the caspase-dependent mitochondrial pathway. Cytochrome c binds

and activates Apaf-1 as well as procaspase-9, forming an “apoptosome” (89, 90). The

second group of pro-apoptotic proteins, include the apoptosis-inducing factor (AIF),

endonuclease G and caspase-activated Dnase (CAD) are released from the mitochondria

19 

 

during apoptosis, but this is a late event that occurs after the cell has been committed to

death. AIF translocates to the nucleus and causes DNA fragmentation into ~50–300 kb

pieces and condensation of peripheral nuclear chromatin (91).

The control and regulation of these apoptotic mitochondrial events occurs through

members of the B-cell lymphoma 2 (Bcl-2) family of proteins (92). The tumour suppressor

protein p53 has a critical role in regulation of the Bcl-2 family of proteins; however the

exact mechanisms have not yet been completely elucidated. The Bcl-2 family of proteins

governs mitochondrial membrane permeability and can be either pro-apoptotic or anti-

apoptotic. It is thought that the main mechanism of action of the Bcl-2 family of proteins is

the regulation of cytochrome c release from the mitochondria via alteration of

mitochondrial membrane permeability (93).

1. 2. 4. 4. Execution pathway of apoptosis

The extrinsic and intrinsic pathways both end at the point of the execution phase which is

the final pathway of apoptosis. It is the activation of the execution caspases that triggers

this phase of apoptosis. Execution caspases activate cytoplasmic endonuclease, which

degrades nuclear material, and proteases that degrade the nuclear and cytoskeletal proteins.

Caspase-3, 6 and 7 act as effectors or “executioner” caspases, cleaving various substrates

including cytokeratins, Poly (ADP-ribose) polymerase (PARP), the plasma membrane

cytoskeletal protein alpha fodrin, the nuclear mitotic apparatus protein (NuMA) and others,

that ultimately cause the morphological and biochemical changes seen in apoptotic cells

(94).

1. 2. 5. Physiologic apoptosis

The role of apoptosis in normal physiology is as significant as that of its counterpart,

mitosis. It demonstrates a complementary but opposite role to mitosis and cell proliferation

in the regulation of various cell populations. It is estimated that to maintain homeostasis in

the adult human body, around 10 billion cells are made each day just to balance those

dying by apoptosis. And that number can increase significantly when there is increased

apoptosis during normal development and aging or during disease (95).

Apoptosis is critically important during various developmental processes. Apoptosis is

necessary to rid the body of pathogen-invaded cells and is a vital component of wound

healing in that it is involved in the removal of inflammatory cells and the evolution of

20 

 

granulation tissue into scar tissue (96). Additionally, apoptosis is central to remodelling in

the adult, such as the follicular atresia of the postovulatory follicle and post-weaning

mammary gland involution (97, 98). Furthermore, as organisms grow older, some cells

begin to deteriorate at a faster rate and are eliminated via apoptosis. It is clear that

apoptosis has to be tightly regulated since too little or too much cell death may lead to

pathology, including developmental defects, autoimmune diseases, neurodegeneration or

cancer (99, 100).

1. 2. 6. Pathologic apoptosis

Abnormalities in cell death regulation can be a significant component of diseases such as

cancer, autoimmune lymphoproliferative syndrome, acquired immune deficiency

syndrome (AIDS), ischemia, and neurode-generative diseases such as Parkinson’s disease,

Alzheimer’s disease, Huntington’s disease, and Amyotrophic Lateral Sclerosis. Some

conditions feature insufficient apoptosis whereas others feature excessive apoptosis.

Cancer is an example where the normal mechanisms of cell cycle regulation are

dysfunctional, with either an over-proliferation of cells and/or decreased removal of cells

(95, 101).

Excessive apoptosis may also be a feature of some conditions such as autoimmune

diseases, neurodegenerative diseases, and ischemia-associated injury. Acquired immune

deficiency syndrome (AIDS) is an example of an autoimmune disease that results from

infection with the human immunodeficiency virus (HIV) (102).

1. 2. 7. Inhibition of apoptosis

There are many pathological conditions that exhibit excessive apoptosis

(neurodegenerative diseases, AIDS, ischemia, etc.) and thus may benefit from artificially

inhibiting apoptosis. However, understanding of the field evolves, the identification and

exploitation of new targets remains a considerable focus of attention (103). A short list of

potential methods of anti-apoptotic therapy includes stimulation of the inhibitors of

apoptosis proteins (IAP) family of proteins, caspase inhibition, PARP (poly [ADP-ribose]

polymerase) inhibition, stimulation of the protein kinase B (PKB) pathway, and inhibition

of Bcl-2 proteins. The inhibitor of apoptosis (IAP) family of proteins is perhaps the most

important regulators of apoptosis due to the fact that they regulate both the intrinsic and

extrinsic pathways (104).

21 

 

1. 2. 8. Assays for apoptosis

Since apoptosis occurs via a complex signalling cascade that is tightly regulated at multiple

points, there are many opportunities to evaluate the activity of the proteins involved. As the

activators, effectors and regulators of this cascade continue to be elucidated, a large

number of apoptosis assays are devised to detect and count apoptotic cells. However, many

features of apoptosis and necrosis can overlap, and it is therefore crucial to employ two or

more distinct assays to confirm that cell death is occurring via apoptosis. One assay may

detect early (initiation) apoptotic events and a different assay may target a later (execution)

event. The second assay, used to confirm apoptosis, is generally based on a different

principle. Multiplexing, which is the ability to gather more than one set of data from the

same sample, is another methodology for apoptosis detection that is becoming increasingly

popular. There are a large variety of assays available, but each assay has advantages and

disadvantages which may make it acceptable to use for one application but inappropriate

for another application (105, 106). Therefore, when choosing methods of apoptosis

detection in cells, tissues or organs, understanding the pros and cons of each assay is

crucial. However, apoptosis assays can be classified into six major groups:

1. Cytomorphological alterations

2. DNA fragmentation

3. Detection of caspases, cleaved substrates, regulators and inhibitors

4. Membrane alterations

5. Detection of apoptosis in whole mounts

6. Mitochondrial assays.

1. 2. 8. 1. Cytomorphological alterations

The evaluation of hematoxylin and eosin-stained tissue sections with light microscopy

does allow the visualization of apoptotic cells. Although a single apoptotic cell can be

detected with this method, confirmation with other methods may be necessary. Because the

morphological events of apoptosis are rapid and the fragments are quickly phagocytosed,

considerable apoptosis may occur in some tissues before it is histologically apparent.

Additionally, this method detects the later events of apoptosis, so cells in the early phase of

apoptosis will not be detected (107).

22 

 

1. 2. 8. 2. DNA fragmentation

The DNA laddering technique is used to visualize the endonuclease cleavage products of

apoptosis. This assay involves extraction of DNA from a lysed cell homogenate followed

by agarose gel electrophoresis. The assay results in a characteristic “DNA ladder” with

each band in the ladder separated in size by approximately 180 base pairs. This method is

easy to perform, has a sensitivity of 1 × 106 cells (i.e., level of detection is as few as

1,000,000 cells), and is useful for tissues and cell cultures with high numbers of apoptotic

cells per tissue mass or volume, respectively. On the other hand, it is not recommended in

cases with low numbers of apoptotic cells. There are other disadvantages to this assay.

Since DNA fragmentation occurs in the later phase of apoptosis, the absence of a DNA

ladder does not eliminate the potential that cells are undergoing early apoptosis.

Additionally, DNA fragmentation can occur during preparation making it difficult to

produce a nucleosome ladder and necrotic cells can also generate DNA fragments (107).

1. 2. 8. 3. Membrane alterations

Externalization of phosphatidylserine residues on the outer plasma membrane of apoptotic

cells allows detection via Annexin V in tissues, embryos or cultured cells. Once the

apoptotic cells are bound with fluorescein isothiocyanate (FITC)-labeled Annexin V, they

can be visualized with fluorescent microscopy. The advantages are sensitivity (can detect a

single apoptotic cell) and the ability to confirm the activity of initiator caspases. The

disadvantage is that the membranes of necrotic cells are labelled as well (108).

1. 2. 8. 4. Detection of apoptosis in whole mounts

Apoptosis can also be visualized in whole mounts of embryos or tissues using dyes such as

acridine orange (AO), Nile blue sulphate (NBS), and neutral red (NR). Since these dyes are

acidophilic, they are concentrated in areas of high lysosomal and phagocytotic activity.

The results would need to be validated with other apoptosis assays because these dyes

cannot distinguish between lysosomes degrading apoptotic debris from degradation of

other debris such as microorganisms. Although all of these dyes are fast and inexpensive,

they have certain disadvantages. AO is toxic and mutagenic and quenches rapidly under

standard conditions whereas NBS and NR do not penetrate thick tissues and can be lost

during preparation for sectioning (109).

23 

 

1. 2. 8. 5. Mitochondrial assays

Mitochondrial assays and cytochrome c release allow the detection of changes in the early

phase of the intrinsic pathway. Laser scanning confocal microscopy (LSCM) creates

submicron thin optical slices through living cells that can be used to monitor several

mitochondrial events in intact single cells over time (110). Mitochondrial permeability

transition (MPT), depolarization of the inner mitochondrial membrane, Ca2+ fluxes,

mitochondrial redox status, and reactive oxygen species can all be monitored with this

methodology. The main disadvantage is that the mitochondrial parameters that this

methodology monitors can also occur during necrosis. The electrochemical gradient across

the mitochondrial outer membrane (MOM) collapses during apoptosis, allowing detection

with a fluorescent cationic dye (69).

1. 3. Erythrocyte suicidal death (Erypyosis)

Erythrocytes are devoid of nuclei and mitochondria and thus lack crucial elements in the

machinery of apoptosis. However, similar to other cell types, erythrocytes have to be

eliminated when they are defective or after their physiological life span of 120 days (111)

in case of human beings. Due to the lack of key organelles involved in the process of

apoptosis it was considered that erythrocytes are unable to undergo apoptosis and hence

have to be eliminated by mechanisms other than apoptosis. It has been observed that

erythrocyte senescence is associated with cell shrinkage, plasma membrane

microvesiculation, a progressive shape change from a discocyte to a spherocyte,

cytoskeleton alterations associated with protein (spectrin) degradation, and loss of plasma

membrane phospholipid asymmetry leading to the externalization of phosphatidylserine in

the erythrocyte membrane (111, 112). The exposure of phosphatidylserine and further eat-

me-signals at the cell surface triggers, and the decrease of cell volume facilitate, the

engulfment of the dying cells by phagocytes (113). Hence, the term “eryptosis” was coined

recently (114) to describe erythrocyte cell death characterised by cell shrinkage, membrane

blebbing, activation of proteases, and phosphatidylserine exposure at the outer membrane

leaflet which are typical features of apoptosis in nucleated cells (115-117).

1. 3. 1. Normal erythrocyte membrane and cation transport

The non-nucleated erythrocyte is unique among human cells in that its plasma membrane

accounts for all of its diverse antigenic, transport, and mechanical characteristics. The

24 

 

discoid shape of the red cell evolves from the multilobulated reticulocyte during 48 hours

of maturation first in the bone marrow and then in the circulation (118).

The structural organization of the human red cell membrane enables it to undergo large

reversible deformations while maintaining its structural integrity during its 4-month life

span in the circulation. The red cell membrane exhibits unique material behaviour. It is

highly elastic (100-fold softer than a latex membrane of comparable thickness), rapidly

responds to applied fluid stresses (time constants in the range of 100 milliseconds), and is

stronger than steel in terms of structural resistance. While a normal red cell can deform

with linear extensions of up to approximately 250%, a 3% to 4% increase in surface area

results in cell lysis. Hence an important feature of induced red cell deformations, both in

vitro and in vivo, is that they involve no significant change in membrane surface area.

These unusual membrane material properties are the result of an evolution-driven

“engineering” process resulting in a composite structure in which a plasma membrane

envelope composed of cholesterol and phospholipids is anchored to a 2-dimensional elastic

network of skeletal proteins through tethering sites on cytoplasmic domains of trans-

membrane proteins embedded in the lipid bilayer (118). Direct interaction of several

skeletal proteins with the anionic phospholipids affords additional attachments of the

skeletal network to the lipid bilayer (113).

The lipid bilayer is composed of equal proportions by weight of cholesterol and

phospholipids (119). While cholesterol is thought to be distributed equally between the two

leaflets, the four major phospholipids are asymmetrically disposed. Phosphatidylcholine

and sphingomyelin are predominantly located in the outer monolayer, while most

phosphatidylethanolamine and all phosphatidylserine (PS), together with the minor

phosphoinositide constituents, are confined to the inner monolayer (120). Several different

types of energy-dependent and energy-independent phospholipid transport proteins have

been implicated in generating and maintaining phospholipid asymmetry. “Flippases” move

phospholipids from the outer to the inner monolayer while “floppases” do the opposite

against a concentration gradient in an energy-dependent manner. In contrast, “scramblases”

move phospholipids bi-directionally down their concentration gradients in an energy-

independent manner (121, 122).

25 

 

The maintenance of asymmetric distribution of phospholipids, in particular exclusive

localization of PS and phosphoinositides to the inner monolayer, has several functional

implications. Because macrophages recognize and phagocytose red cells that expose PS at

their outer surface, the confinement of this lipid in the inner monolayer is essential if the

cell is to survive its frequent encounter with macrophages of the reticuloendothelial

system, especially the spleen. Loss of lipid asymmetry leading to exposure of PS on the

outer monolayer has been suggested to play a role in premature destruction of thalassemic

and sickle red cells. Furthermore, the restriction of PS to the inner monolayer also inhibits

the adhesion of normal red cells to vascular endothelial cells, thereby ensuring unimpeded

transit through the microvasculature (123, 124).

More than fifty trans-membrane proteins with variable abundance, ranging from a few

hundred to a million copies per red cell, have been characterized. A large fraction of these

trans-membrane proteins define the various blood group antigens. The membrane proteins

exhibit diverse functional heterogeneity, serving as transport proteins. Most proteins

associated with the erythrocyte membrane are involved in active and passive ion transport,

i.e. processes to maintain erythrocyte homeostasis. Other erythrocyte membrane proteins

may act as adhesion proteins involved in interactions of red cells with other blood cells and

endothelial cells, and also as signalling receptors (123, 125).

Membrane proteins with transport function include band 3 (anion transporter), aquaporin 1

(water transporter), GLUT-1 (glucose and L-dehydroascorbic acid transporter), Kidd

antigen protein (urea transporter), RhAG (gas transporter, probably of carbon dioxide),

Na+-K+-ATPase, Ca++ ATPase, Na+-K+-2Cl− cotransporter, Na+-Cl− cotransporter, Na+-K+

cotransporter, K+-Cl− cotransporter, and Gardos Channel (123, 126).

1. 3. 1. 1. Na+ / K+ pump in non-infected erythrocytes

The Na+/K+ pump mechanism of human erythrocytes is well understood. It builds up and

maintains a high intracellular K+ ([K+]i) and a low intracellular Na+ ([Na+]i) concentration.

The ouabain-sensitive Na+/K+-ATPase in the erythrocyte membrane pumps 2 K+ ions into

versus 3 Na+ out of the cell, thereby generating opposing chemical gradients for the two

ions. The pumping helps to build up the membrane potential and counterbalances the

"leak" of the Na+ and K+ ions down their respective concentration gradients via various

cotransporters, exchangers, and channels that adjust a steady-state cytoplasmic [Na+]-to-

26 

 

[K+] ratio of 0.12-0.16 in normal human erythrocytes (127). Intraerythrocytic Na+ and K+

concentrations have been determined as approximately 10 to 20 mM and 140 mM,

respectively. Several cation channels that may contribute to the cation leak have been

identified electrophysiologically (128).

1. 3. 1. 2. Nonselective cation channels in non-infected erythrocytes

Whole-state and nystatin-perforated patch-clamp single-channel recordings have led to the

discovery of nonselective cation (NSC) channels in the human RBC membrane that are

activated by strong depolarization of the membrane potential (129, 130). These voltage-

gated NSC channels are stimulated through nicotinic acetylcholine and PGE2 receptors.

They are permeable to divalent cations such as Ca2+, Ba2+, and Mg2+ (131, 132), and

exhibit a hysteresis-like voltage-dependent gating (133). In addition, voltage-independent

NSC channels have been discovered. Experiments with simultaneous activity of both

channel types suggest their differential regulation (114).

Activation of the voltage-independent NSC channels occurs within minutes upon

replacement of extracellular Cl- by gluconate (115, 116), similar when extracellular Cl- is

replaced by NO3-, Br-, or SCN-. Furthermore, voltage-independent NSC channels are

activated when the ionic strength of the bath solution is lowered by isosmotic substitution

of NaCl with sorbitol, indicating that the underlying process is probably identical with that

of the Na+ and K+ permeability increase upon incubation of human erythrocytes in low

ionic strength medium (129, 132). Decrease of the extracellular Cl- concentration activates

these cation channels. Remarkably, the anion channel/transporter inhibitor

diisothiocyanatostilbene-2′,2-disulfonic acid (DIDS) directly inhibits K+ efflux in low ionic

strength medium paralleled by inhibition of Cl- exchange. Moreover, whole-cell recordings

show that upon extracellular Cl- removal DIDS prevents activation of the NSC channel,

while having no effect on the activated cation channels (134). Furthermore, cation channel

activity strongly depends on the cytosolic (i.e., pipette) Cl- concentration (135), suggesting

that a DIDS-sensitive pathway equilibrates the Cl- concentrations between the cytoplasmic

and extracellular membrane face and that intracellular rather than extracellular Cl- ions

modulate the cation channel activity (135). The voltage-independent channels hardly

discriminate between monovalent cation channels (cation permselectivity in the rank order

of Cs+ > K+ > Na+ = Li+ >> NMDG+) and have a Ca2+-permeability similar to that of

voltage-gated NSC channels (128).

27 

 

1. 3. 1. 3. Ca2+ activated K+ channels (Gardos channels)

The erythrocyte uses at least four well-characterized K+ transport mechanisms: the K+Cl-

cotransporter, a NaK2Cl cotransporter, a Na+/K+ ATPase (a sodium pump) and a Ca2+

activated K+ channel (Gardos channels) (136). The gene for the Gardos channel in

erythrocytes is the 4th member of the potassium intermediate/small conductance calcium-

activated channel subfamily N (KCNN4; aliases: hsK4 (human small conductance

potassium channel 4), hIK1 (human intermediate conductance potassium channel) (115,

137). Gardos channels have been characterized by single channel recordings (138). They

play an important role in the regulation of cell volume and are major factors in HbS/S

RBC dehydration (125).

Gardos channel activity is dependent on the free Ca2+ concentration at the cytoplasmic

membrane face. , Concomitant with an increase in free [Ca2+]i from 500 nM to 60 µM its

open probability increases from 0.1 to 0.9. Ca2+ acts via binding to calmodulin

constitutively associated with the Gardos channels (139). In addition to Ca2+, channel

activity is dependent on intracellular K+, and inhibited by extracellular Na+, K+. In

addition, protein kinase A (PKA) reportedly induces a dramatic enhancement of Gardos

channel activity, possibly by modulating the Ca2+ sensitivity (138).

In non-stressed and non-stimulated human erythrocytes the fractional Gardos K+ channel

activity is low, resulting in a very low membrane potential (≤ -10 mV) (140). As a

consequence, human erythrocytes relatively have high intracellular Cl- concentrations.

Activation of Gardos channels by increased intraerythrocytic Ca2+ levels (≥ 150 nM) leads

to hyperpolarization of the RBC membrane potential close to the K+ equilibrium potential

(115, 141). This imposes an outwardly directed electrochemical Cl- gradient across the

RBC membrane, driving Cl- and K+ into the extracellular space, followed by osmotically

obliged water efflux. The Gardos channel inhibitors clotrimazole and charybdotoxin (138,

142), or Cl- channel blocker 5-Nitro-2- (3-phenylpropylamino) benzoic acid (NPPB) blunt

phosphatidylserine (PS) exposure, hyperosmotic and Ca2+-stimulated isosmotic cell

shrinkage, confirming the contribution of Gardos and anion channels to KCl and water

efflux during RBC shrinkage (143, 144).

Increase in free [Ca2+]i therefore favors RBC shrinkage (Gardos effect) (144) subsequently

decreased RBC deformability under isosmostic conditions, measured as decreased cell size

28 

 

or filterability (141). The Gardos effect is dependent on the extracellular Ca2+

concentration and is sensitive to amiloride (115). During complement activation rising

cytosolic calcium triggers the Gardos effect, thus limiting the colloidosmotic swelling and

lysis of erythrocytes (145).

1. 3. 2. Mechanisms of erythrocyte cell death or eryptosis

Eryptosis is triggered by erythrocyte injury after several stressors, including oxidative

stress. Besides caspase activation after oxidative stress, two signalling pathways converge

to trigger eryptosis: (a) formation of prostaglandin E2 leads to activation of Ca2+

permeable cation channels, and (b) the phospholipase-A2 mediated release of platelet-

activating factor activates a sphingomyelinase, leading to formation of ceramide. Increased

cytosolic Ca2+ activity and enhanced ceramide levels lead to membrane scrambling with

subsequent phosphatidylserine exposure. Moreover, Ca2+ activates Ca2+-sensitive K+

channels, leading to cellular KCl loss and cell shrinkage. In addition, Ca2+ stimulates the

protease calpain, resulting in degradation of the cytoskeleton (108, 146).

1. 3. 2. 1. Role of prostaglandins in stimulation of erythrocyte cation channels

Intriguing evidence points to a role of prostaglandins in the regulation of eryptosis.

Hyperosmotic shock and Cl--removal trigger the release of prostaglandin E2 (PGE2).

PGE2 in turn activates the cation channels, increases the cytosolic Ca2+ concentration, and

stimulates phosphatidylserine exposure at the erythrocyte surface. The activation of the

cation channels by Cl- -removal is abolished by the cyclooxygenase inhibitor diclofenac.

Moreover, phospholipase-A2 inhibitors quinacrine and palmitoyl trifluoromethyl ketone

and cyclooxygenase inhibitors acetylsalicylic acid and diclofenac blunt the increase of

phosphatidylserine exposure following Cl- removal. PGE2 further activates the Ca2+

dependent cysteine endopeptidase calpain, an effect, however, apparently not required for

stimulation of phosphatidylserine exposure but playing a role in the degradation of the

cytoskeleton (141).

1. 3. 2. 2. Role of Ca2+ sensitive K+ channels (Gardos channels) in eryptosis

Calcium ions entering erythrocytes do not only activate the scramblase but in addition

stimulates the Ca2+ sensitive “Gardos” K+ channels in erythrocytes (147). The activation of

the channels leads to hyperpolarization of the cell membrane driving Cl- in parallel to K+

into the extracellular space. The cellular loss of KCl favours cell shrinkage. Moreover, the

29 

 

cellular loss of K+ presumably participates in the triggering of “eryptosis”. Increase of

extracellular K+ or pharmacological inhibition of the Gardos channels by clotrimazole or

charybdotoxin do not only blunt the cell shrinkage but also decrease the phosphatidylserine

exposure following exposure to ionomycin. Presumably, cellular loss of K+ somehow

stimulates “eryptosis” as has been shown for apoptosis of nucleated cells (148). As PGE2

increases cytosolic Ca2+ activity, it similarly activates the Ca2+ sensitive “Gardos” K+

channels with subsequent cell shrinkage (149).

1. 3. 2. 3. Role of protein kinase C in eryptosis

Erythrocyte energy depletion enhances phosphorylation of membrane proteins by protein

kinase C (PKC), leads to subsequent phosphatidylserine exposure at the cell surface and

triggers cell shrinkage, effects mimicked by stimulation of protein kinase C (PKC) with

phorbol esters or inhibition of protein phosphatises with okadaic acid (150). PKC

activation has previously been shown to stimulate erythrocyte Ca2+ entry and

phosphatidylserine exposure (151). PKC is a family of serine/threonine-specific protein

kinases consisting of 10 members and requiring Ca2+, diacylglycerol, and a phospholipid

for activation. PKC isoenzymes play an essential role in the regulation of diverse cellular

functions including proliferation, differentiation, and apoptosis. Human erythrocytes

express PKC isoenzymes mediating the phosphorylation of cytoskeletal proteins, such as

band 4.1, 4.9, and adducing the human Na+/H+ exchanger (NHE 1) (151, 152).

1. 3. 2. 4. Role of oxidative stress and caspase activation in eryptosis

Oxidative stress resulting from exposure to tert-butylhydroperoxide or peroxynitrite, for

instance, is a major cause of erythrocyte injury (153, 154). It has been shown to activate

aspartyl and cysteinyl proteases (155). Caspases have been shown to be expressed in

erythrocytes, to cleave the anion exchanger band 3 in vitro (155), and to stimulate

phosphatidylserine exposure of erythrocytes (156). Conversely, eryptosis after ionomycin

or hyperosmotic shock does not require activation of caspases (157). Besides its effect on

caspases, oxidative stress or defects of antioxidative defence, enhance Ca2+ entry via

activation of the cation channels and thus stimulate eryptosis at least partially through

channel activation (158). Oxidation of erythrocytes leads to an increase of the cation

permeability of the membrane, Oxidation with tert-butylhydroperoxide also enhanced

erythrocyte annexin-V binding as a measure of phosphatidylserine exposure by some six

folds. Interestingly, not all erythrocytes of one population showed the same sensitivity

30 

 

against oxidative stress, and only one third of the population was shown to be annexin-V

positive (141). Oxidative stress further activates erythrocyte Cl- channels (159), which are

required for erythrocyte shrinkage and thus also participate in the triggering of eryptosis

(143). Antioxidants, such as vitamin E (160, 161), glutathione (162), or the semi-synthetic

flavonoid 7-monohydroxyethylrutoside may protect erythrocytes from oxidative stress and

thus presumably from eryptosis (163).

1. 3. 2. 5. Role of platelet-activating factor (PAF) and stimulation of sphingomyelinase

in eryptosis

Erythrocyte shrinkage triggers the formation of platelet-activating factor (PAF) which is

involved in the regulation of inflammation, thrombosis, atherogenesis, and cardiovascular

function (164). PAF then stimulates a sphingomyelinase, leading to the breakdown of

sphingomyelin and release of ceramide from erythrocytes (122). Osmotic shock thus leads

to the appearance of ceramide at the erythrocyte surface. At least partially because of

ceramide formation, PAF triggers scrambling of the cell membrane with

phosphatidylserine exposure at the erythrocyte surface. C6- ceramide as well as treatment

with purified, bacterial sphingomyelinase similarly triggers phosphatidylserine scrambling

(165). Moreover, eryptosis after osmotic shock is blunted by the sphingomyelinase

inhibitor 3,4-dichloroisocoumarin. PAF further activates Ca2+-sensitive K+ channels

(Gardos channels) in the erythrocyte cell membrane (166). Conversely, PAF is released

from erythrocyte progenitor cells on increase of cytosolic Ca2+ activity (114). The

signalling through PAF does, however, not necessarily require elevated cytosolic Ca2+

concentrations, and enhanced PAF levels at least partially account for Ca2+-independent

eryptosis (164).

1. 3. 3. Eryptosis inhibitors

Erythropoietin is a potent inhibitor of erythrocytic progenitor cells apoptosis, and also

inhibits the suicidal death of mature erythrocytes. The hormone is effective through

inhibition of the Ca2+-permeable cation channels. The anti-eryptotic effect of

erythropoietin presumably accounts for its ability to increase the life span of circulating

cells. Dopamine, isoproterenol, and epinephrine similarly inhibit the Ca2+-permeable cation

channels and thus interfere with eryptosis. The effect is presumably not relevant for

physiologic regulation, as the concentrations of catecholamines required exceed those

encountered in vivo. However, animal experiments revealed that the hematotoxicity

31 

 

(including red blood cell counts) of cyclophosphamide has been reduced by simultaneous

dopamine treatment. Thus, administration of dopamine at pharmacologic doses may well

be therapeutically applicable for the suppression of eryptosis (167).

1. 3. 4. Clinical conditions associated with eryptosis

A variety of clinical conditions decrease the life span of mature erythrocytes by facilitating

eryptosis. Increased sensitivity of sickle cells and of glucose 6-phosphate dehydrogenase–

deficient cells to osmotic shock, oxidative stress and glucose depletion has been observed

previously. The enhanced susceptibility to eryptosis is believed to decrease the erythrocyte

life span in those disorders (168).

Similarly, iron-deficient erythrocytes are more sensitive to eryptosis. The enhanced

eryptosis is at least partially the result of enhanced cation channel activity. Presumably, the

decreased volume of iron-deficient erythrocytes decreases the threshold for the activation

of the channel. Interestingly, the enhanced exposure of phosphatidylserine on the surface

of iron-deficient erythrocytes coincides with a substantial decrease of the life span of iron-

deficient erythrocytes (169). Eryptosis is further triggered by exposure of erythrocytes to

lead or mercury, or by treatment with plasma from patients with recurrent hemolytic-

uremic syndrome. Thus, the typical anemia after lead intoxication is at least partially due to

enhanced eryptosis. Oxidant injury of erythrocytes and/or erythrocyte precursors may

further occur in HIV (162).

Even though it is a pathophysiologic mechanism, eryptosis serves an important physiologic

function (i.e., the prevention of hemolysis). Energy depletion, defective Na+/K+ATPase, or

enhanced leakiness of the cell membrane eventually leads to cellular gain of Na+ and Cl–

with osmotically obliged water, resulting in subsequent cell swelling (170). Initially, the

entry of Na+ is compensated by cellular loss of K+; the decrease of the K+ equilibrium

potential leads, however, to gradual depolarization, which favours the entry of Cl-. The cell

swelling jeopardizes the integrity of the cell membrane. Rupture of the cell leads to release

of hemoglobin, which may be filtered at the glomeruli of the kidney, precipitate in the acid

lumen of the tubules, obliterate the tubules, and thus lead to renal failure. The

phosphatidylserine exposure at the cell surface allows the macrophages to recognize

defective erythrocytes and to clear them from circulating blood before hemolysis.

Moreover, the activation of the Gardos K+ channel delays swelling and disruption of

32 

 

defective erythrocytes, thus expanding the time allowed for macrophages to clear the

injured erythrocytes from circulating blood (167).

1. 3. 5. Eryptosis in malaria

1. 3. 5. 1. Stimulation of membrane transport in P. falciparum infected erythrocytes

and activation of ion channels

A prerequisite for intracellular survival of pathogens is an adequate supply of nutrients and

disposal of waste products (171). Additional transport systems for nutrient supply and

disposal of waste products are needed particularly during infection of erythrocytes with

Plasmodium species. Before infection, the substrate turnover of erythrocytes is low. Since

they do not synthesize proteins, DNA or membrane components, they have no need for

amino acids, nucleic acids, lipids or most of the vitamins. The replicating Plasmodium,

however, has extensive requirements for all of these nutrients and, to fuel the replication

process, Plasmodium parasites consume large amounts of glucose. An infected erythrocyte

takes up 40–100 times more glucose than a non-infected cell and releases the

corresponding amounts of lactic acid (172). To gain access to the necessary nutrients and

to dispose the waste products, Plasmodium induces the so-called new permeability

pathway (NPP). This pathway mediates the transport of electrolytes, sugars, nucleic acids,

membrane components and other substances across the erythrocyte membrane. In addition

to nutrient uptake and lactate disposal the NPP contributes to the strategy of the pathogen

to counteract swelling of the host cell by decreasing the colloid-osmotic pressure in the

host cytosol. The parasite digests haemoglobin in excess (i.e. more than needed for its

protein biosynthesis) and exports the haemoglobin-derived amino acids via the NPP thus

preventing host cell swelling and lysis (173). NPP further allows the entry of Na+ and Ca2+

into the host cell, thus adjusting the electrolyte composition of the host cytosol to the needs

of the parasite. Starting >15 h post-infection, K+ in the host cytosol is replaced increasingly

by Na+. The parasite most probably requires an outwardly directed K+ and an inwardly

directed Na+ gradient across its plasma membrane since decreasing these gradients

experimentally in vitro abolishes parasite growth. In addition, absence of extracellular Ca2+

disrupts the intraerythrocyte survival of the pathogen (174). The necessity of the NPP for

Plasmodium’s survival is illustrated by the fact that several inhibitors of this pathway

eventually kill the pathogen within the erythrocyte (175). On the other hand, the

33 

 

Plasmodium species imposes oxidative stress on host cells, and this has been proposed to

be required for the activation of the cation channels (135).

1. 3. 5. 2. Induction of eryptosis by P. falciparum infection

The opening of cation channels in the host cell by P. falciparum is expected to increase the

cytosolic Ca2+ activity and thus to induce eryptosis. However, most of the Ca2+ entering the

parasitized erythrocyte is either extruded across the host cell membrane or taken up by the

pathogen so that cytosolic free Ca2+ activity remains low (176). Nevertheless, some in vitro

studies have shown infection with P. falciparum increase the number of erythrocytes that

expose phosphatidylserine at the outer membrane leaflet, (177) while another study could

not detect breakdown of the phospholipid asymmetry. Infection with P. falciparum

increases the formation of ceramide (122) that in turn increases the Ca2+ sensitivity of the

erythrocyte scramblase and may account for the stimulation of the scramblase even at low

or only slightly enhanced cytosolic Ca2+ concentrations. Beyond that, Ca2+ independent

mechanisms may participate in the triggering of eryptosis. As observed in nucleated cells

the cellular loss of K+ via the induced cation channel is expected to augment the activation

of the erythrocyte scramblase and thus adds to the stimulation of the phosphatidylserine

exposure (178).

1. 3. 5. 3. Role of eryptosis in parasitized erythrocytes

Human erythrocytes infected with trophozoite and shizont stages of P. falciparum may

adhere to endothelial cells lining the post-capillary venules. Adhesion of the infected

erythrocyte to the endothelium involves receptors in the endothelial membrane such as

CD36, secreted proteins such as thrombospondin, parasite-encoded molecules such as P.

falciparum erythrocyte membrane 1 (PfEMP1) and a modified erythrocyte anion

exchanger (AE1, band 3) (177). In particular, exposure of phosphatidylserine at the outer

membrane leaflet contributes to erythrocyte-endothelium adhesion via binding to

thrombospondin and CD36 receptor. Thus “eryptosis” of parasitized erythrocytes may

contribute to tissue sequestration, which is thought to avoid splenic clearance of the

parasitized erythrocyte and to favour parasite development in a low oxygen pressure

microenvironment. On the other hand “eryptosis” may result in clearance of parasitized

erythrocytes. As macrophages are equipped with receptors specific for phosphatidylserine,

erythrocytes exposing phosphatidylserine at their surface will be recognized, bound,

engulfed and degraded rapidly. Thus, infected erythrocytes are liable to be eliminated as

34 

 

soon as they expose phosphatidylserine. The removal of the infected erythrocyte would of

course clear the pathogen as well, since it would be similarly degraded by lysosomal

enzymes of the macrophages (179, 180).

1. 4. Rodent malaria parasites as models for human malaria

A wide range of investigations using rodent parasites have contributed to development and

shaping of basic concepts in research of human diseases. Rodent malaria parasites are

practical models for the experimental study of mammalian malaria. These parasites have

proved to be analogous to human malaria in most essential aspects of structure, physiology

and life cycle (181). Metabolic pathways are conserved between rodent and human malaria

parasites. No gross differences in metabolic pathways have been reported in the various

malaria parasites. The similarity in sensitivity of these parasites to anti-malarial drugs and

other specific inhibitors emphasises the similarities in their metabolic processes. The life

cycles and the different developmental stages of all rodent and human malaria parasites are

largely comparable (182, 183). Rodent parasites are recognized as valuable model parasites

for the investigation of the developmental biology of malaria parasites, parasite-host

interactions, vaccine development and drug testing. The relationships of rodent parasites to

other human malaria parasites in brief are:

• The basic biology of rodent and human parasites is similar.

• The genome organization and genetics is conserved between rodent and human parasites

(184).

• Housekeeping genes and biochemical processes are conserved between rodent and human

parasites.

• Methodologies for genetic modification are available for both types of parasites.

• Rodent hosts with extensively characterised genetic backgrounds and transgenic lines are

valuable and available tools for immunological studies.

• The structure and function of vaccine candidate target antigens are conserved between

rodent and human parasites.

• The manipulation of the complete lifecycle of rodent parasites, including mosquito

infections is simple and safe.

• In vitro culture techniques for large-scale production and manipulation of different life

cycle stages are available. For example, in vitro cultures of liver and mosquito stages

provide tools to investigate the less accessible parts of the life cycle in the human

35 

 

• The molecular basis of drug-sensitivity and resistance show similar characteristics in

rodent and human parasites.

• Rodent parasites allow in vivo investigations of parasite-host interactions and in vivo drug

testing.

The malaria parasites P. berghei which infects hamsters, rats and mice, is one of the many

species of malaria parasites that infect mammals other than humans. A susceptible

mosquito vector for P.berghei, which is widely used in the laboratory, is Anopheles

stephensi (182, 184).

1. 5. Pharmacological agents

1. 5. 1. Gum Arabic

Gum Arabic (GA) is a water-soluble dietary fibre derived from the dried gummy exudates

of the stems and branches of Acacia Senegal (185). Chemically, GA is a polysaccharide of

(1-3) linked β-D-galactopyranosyl units with branches of two to five units in length

attached by (1-6) links to the main chain. Both the main chain and side chains contain α-L-

arabinofuranosyl, α-L-rhamnopyranosyl, β-D-glucuronopyranosyl, and 4-O-methyl-β-D-

glucuronopyranosyl units. Gum arabic is readily soluble in water without increasing

viscosity (186).

Gum Arabic is a highly heterogeneous material, but was separated into three major

fractions by hydrophobic affinity chromatography (187). Most of the gum (88.4% of total),

an arabinogalactan (AG), had a very low protein content (0.35%) and a molecular mass of

3.8 - 105 Da. The second fraction (10.4% of total), an arabinogalactan-protein complex

(AGP), contained 11.8% protein and had a molecular mass of 1.45 - 106 Da. The third

fraction (1.2% of total gum), referred to as a low molecular weight glycoprotein (GP), had

a protein content of 47.3%, and a molecular mass of 2.5 - 105 Da (GPC data). The major

amino acids present in the protein of AG and AGP were hydroxyproline, serine and

proline, whereas in GP, aspartic acid is the most abundant (188).

Gum Arabic is primarily indigestible to both humans and animals. It is not degraded in the

small intestine, but fermented in the large intestine by microorganisms to short-chain fatty

acids, particularly butyric and propionic acids. Such degradation products are absorbed in

the human colon and subsequently metabolized. In an experiment using an enrichment

36 

 

culture of pig cecal bacteria, showed that a Prevotella ruminicola-like bacterium was the

predominant organism that is most likely to be responsible for the fermentation of GA to

butyrate and propionate (189, 190).

Gum Arabic is widely used in both the pharmaceutical and food industries as an emulsifier

and stabilizer of various products for human consumption. Examples include tablets or

food applications such as puddings and fillings, frostings, candy, and chewing gum. The

United States Food and Drug Administration recognize it as one of the safest dietary fibers

(189, 191).

Pharmacologically, GA has been claimed to act as an anti-oxidant, and to protect against

experimental hepatic, renal and cardiac toxicities in rats. GA has been claimed to alleviate

the adverse effects of chronic renal failure in humans. This could not be corroborated

experimentally in rats (183). Reports on the effects of GA on lipid metabolism in humans

and rats are contradicting, but mostly suggest that GA ingestion can reduce plasma

cholesterol concentrations. GA has proabsorptive properties and can be used in diarrhoea.

It enhances dental remineralization, and has some antimicrobial activity, suggesting a

possible use in dentistry. GA has been shown to have an adverse effect on electrolyte

balance and vitamin D in mice, and to cause hypersensitivity in humans (183).

1. 5. 2. Amphotericin B

Amphotericin B is a polyene antifungal agent with an in vitro activity against a wide

variety of fungal pathogens. Amphotericin B exerts its antifungal effect by disruption of

fungal cell wall synthesis because of its ability to bind to sterols, primarily ergosterol. This

affinity may also account for its toxic effects against selected mammalian cells.

Amphotericin B may be fungistatic or fungicidal, depending upon drug concentration and

sensitivity of the pathogen (192).

Oral preparations of amphotericin B are used to treat oral thrush; these are virtually

nontoxic. The main i.v. use is in systemic fungal infections (eg. in immunocompromised

patients), and in visceral Leishmaniasis. Aspergillosis, Cryptococcus infections (eg.

meningitis) and Candidiasis are treated with amphotericin B. It is also used empirically in

febrile immunocompromised patients who do not respond to broad-spectrum antibiotics

(193).

37 

 

1. 5. 2. 1. Mode of action

As with other polyene antifungals, amphotericin B associates with ergosterol, a membrane

chemical of fungi, forming a pore that leads to K+ leakage and fungal cell death. Recently,

however, researchers found evidence that pore formation is not necessarily linked to cell

death. The actual mechanism of action may be more complex and multi-faceted.

Amphotericin B is believed to interact with membrane sterols (ergosterol) to produce an

aggregate that forms a trans-membrane channel. Intermolecular hydrogen bonding

interactions among hydroxyl, carboxyl and amino groups stabilize the channel in its open

form, destroying activity and allowing the cytoplasmic contents to leak out (194).

1. 5. 2. 2. Side effects

Very often a most serious acute reaction after the infusion (1 to 3 hours later) is noted

consisting of fever, shaking chills, hypotension, anorexia, nausea, vomiting, headache,

dyspnea, and tachypnea. This reaction sometimes subsides with later applications of the

drug and may in part be due to histamine liberation. An increase in prostaglandin-synthesis

may also play a role. Often the most difficult decision has to be made, whether the fever is

disease or drug-related. In order to decrease the likelihood and severity of the symptoms,

initial doses should be low and increased slowly. The liposomal preparation obviously has

a lower incidence of the syndrome. Acetaminophen, pethidine, diphenhydramine and/or

hydrocortisone have all been used to treat or prevent the syndrome, but the prophylactic

use of these drugs should be limited (195).

Nephrotoxicity (kidney damage) is a major issue and can be severe and/or irreversible. It is

much milder when amphotericin B is delivered in liposomes. Electrolyte imbalances (eg.

hypokalemia and hypocalcemia) may also occur (196).

1. 6. Rational of the study

Targeting the pathogen with antimalarial drugs has only been partially successful because

of the spread of drug-resistant parasites and the optimal use of effective drugs has always

been a major concern. Hence all possibilities to combat this disease must be explored. The

course of the disease is not only a function of the pathogen but is heavily influenced by

properties of the host. Hence, studies must be carried out to fight infection by altering host

physiology which decimates the problem of drug resistance and if successful, these

38 

 

approaches may prove extremely useful, particularly in the treatment of resistant

pathogens. Targeting at the accelerated clearance of the infected host cells is expected to

provide protection against malaria.

Pharmacological induction of phosphatidylserine exposure of ring stage-infected

erythrocytes reportedly accelerates their clearance. Thus, manoeuvres accelerating

eryptosis may result in premature clearance of the intraerythrocytic parasite and partially

enhance the survival of Plasmodium berghei-infected mice. Importantly, counteracting

Plasmodia by inducing eryptosis is not expected to generate resistance of the pathogen, as

the proteins involved in suicidal death of the host cell are not encoded by the pathogen and

thus cannot be modified by mutations of its genes.

In this study we are interested in investigating the role of phosphatidylserine (PS) exposure

and eryptosis which may allow the pharmacological manipulation of erythrocyte survival

and hence of the malaria disease course. This may be achieved by determining the effect of

eryptosis inducing drugs on the proliferation and survival of the Plasmodium pathogen.

39 

 

1. 7. The objectives of the study

1. 7. 1. General objective

The aim of the present study is to investigate the effect of amphotericin B or gum Arabic

administration on eryptosis of infected cells and their influence on the course of malaria in

P. berghei infected mice.

1. 7. 2. Specific objectives

1. To determine the effect of amphotericin B or butyrate on in vitro growth and DNA

amplification of P. falciparm infected erythrocytes.

2. To determine the effect of amphotericin B or butyrate on the cytosolic Ca2+ activity

of P. falciparm infected and non-infected erythrocytes.

3. To determine the effect of amphotericin B or butyrate on the phosphatidylserine

exposure on P. falciparm infected and non-infected erythrocytes.

4. To determine the effect of amphotericin B or gum Arabic administration on

eryptosis of infected cells and their influence on the course of malaria in P. berghei

infected mice.

5. To determine the significance of phosphatidylserine exposure in clearance and

elimination of infected erythrocytes.

40 

 

2. Materials and methods

2. 1. Animals

Experiments were carried out on 4 to 5-month-old wild-type SV129/mJ mice of either sex

(n = 6 male, 8 female for each one of the three series) in gum Arabic and amphotericin B

experiments. The animals were housed under controlled environmental conditions (22-

24°C, 50-70% humidity, and a 12-h light/dark cycle). Throughout the study, mice had free

access to standard pelleted food (C1310, Altromin, Heidenau, Germany) and tap water or

GA solution, as indicated. All animal experiments were conducted according to the

guidelines of the American Physiological Society and the German law for the care and

welfare of animals, and were approved by local authorities in Eberhard Karls University of

Tübingen, Germany. (Registration number PY 2/06). Mouse erythrocytes were drawn from

animals by retroorbital venopuncture or by incision of the tail vein.

2. 2. Parasites

Plasmodium falciparum BINH (197) and Plasmodium berghei ANKA (198) were a kind

gift from the Institute of Tropical Medicine in Tuebingen, Germany. All chemicals and

equipments used in this study are shown in appendix 1.

2. 3. Infection of mice

For infection of mice Plasmodium berghei ANKA-parasitized murine erythrocytes (1x106)

were injected intraperitoneally into wild-type mice (199, 200).

2. 4. Gum Arabic treatment

Gum Arabic in powder form was provided as a generous gift from Dar Savanna, Ltd.,

Khartoum, Sudan (www.ssgums.com). It is a 100% natural extract powder produced

mechanically from the wildly grown Acacia senegal tree, with a particle size of <210 µm.

Its quality conforms to the food and pharmaceutical requirements of the Food and

Agriculture Organization of the United Nations (FAO), the British Pharmacopoiea (BP),

the United States Pharmacopoiea (USP), and the Joint FAO/WHO Expert Committee on

Food Additives (JECFA).

Experiments were carried out on wild-type SV129/mJ mice of either sex (n = 6 male, 8

female for each one of the three series). Starting from the day of infection, animals were

provided with 10% (w/w) GA dissolved in tap water (100 g/L). Preparations were

41 

 

refreshed every day during the treatment. The effects of GA treatment were investigated

starting from day 8 after the infection. The intake corresponded to a dose of approximately

10 g/kg body weight/day. Blood was collected from the mice 8 days after infection by

incision of the tail. Parasitemia was determined by Syto-16 staining in FACS analysis or

using Giemsa stain.

2. 5. Amphotericin B administration

Experiments were carried out on six male mice for control and other six mice for

Plasmodium berghei infected group in each one of the three series of the experiment.

Amphotericin B (1.4 mg/kg body weight for three successive days) was administered

subcutaneously from the eighth day of infection. Blood was collected from the mice 8 days

after infection by the incision of the tail. Parasitemia was determined by Syto-16 staining

in FACS analysis and Giemsa stain.

2. 6. Preparation of human erythrocytes

Human erythrocytes were drawn from healthy volunteers. They were used after separation

by centrifugation for 25 min; 2000 g over Ficoll (Biochrom KG, Berlin, Germany). The

buffy coat and upper 10-20% of the red blood cells were discarded; the remaining pellet

was used for experiments after washing three times in phosphate buffer saline. The RBCs

were stored at 4°C until use (2-5 days). Experiments were performed at 37°C in Ringer

solution containing (in mM) 125 NaCl, 5 KCl, 1 MgSO4, 32 N-2-hydroxyethylpiperazine-

N-2-ethanesulfonic acid (HEPES)/NaOH (pH 7.4), 5 glucose, 1 CaCl2

or RPMI 1640

medium. Blood donors gave their informed consent and the study was approved by the

Ethical Committee of Eberhard Karls University of Tübingen, Germany (project number:

184/2006V). (Appendix-2 protocol 1)

2.7. Investigations

2.7.1. In vitro culture of Plasmodium falciparum infected human erythrocytes

For infection of human erythrocytes, the P. falciparum strain BinH (197) was grown in

vitro (135) in human erythrocytes. Cultures were maintained continuously by routine

passage in fresh and stored human erythrocytes. Parasites were cultured as described

earlier (201, 202) at a hematocrit of 2-5% and a parasitemia of 2-10% in RPMI 1640

medium supplemented with 0.5% Albumax II (Gibco, Karlsruhe, Germany), 0.13 µM

42 

 

hypoxanthine, 2 mM L-glutamine (Gibco, Karlsruhe, Germany), 25 mM HEPES/NaOH

pH 7.4 (Sigma-Aldrich, Schnelldorf, Germany), 20 µg/ml gentamycin (Gibco, Karlsruhe,

Germany) in an atmosphere of 90 % N2, 5 % CO

2, 5 % O

2.

The dilution factors (Df) for splitting the infected RBCs suspension in order to maintain

the P. falciparum in vitro culture were calculated as follows:

Df = mP * V(RBC) / dP * dV(RBC)

Fresh blood was added according to the following formula:

dV(RBC) – (V(RBC) )/ Df

Where mP: measured parasitemia; V(RBC): volume of packed erythrocytes in the culture

flask; dP: desired parasitemia; dV(RBC): desired volume of packed erythrocytes in the

flask. The volume of the medium was adjusted according to the degree of parasitemia and

the time until the next medium change (203) (Appendix-2 protocol 2).

2.7. 1. 1. Freezing and sawing of parasites

From time to time samples with high, i.e. ≥ 20 % parasitemia, predominantly containing

ring stages were deep frozen in liquid nitrogen for stock purposes. The cell pellet obtained

after centrifugation (450g / 8 min at RT) was mixed with an equal volume of freezing

solution, sterile transferred to a 2.0 ml cryotube vial and directly deep frozen in liquid

nitrogen. For defreezing the parasites from cryotubes containing infected RBCs (> 20 %

parasitemia) were thawed quickly at 37°C in a water bath (3-5 min). The contents were

transferred to a 15 ml falcon tube and centrifuged (450g / 4 min at room temperature). The

supernatant was discarded. A volume of sterile filtered 3.5 % NaCl solution equal to cell

pellet volume (V(RBC)) was added at a rate of 1 - 2 drops per second while gently shaking

the tube. Finally the liquid was mixed with a pipette. After centrifugation at 450g for 4 min

the supernatant was discarded. The RBC pellet was carefully suspended in original RPMI

1640 medium (2*V (RBC)). Washing was repeated twice. The remaining cell pellet was

re-suspended in complete medium and transferred to a culture flask. Fresh RBCs were

added (amount depending on V(RBC)). The culture flask was filled with 90% N2

/5% O2

/

5% CO2 (Appendix-2 protocol 3).

43 

 

2. 7. 1. 2. Isosmotic sorbitol synchronization of P. falciparum infected human

erythrocytes

The P. falciparum BinH strain was cultured and synchronized to the ring stage by sorbitol

treatment (197). Briefly, the infected RBCs (>5% parasitemia) were spinned down at

600×g and re-suspended in isoosmotic sorbitol solution (in mM: 290 sorbitol, 5 glucose, 5

HEPES/NaOH, pH 7.4) for 20 min at 21°C in continuous shaking. Then the cells were

washed twice in malaria culture medium and sub-culutred for further experiments

(Appendix-2 protocol 4).

2. 7. 1. 3. In vitro P. falciparum growth assay

For the in vitro growth assay, synchronized parasitized erythrocytes were aliquoted in 96-

well plates (200 µl aliquots,0.5- 1% hematocrit, 0.5 –2 % parasitemia) and grown for 48 h

in serum-free Albumax II (0.5%)-supplemented RPMI medium (159) (Appendix-2

protocol 5). The erythrocytes were grown in the presence or absence of different

concentrations of sodium butyrate or amphotericin B. The parasitemia of human

erythrocytes were assessed at time 0 and after 48 h of culture by flow cytometry (FACS

Calibur, Becton Dickinson, Heidelberg, Germany). Parasitemia was defined by the

percentage of erythrocytes stained with the DNA/RNA specific fluorescence dye Syto16

(20 nM final concentration, Molecular Probes, Göttingen, Germany). Briefly, RBCs were

incubated with Syto16, diluted in PBS or annexin binding buffer at 37°C. The staining

procedure was performed for around 30 – 40 min for infected human RBCs. Syto16 green

fluorescent nucleic acid stain bound to DNA has a maximum excitation/absorption

wavelength of 488 nm, which corresponds to the argon line of the single-laser of the FACS

Calibur used, and a maximum emission wavelength of 518 nm. Bound to RNA the

absorption maximum is at 494 nm, the emission maximum at 525 nm. This green emission

is detected in the Fluorescence 1 (FL-1) channel with a detector for an emission

wavelength of 530 ± 15nm. FACS analysis proved a more sensitive technique for

determining parasitemia than either Giemsa or Field´s rapid staining. (Appendix-2 protocol

6). In mice the course of parasitemia (=percentage of infected cells) was determined in

Giemsa stained blood films made from tail blood. The thick blood film was air-dried and

fixed with methanol. 2% Giemsa solution (Sigma) was added for 30 min. The slide was

rinsed with water and again dried. Then, the slides were analysed under a Leica CM E light

microscope (100 X, oil immersion) (204) (Appendix-2 protocol 7).

44 

 

2. 7. 2. Intraerythrocytic DNA amplification of P. falciparum

To estimate the DNA/RNA amplification in a further series of experiments, the culture was

ring stage-synchronized and re-synchronized after 6 h of culture (to narrow the

developmental parasite stage), aliquoted (200 µl aliquots, 2 % hematocrit and 10 %

parasitemia) and cultured for further 16 h. The erythrocytes were cultured in the presence

or absence of different concentrations of sodium butyrate or amphotericin B. Thereafter,

the DNA/RNA amount of the parasitized erythrocytes was determined by Syto16

fluorescence as a measure of intraerythrocytic parasite copies.

2. 7. 3. Annexin binding experiments (Determination of phosphatidylserine exposure)

Suspensions with non-infected RBCs were stained with annexin V-FLUOS. Suspensions

with P. falciparum infected RBCs were stained with annexin V-568 and/or with the DNA

dye Syto16 to assess PS exposure in the outer leaflet of the RBC membrane and the

percentage of infected RBCs, respectively (205). (Appendix-2 protocol 8)

For annexin binding, RBCs were washed, re-suspended in annexin-binding buffer, stained

with annexin V-568 (dilution 1:50) or annexin V-FLUOS (dilution 1:100), and incubated

for 20 min at RT in Ca2+-free annexin binding buffer. AnnexinV binding to negatively

charged phospholipids (with a high specificity for PS) is Ca2+-dependent. Therefore an

annexin binding buffer containing Ca2+ was used. Syto16 (final concentration of 30 nM)

was co-incubated in the annexin binding buffer for 20 min at RT for double-staining

purposes. Samples were diluted 1:5 with annexin binding buffer just before FACS analysis.

Cells were analyzed by flow cytometry. Fluorescence 1, FL-1 (detector for an emission

wavelength of 530 ± 15 nm) was either an indicator of annexin V-FLUOS fluorescence

intensity (excitation: 450 - 500 nm, emission: 515 -565 nm) (simple staining for annexin);

or of Syto16 fluorescence (maximum excitation wavelength: 488 nm for DNA, 494 nm for

RNA, maximum emission wavelength: 518 nm for DNA, 525 nm for RNA). Syto16

fluorescence was taken to reflect the degree of parasitemia. Fluorescence 2, FL-2 (detector

for an emission wavelength of 585 ± 21 nm) or FL-1 was used to determine the annexin

fluorescence of Annexin V-568 or Annexin Fluos, respectively, i.e. the percentage of PS

exposing cells. Annexin V-568 has an absorption spectrum from 450 – 650 nm, a

maximum at 578 nm, an emission spectrum from 560 nm up to > 700 nm with a maximum

at 603 nm. Double staining necessitated compensation for overlapping emission spectra by

45 

 

electronic subtraction of any unwanted spectral “spillover” from the signal of interest.

Simple stained infected erythrocytes (with Syto16 or annexin V-568) containing dye in the

relevant concentrations were used as compensation controls for this multicolor study.

To induce annexin binding according to Lang et al. (116), non-infected RBCs were

incubated for 1 h at 37 °C with the Ca2+ ionophore ionomycin (0 and 1 µM, respectively)

or oxidized by t-butylhydroperoxide (t-BHP, 1 mM for 15 min followed by 24h of post-

incubation) in a modified NaCl test solution consisting of ( in mM): 125 NaCl, 5 KCl, 5 D

(+) glucose, 1 CaCl2, 1 MgSO4, 32 HEPES titrated to pH 7.4 with NaOH.

2. 7. 4. Determination of intracellular Ca2+

influx in the erythrocytes

Fluo 3 acetoxymethyl AM ester was used as the fluorescent indicator. Fluo-3 (1- [2-amino-

5-(2,7-dichloro-6-hydroxy-3-oxo-9-xanthenyl) phenoxy] -2-(2-amino-5-methylphenoxy)

ethane-N,N,N’,N’-tetra-acetic acid, C51H50C12N2O23 =1129.85) is a fluorescent cheater

excited by visible light (~488nm), and emits a yellowish green fluorescence (~525nm)

when bound to a calcium ion (198). The intensity of fluorescence depends on the free

calcium concentration. Fluo-3 does not fluoresce unless bound to calcium ions.

Intracellular Ca2+

measurements were performed as described previously by Andrews et

al., (2002) (206). Erythrocytes were loaded with fluo-3 AM (Calbiochem, Bad Soden,

Germany) by addition of fluo-3 AM stock solution (2 mM diluted in DMSO) to 1 ml of

erythrocyte suspension (0.16% hematocrit in Ringer solution; 4 µM fluo-3 AM final

concentration). The cells were incubated at 37°C for 15 min under protection from light.

An additional 2-µl aliquot of fluo-3 AM was added, and then the mixture was incubated for

25 min. Fluo-3-loaded erythrocytes were centrifuged at 1,000 g for 5 min at 22°C and then

washed twice with Ringer solution containing 0.5% bovine serum albumin (Sigma) and

once with Ringer solution and incubated. For flow cytometry, fluo-3-loaded erythrocytes

were re-suspended in 1 ml of Ringer solution (0.16% hematocrit) containing serial

concentrations of amphotericin B or sodium butyrate and incubated at 37°C for 30 min. As

a positive control, erythrocytes were stimulated with 1 µM Ca2+

ionophore ionomycin

(Sigma) for 3 min prior to analysis to increase intracellular Ca2+

activity. For negative

control, cells were incubated for 30 min at 37°C with vehicle alone. Subsequently, Ca2+

-

46 

 

dependent fluorescence intensity was measured in fluorescence channel FL-1 with an

excitation wavelength of 488 nm and an emission wavelength of 530 nm (115).

2. 7. 5. Measurement of hemolysis

After incubation of infected erythrocytes for 48 hours at 37°C, the samples were

centrifuged (3 min at 400 g, RT), and the supernatants were harvested. As a measure of

hemolysis, the hemoglobin (Hb) concentration of the supernatants was determined

photometrically at 405 nm. The absorption of the supernatant of erythrocytes lysed in

distilled water was defined as 100% hemolysis (207, 208) (Appendix-2 protocol 9).

2. 7. 6. In vivo proliferation of P. berghei ANKA

For infection of mice P. berghei ANKA-parasitized mouse erythrocytes (1x106) were

injected intraperitoneally (209) into sex- and age-matched mice fed on control diet and

with free access to drinking water or 10% (w/w) GA dissolved in tap water (100 g/L) or

amphotericin B (1.4 mg/kg body weight for three successive days after day 8) administered

subcutaneously. Parasitemia was determined daily from the 8th

day of infection by flow

cytometry. The blood samples were stained with Giemsa stain and the DNA/RNA-specific

fluorescence dye Syto-16 and the Syto-16 fluorescence was analysed with a FACS Calibur

cytometer (Becton Dickinson , Heidelberg, Germany) at fluorescence channel 1 (FL1 488

nm excitation and 530 nm emission as described above).

2. 7. 7. Measurement of the in vivo clearance of fluorescence-labelled erythrocytes

Fluorescence-labelled erythrocytes from mice were obtained by staining erythrocytes with

carboxyfluorescein diacetate, succinimidyl ester (CFSE) from Molecular Probes (Leiden,

The Netherlands). The labelling solution was prepared by addition of adequate amounts of

a CFSE stock solution (10 mM in DMSO) to phosphate-buffered saline (PBS) to yield a

final concentration of 5 µM. Then, the cells were incubated with labelling solution for 30

min at 37°C. The cells were pelleted at 1000 x g for 5 min, washed and re-suspended in

fresh, pre-warmed PBS. The fluorescence-labelled erythrocytes were injected into the tail

veins of mice. At indicated time points, blood was taken from the mice and CFSE-

dependent fluorescence intensity of the erythrocytes was measured in the fluorescence

channel FL-1. The percentage of CFSE-positive erythrocytes was calculated in % of the

whole population (169).

47 

 

2. 7. 8. Measurement of the fluorescence-labelled erythrocytes in the spleens of mice

The fluorescence-labelled erythrocytes were injected into the tail veins of mice. After 2

hours the mice were sacrificed and the spleens were dissected and carefully mashed

through a net. Finally, CFSE-dependent fluorescence intensity of the erythrocytes in spleen

suspensions was measured in the fluorescence channel FL-1 (209).

2. 7. 9. Data analysis and statistics

Data are expressed as means ± SEM, and (n) represents the number of independent

experiments. Statistical analysis was made by paired or unpaired t test or by ANOVA

using Dunnett’s or Tukey’s test as post hoc test, where appropriate. P ≤ 0.05 was

considered statistically significant. Statistical analysis was performed with GraphPad

InStat version 3.00 for Windows (GraphPad Software, San Diego, CA;

www.graphpad.com).

48 

 

3. Results

3. 1. Influence of amphotericin B on the course of malaria Effect of amphotericin B on in vitro growth of P. falciparum in human erythrocytes.

In vitro growth of P. falciparum in human erythrocytes was significantly decreased at

amphotericin B started from concentration of 0.01 µM (Fig. 3. 3). Within 48 hours the

percentage of infected erythrocytes increased from 4 to 23%. Increased amphotericin B

concentration decreases percentage parasitemia in a dosage dependant manner, an effect

reaching statistical significance at ≥ 0.01 µM amphotericin B concentrations (p< 0.05).

Effect of amphotericin B on P. falciparum DNA amplification

Intraerythrocytic DNA amplification in erythrocytes was not significantly influenced by

amphotericin B at the concentrations employed but only at the highest one (1µM). (Fig. 3.

4).

Effects of amphotericin B on erythrocytes hemolysis.

At the applied concentrations amphotericin B did not disrupt the integrity of the cell

membrane and did not induce significant hemolysis (Fig. 3. 5).

Fluo 3 fluorescence intensity in control, ionophore ionomycin and amphotericin B treated erythrocytes Erythrocyte shrinkage induces activation of Ca

2+-permeable non-selective cation channels.

Thus erythrocytes were loaded with the Ca2+

-sensitive fluorescence dye fluo-3 (2 µM fluo-

3 AM) to determine the effect of amphotericin B on cytosolic Ca2+

activity. Cells were

incubated in the absence or presence of amphotericin B (1 µM for 30 min) or the Ca

2+

ionophore ionomycin (1 µM), and fluo-3 fluorescence intensity was monitored using

FACS. As illustrated in Fig. 3. 6, amphotericin B increased fluo-3 fluorescence intensity.

As a positive control, the Ca2+

ionophore ionomycin (1 µM) similarly enhanced the fluo-3

fluorescence intensity. The data strongly suggest that the observed amphotericin B effect

on fluo-3-loaded erythrocytes reflect an increase in cytosolic free Ca2+

concentration.

Fluo 3 fluorescence intensity in control and amphotericin B treated erythrocytes

Incubation of erythrocytes in Ringer solution with amphotericin B for 48 h indicates

significant difference from the respective value of Fluo 3 fluorescence intensity in the

absence of amphotericin B (Fig. 3. 7). Fluo3 fluorescence intensity increased by increased

49 

 

amphotericin B concentration. It was found to be 19.73 (arb. units) in the absence of

amphotericin B, increases in Fluo3 fluorescence reaching statistical significance at ≥ 0.1

µM amphotericin B concentration (p< 0.01)..

Effects of amphotericin B on phosphatidylserine exposure of P. falciparum infected and non-infected erythrocytes.

To identify phosphatidylserine exposing erythrocytes, annexin V binding has been

determined in FACS analysis. The percentage of annexin-binding erythrocytes was low in

fresh erythrocytes (1.24 ± 0.24%, n = 6). Exposure of uninfected erythrocytes to

amphotericin B-free Ringer solution increased the percentage of annexin binding cells

within 24 h to 4.14 ± 0.50% (n = 6) (Fig. 3. 8 A). Infection of erythrocytes with P.

falciparum significantly enhanced the percentage of annexin binding erythrocytes.

Phosphatidylserine exposure of infected erythrocytes was augmented by amphotericin B,

an effect reaching statistical significance at 0.01 µM amphotericin B (Fig. 3. 8 B).

Effects of amphotericin B on forward scatter of infected and non-infected erythrocytes.

Infection of human erythrocytes with P. falciparum induces a biphasic change of the host

erythrocyte volume. In the early trophozoite stage infected cells loose KCl and water

leading to cell shrinkage. In the present study, ring-stage-synchronized erythrocytes when

grown for 48h re-invaded new erythrocytes in which the parasite at the early trophozoite

stages. Accordingly, infected cells had a lower forward scatter than uninfected

erythrocytes. Amphotericin B leads to farther significant decrease in infected erythrocyte

volume illustrated in Fig. 3. 9 A & B, which shows decreased forward scatter.

In vivo proliferation of P. berghei in control and amphotericin B treated mice.

Further experiments were performed on the impact of amphotericin B treatment on the

course of malaria infection in vivo. To this end mice were infected with P. berghei. As

shown in Fig. 3. 10, parasitemia in control and amphotericin B mice was (2.61 ± 0.18 and

2.33 ± 0.12%) respectively 8 days after infection but gradually increased in the next 12

days. In untreated animals, the percentage of parasitized erythrocytes increased up to 50.36

± 3.98%. The addition of 0.7 mg/kg b.w. amphotericin B had only a slight effect on

parasitemia. Thus, the concentration of amphotericin B was increased to 1.4 mg/kg b.w.,

which indeed significantly blunted the increase of parasitemia compared to non treated

animals to 39 ± 2.46 % (mean ± SEM, n = 6), (p<0.05) (Fig. 3. 11).

50 

 

Clearance of amphotericin treated erythrocytes from circulating blood

A next series of experiments explored, whether the enhanced phosphatidylserine exposure

accelerates clearance of erythrocytes from circulating blood in vivo. For this purpose mice

were either left untreated or treated with 1.4 mg/Kg b w amphotericin B intraperitoneally,

their erythrocytes were labelled with the dye CFSE and injected into the tail vein. As

illustrated in Fig. 3. 12 A, the clearance of erythrocytes in amphotericin B treated mice was

slightly, but significantly faster than erythrocyte clearance in untreated mice. On the other

hand, the fraction of labelled erythrocytes recovered in the spleen was significantly

enhanced by amphotericin B treatment (Fig. 3. 12 B).

The effect of amphotericin B on survival of P. berghei infected mice.

The effect of amphotericin B on the parasitemia was paralleled by enhanced survival of the

amphotericin B treated animals. Whereas all untreated animals died within 27 days after

the infection, 80% of the amphotericin B treated animals survived the infection for more

than 27 days (Fig. 3. 13). Thus, amphotericin B treatment indeed significantly modified the

course of malaria.

3. 2. Influence of butyrate and gum Arabic on the course of malaria Effect of butyrate on in vitro growth of P. falciparum in human erythrocytes.

The influence of butyrate on the in vitro growth of the parasite was analyzed. P.

falciparum-infected erythrocytes were cultured in human erythrocytes and synchronized to

ring stage by sorbitol treatment. Within 48 hours the percentage of infected erythrocytes

increased from 5 to 26 %. Increased butyrate concentration decreases percentage

parasitemia in a dosage dependant manner, an effect reaching statistical significance at ≥

0.5 mM butyrate concentration (Fig. 3. 14) (p< 0.05).

Effect of butyrate on P. falciparum DNA amplification.

To test for intraerythrocytic DNA amplification infected erythrocytes were ring-stage

synchronized and cultured for further 24h. As a result of increased butyrate concentration,

statistically significant decrease in DNA amplification was observed only at the highest

concentration (10 mM) (Fig. 3. 15).

51 

 

Effect of butyrate on erythrocytes cytosolic Ca2+ concentration.

Activation of cation channels is expected to increase cytosolic Ca2+ activity. Fluo3

fluorescence was employed to test, whether butyrate modifies the cytosolic Ca2+

concentration. As illustrated in Fig. 3. 16, 48 hours incubation of erythrocytes in the

presence of different concentrations of amphotericin B was followed by a significant

increase in Fluo3 fluorescence. Accordingly, exposure to amphotericin B increases

cytosolic Ca2+ activity of human erythrocytes.

Effects of butyrate on phosphatidylserine exposure of infected and non-infected erythrocytes.

To determine the effect of infection and butyrate on suicidal erythrocyte death (eryptosis),

the percentage of phosphatidylserine-exposing erythrocytes was estimated by measurement

of annexin V-binding in FACS analysis. Prior to the infection the percentage of annexin V-

binding erythrocytes was low (1.2 ± 0.11%, n = 6). Within 24 hours, infection with P.

falciparum markedly increased annexin V-binding of infected erythrocytes compeered to

non infected mean ± SEM, (26.5 ± 4.3 and 15.9 ± 3.2) respectively (p≤0.05; paired

ANOVA). Both in the absence and presence of butyrate the percentage of annexin V-

binding was significantly higher in infected than non-infected erythrocytes. The

phosphatidylserine exposure of infected erythrocytes was significantly increased by

increased butyrate concentration, an effect reaching statistical significance at 5 mM

butyrate (p≤0.05; paired ANOVA) (Fig. 3. 17).

Effects of butyrate on forward scatter of infected and non-infected erythrocytes.

At early stages of the parasite development, infection of erythrocytes decreased erythrocyte

forward scatter, pointing to cell shrinkage. At late stages of the parasite development

(trophozoites), infection increased the forward scatter, pointing to cell swelling. Butyrate

decreased the forward scatter of the early and late stage infected erythrocytes. Fig. 3. 18 A

& B shows significant difference (p < 0.05) from absence of butyrate at early stages, and

also indicates significant difference to non-infected erythrocytes (p < 0.05).

In vivo effect of gum Arabic on parasitemia of P. berghei infected mice.

To determine the in vivo efficacy of gum Arabic, mice were infected with P. berghei with

or without gum Arabic treatment. Gum Arabic (10 % in drinking water) was administered

52 

 

from the 8th day of infection, when parasitemia was still low in control and infected mice,

mean ± SEM, (3.44 ± 0.63 and 5.09 ± 0.21%) respectively, no significant deferens was

observed. The percentage of infected erythrocytes increased gradually in mice without or

with gum Arabic treatment reaching 43.6 ± 4.39 and 58.08 ± 3.03 respectively at day 21

(Fig. 3. 19). However, the percentage of parasitized erythrocytes was slightly lower in gum

Arabic-treated mice than in mice without gum Arabic treatment, an effect reaching

statistical significance between day 16 to day 20 of infection (p≤0.05; paired ANOVA)

(Fig. 3. 19).

Effects of gum Arabic on survival of P. berghei infected mice.

The treatment with gum Arabic further influenced the survival of P. berghei-infected mice.

As shown in Fig. 3. 20, all of the gum treated animal survived up to day 21 after the

infection, were the untreated animals number decreased by 30 %. All of the untreated

animals died within 26 days after the infection. In contrast as many as 70% of the gum

Arabic-treated animals survived the infection for more than 26 days.

53 

 

Fig. 3. 3: In vitro growth of Plasmodium falciparum in human erythrocytes as a function of

amphotericin B concentration.

Arithmetic means ± SEM (n = 8) of in vitro growth of Plasmodium falciparum in human

erythrocytes as a function of amphotericin B concentration. *, **Indicates significant difference (p

< 0.05; p < 0.01) from absence of amphotericin B.

54 

 

Fig. 3. 4: DNA amplification of Plasmodium falciparum in human infected erythrocytes as a function of amphotericin B concentration (arithmetic means ± SEM, n = 8). *Indicates significant difference (p < 0.05;) from absence of amphotericin B.

55 

 

0

1

2

0 0.001 0.01 0.05 0.1 1

Hem

olys

is [%

]

Conc. of amphotericin B µM

Fig. 3. 5: Effect of amphotericin B on erythrocyte membrane integrity. Arithmetic means ± SEM (n= 4) of the percentage of haemolysed erythrocytes exposed for 48 hours to Ringer solution without (white bar) or with (black bars) amphotericin B at the indicated concentrations.

56 

 

Fig. 3. 6: Effect of amphotericin B and ionomycin cytosolic fluo-3 fluorescence intensity in non-infected erythrocytes. Mean Ca

2+-sensitive fluo-3 fluorescence

(arithmetic means ± SE; n = 6) of erythrocytes incubated either in Ringer solution (Control) or in Ringer solution containing amphotericin B

(1 µM) or ionomycin (1 µM). *

Indicates significant difference (p < 0.05) from control.

57 

 

0

20

40

60

80

0 0.01 0.05 0.1 1

Fluo

-3 fl

uore

scen

ce [a

rb. u

nits

]

amphotericin B µM

** **

Fluo

-3 fl

uore

scen

ce [a

rb. u

nits

]

** **

Fig. 3. 7: Effect of amphotericin B on cytosolic Ca2+ concentration in non-infected erythrocytes. Arithmetic means ± SEM (n = 6) of the geo. means of Fluo3 fluorescence in erythrocytes exposed for 48 hours to Ringer without (white bar) or with (black bars) different concentrations of amphotericin B . ** (p<0.01) indicates significant difference from the respective value in the absence of amphotericin B.

58 

 

0

1

2

3

4

5

Fresh 24 h

Ann

exin

bin

ding

[%]

#A)

0

10

20

30

0 0.001 0.01 0.05 0.1 1

Ann

exin binding

 [%]

Conc. of amphotericin B (µM)

Non-infectedInfected

*

* *B)

*

Fig. 3. 8 A & B: Annexin binding of non-infected and Plasmodium falciparum infected erythrocytes from control and amphotericin B treated samples. A) Arithmetic means ± SEM (n = 4) of annexin binding to erythrocytes in freshly drawn blood (open bar) and 24 hour after incubation in RPMI 1640 medium containing 0.05 µM amphotericin B (closed bar) #Indicates significant difference between two groups(p ≤ 0.01). B) Arithmetic means ± SEM (n = 4) of annexin binding of infected (closed bar) and non-infected (open bar) erythrocytes before and after treatment with different concentrations of amphotericin B. *Indicates significant difference (p < 0.05) to non-infected erythrocytes.

59 

 

200

250

300

350

400

0 0.001 0.01 0.05 0.1 1

Forw

ard

Scat

ter

[rel

.uni

ts]

Conc. of amphotericin B (µM)

A)* #

* #A)* #

* #A)# *

# *# **

# *

200

240

280

320

360

0 0.001 0.01 0.05 0.1 1

Forw

ard

Scat

ter

[rel

.uni

ts]

Conc. of amphotericin B (µM)

Noninfected

Infected

* # * #* # * #

B)

Fig. 3. 9 A & B: Effects of amphotericin B on forward scatter of Plasmodium falciparum infected and noninfected erythrocytes. A). Arithmetic means ± SEM (n = 6) forward scatter of the early stage-infected erythrocytes (closed symbols) and non-infected (open symbols) erythrocytes as a function of the butyrate concentration. * indicates significant difference (p≤0.05; ANOVA) from absence of butyrate, # indicate significant difference (p≤0.05; ANOVA) from non-infected erythrocytes. B). Arithmetic means ± SEM (n = 6) forward scatter of late stage-infected erythrocytes (closed symbols) and non-infected (open symbols) erythrocytes as a function of the butyrate concentration. * indicates significant difference (p≤0.05; ANOVA) from absence of butyrate. # indicates significant difference to non-infected erythrocytes.

60 

 

Con

trol

amph

oter

icin

B

Day 20 Day 8

Fig. 3. 10: Effect of amphotericin B treatment on parasitemia of Plasmodium berghei infected mice: Original histograms of parasitemia-dependent Syto 16 fluorescence in untreated animals (upper panels) and animals treated from day 8 until day 20 with 1.4 mg amphotericin B intraperitoneally (lower panels) 10 (left panels) and 20 (right panels) days after infection with P. berghei.

61 

 

Fig. 3. 11: Effect of amphotericin B on parasitemia of P. berghei infected mice. Arithmetic means ± SEM (n = 6) of parasitemia in mice without treatment (open circles) or treated with amphotericin B (closed circles) as a function of days after infection with P. berghei. * indicates significant difference (p ≤ 0.05) from iron replete animals.

62 

 

0

2

4

6

8

0 10 30 40 60 80 100 120

CFS

E p

ositi

ve c

ells

[%]

Time (min)

NoninfectedInfected

A)

*

* * * * *

0

4

8

12

16

CFS

E p

ositi

ve c

ells

[%]

ControlAmphotericin B

B)*

Fig. 3. 12 A & B: In vivo clearance of fluorescence labelled erythrocytes from circulating blood. Disappearance of CFSE labelled non infected and infected mouse erythrocytes from circulating blood following injection into the tail vein of untreated and amphotericin B (1.4 mg i.p.) treated mice. A): Arithmetic means ± SEM (n = 6) of the percentage of CFSE-labelled non-infected erythrocytes in circulating blood of untreated (open symbols) or amphotericin B (closed symbols) treated animals. * indicates significant difference (p<0.05) from untreated animals. B): Abundance of CFSE labelled untreated (open bar) or amphotericin B treated (closed bar) erythrocytes in spleen. Arithmetic means ± SEM (n=6). *indicates significant difference from the untreated animals.

63 

 

0

20

40

60

80

100

8 11 14 17 20 23 26 29

Surv

ival

[%]

Days after infection

control

Amphotericin B

Fig. 3. 13: Effect of amphotericin B on survival of P. berghei infected mice. Arithmetic means ± SEM (n = 6) represent survival of mice without treatment (open circles) or treated with amphotericin B

(closed squares) as a function of days after infection with P. berghei.

64 

 

Fig. 3. 14: In vitro growth of P. falciparum in human erythrocytes as a function of butyrate concentration (arithmetic means ± SEM, n = 8). *, ** indicates significant difference (p≤0.05, p≤0.01) from absence of butyrate.

65 

 

Fig. 3. 15: Intraerythrocytic DNA amplification of P. falciparum in human erythrocytes as a function of butyrate concentration (arithmetic means ± SEM, n = 6). * indicates significant difference (p≤0.05) from absence of butyrate.

66 

 

Fig. 3. 16: Effect of butyrate on cytosolic Ca2+ activity in non-infected erythrocytes. Arithmetic means ± SEM (n = 6) of the geometric means of Fluo3 fluorescence in erythrocytes exposed for 48 hours to Ringer without (white bar) or with (black bars) different concentrations of butyrate. * (p<0.05) indicates significant difference from the respective value in the absence of butyrate.

67 

 

Fig. 3. 17: Effects of butyrate on phosphatidylserine exposure of infected and non-infected erythrocytes. Arithmetic means ± SEM (n = 6) of annexin V-binding of infected (closed bars) and non-infected (open bars) erythrocytes following infection of human erythrocytes with P. falciparum at 0 mM (left bars) and 10 mM butyrate. * indicates significant difference (p≤0.05; paired ANOVA) from non-infected erythrocytes, # *Indicates significant difference (p < 0.05) from absence of butyrate

 

68 

 

280

300

320

340

360

0 0.3 0.5 1 3 5 7 10

Forw

ard

Scat

ter

(rel

.uni

ts)

Conc. of Butyrate (mM)

NoninfectedInfected early stages

* # * # * # * ##           #            #

#

A)

280

300

320

340

360

0 0.3 0.5 1 3 5 7 10

Forw

ard

Scat

ter

(rel

.uni

ts)

Conc. of Butyrate (mM)

NoninfectedInfected late stagesB)

* # * # * # * # * #

Fig. 3. 18 A & B: Effects of butyrate on forward scatter of infected and non-infected erythrocytes. A): Arithmetic means ± SEM (n = 4) of forward scatter of the early stage-infected erythrocytes (closed symbols) and non-infected (open symbols) erythrocytes as a function of the butyrate concentration. *Indicates significant difference (p < 0.05) from absence of butyrate, #Indicates significant difference to non-infected erythrocytes. B): Arithmetic means ± SEM (n = 4) of forward scatter of late stage-infected erythrocytes (closed symbols) and non-infected (open symbols) erythrocytes as a function of the butyrate concentration. *Indicates significant difference (p < 0.05) from absence of butyrate, #Indicates significant difference to non-infected erythrocytes.

69 

 

0

10

20

30

40

50

60

70

8 10 12 14 16 18 20 22

Para

sitem

ia(%

)

Days after infection

Control

Gum arabica

**

**

*

Fig. 3. 19: Parasitemia of Plasmodium berghei infected mice. Arithmetic means ± SEM of parasitemia in mice without treatment (open circles, n = 8) or with 10 mg/kg B. W. of gum Arabic (closed circles, n = 6) as a function of days after infection with Plasmodium berghei. *Indicates significant difference from the untreated animals.

70 

 

Fig. 3. 20: Effect of gum Arabic treatment on survival of Plasmodium berghei infected mice. Survival of mice without treatment (open circles) or with 10 % gum Arabic in drinking water (closed squares) as a function of days after infection with Plasmodium berghei.

71 

 

4. Discussion

Over the past centuries malaria, has been the major cause of morbidity and mortality

among the mankind. As the time passed the pathogens coevolved with their hosts,

simultaneously manipulating the hosts defence mechanisms by intruding and mimicking

various host related metabolic and signalling pathways. In the course of their parasitic

effect they modified themselves to proliferate and destruct the host to death. The present

strategy for controlling and curing infectious diseases has targeted various metabolic or

enzymatic systems within the parasite. The most severe drawback of this kind of

controlling the diseases has lead to the development of parasitic resistance and consequent

relapse of once-contained infectious diseases amidst the host. This study intended for a

novel drug discovery in order to prevent the pathogen related resistance focusing on

identifying and targeting host factors essential for pathogen entry, survival and

proliferation. The innovative methods involved stimulation of the infected erythrocytes and

recognition by the spleen macrophages to get rid of the pathogen and prevent the further

course of the disease.

Malaria pathogen, Plasmodium, enters erythrocytes and thus escapes recognition by the

immune system. The pathogen induces oxidative stress to the host erythrocyte, which

triggers eryptosis, the suicidal death of erythrocytes (167, 178). Eryptosis is characterized

by cell shrinkage, membrane blebbing and cell membrane phospholipids scrambling with

phosphatidylserine exposure at the cell surface. Phosphatidylserine-exposing erythrocytes

are identified by macrophages which engulf and degrade the eryptotic cells. To the extent

that infected erythrocytes undergo eryptosis prior to subsequent infection of other

erythrocytes by Plasmodia, hence, the premature eryptosis may protect against malaria.

Accordingly, any therapeutic intervention accelerating suicidal death of infected

erythrocytes has the potential to foster elimination of infected erythrocytes, delay the

development of parasitemia and favourably influence the course of malaria.

Observations of this study indicate that Plasmodium infected erythrocytes can be targeted

for induction of eryptosis in vitro and in vivo. This accelerated eryptosis of Plasmodium-

infected erythrocytes results in rapid clearance of infected erythrocytes and confers

protection against severe course of malaria. Plasmodium imposes oxidative stress to the

host cell leading to cation channel activation and Ca2+

entry (210, 211). Thus, infection was

72 

 

expected to trigger phosphatidylserine exposure. Infection of erythrocytes with

Plasmodium was indeed found to trigger eryptosis (135). If the stimulation of eryptosis

remains an abated, it may proceed too fast for the pathogen to allow intraerythrocytic

maturation.

4. 1. Influence of amphotericin B on the course of malaria

According to the present observations, amphotericin B in different concentration exerted

an inhibitory effect on intraerythrocytic P. falciparum proliferation in vitro.(Fig. 3. 3)

Moreover, presence of amphotericin B further decreased the intraerythrocytic DNA

amplification of the parasite, an effect reaching statistical significance only at the highest

concentration in the experiment (Fig. 3. 4). In theory, the positive effect of amphotericin B

treatment could have been due to amphotericin B induced lysis of the trophozoite stage of

P. falciparum ](212). However, at the applied concentrations in this study, amphotericin B

did not disrupt the integrity of the cell membrane and did not induce significant hemolysis

(Fig. 3. 5). The concentrations required to elicit eryptosis are well within the range of those

reached under treatment with amphotericin (213, 214). Eryptosis clearly exceeds hemolysis

and thus, the observed phosphatidylserine exposure is not secondary to hemolysis. Higher

levels of hemolysis have been reported earlier following exposure of erythrocytes to

amphotericin B formulations (215).

P. falciparum survival depends on the activation of several ion channels in the erythrocyte

cell membrane, as they allow the uptake of nutrients, Na+ and Ca

2+ and the disposal of

waste products (172). Increases in cytosolic Ca2+ activity is one of the major stimulators of

eryptosis, which leads to cell membrane vesiculation, stimulates cell membrane scrambling

and activates the cysteine endopeptidase calpain, an enzyme degrading the cytoskeleton

and thus causing cell membrane blebbing (216). Ca2+

may enter through non-selective

cation channels (132). The molecular identity of those channels is incompletely understood

but may include transient receptor potential channels (TRPC6). The cation channels are

activated by osmotic shock, oxidative stress and Cl- removal (134).

In this study to determine the effect of amphotericin B on cytosolic Ca2+

activity, cells

were incubated in the absence or presence of amphotericin B or the Ca2+

ionophore

73 

 

ionomycin, and fluo-3 fluorescence intensity was monitored using FACS. As illustrated in

Fig. 3. 6, amphotericin B indeed increased fluo-3 fluorescence intensity. As a positive

control, the Ca2+

ionophore ionomycin similarly enhanced the fluo-3 fluorescence

intensity. The data strongly suggest that the observed amphotericin B effect on fluo-3-

loaded erythrocytes reflect an increase in cytosolic free Ca2+

concentration. Moreover,

amphotericin B farther increased fluo-3 fluorescence intensity in a dose dependant manner

(Fig. 3. 7). These results were in line with studies by Lang et al, which demonstrated the

effect of amphotericin B on cytosolic free Ca2+ (205, 208).

An increase in cytosolic Ca2+-activity is expected to stimulate cell membrane scrambling

with phosphatidylserine exposure, which could be quantified by determination of annexin

V-binding. As shown in Fig. 3. 8 A, the percentage of phosphatidylserine-exposing

erythrocytes was low in the absence of amphotericin B. Exposure of erythrocytes for 48

hours to Ringer solution containing amphotericin B (0.1 µM) was followed by a significant

increase in annexin V-binding. Phosphatidylserine exposure was used as indicator of

apoptosis (217). In this study amphotericin B was found to increase the percentage of

annexin V binding which indicate phosphatidylserine exposure. As shown in Fig. 3. 8 B.

Furthermore, the effect was particularly prominent in infected erythrocytes. Thus,

amphotericin B preferably triggered phosphatidylserine exposure of infected cells.

Infection of human erythrocytes with P. falciparum induces a biphasic change of the host

erythrocyte volume. In the early trophozoite stage infected cells loose KCl and water

leading to cell shrinkage. An increase in cytosolic Ca2+

activity is further expected to

induce cell shrinkage, which should be reflected by a decrease of forward scatter in FACS

analysis. Amphotericin B further decreased forward scatter pointing to cell shrinkage, the

effect which is more prominent in early stages P. falciparum infected erythrocytes (Fig. 3.

9 A & B). The present experiments confirm the ability of amphotericin B to induce

eryptosis (207). The decrease in forward scatter during eryptosis is paralleled by a decrease

of packed cell volume and thus indeed reflects cell shrinkage (218). It was early shown

that, the decrease in forward scatter is reversed by increased extracellular K+ concentration,

which dissipates the chemical driving force for K+ exit. The decrease in forward scatter is

further blunted by inhibitors of Ca2+-sensitive K+ channels. Thus the decrease in forward

74 

 

scatter following entry of Ca2+ is most likely due to activation of Ca2+-sensitive K+

channels, with subsequent exit of K+, hyperpolarization, and exit of Cl– and H2O (144).

The ability of amphotericin B to trigger scrambling of the cell membrane is shared by

several amphiphiles (219).

Obviously, increasing cytosolic Ca2+ concentrations leads to erythrocyte hyperpolarization,

and activation of Ca2+-sensitive K+ channels (Gardos channels). The parallel stimulation of

erythrocyte scramblase and Ca2+-sensitive K+ channels by an increase of cytosolic Ca2+

activity presumably serves to circumvent hemolysis of defective erythrocytes. Leakage of

the cell membrane or impairment of Na+-K+-ATPase is expected to result in gain of NaCl,

cell swelling, and, eventually, cell lysis (220). Even in normal erythrocytes, cytosolic Cl-

activity is high, and the potential difference across the cell membrane is low. Thus the gain

of Na+ and loss of K+ must be excessive to compromise volume constancy. Prior to that the

leaky cell membrane increases cytosolic Ca2+ activity which activates the erythrocyte

scramblase, leading to breakdown of phosphatidylserine asymmetry. The

phosphatidylserine-exposing erythrocytes are detected by the macrophages and, thus,

cleared from the circulation before hemolysis. The parallel activation of the Ca2+-sensitive

K+ channels, hyperpolarization, and cellular loss of K+ and Cl- counteract cell swelling and,

thus, may serve to prevent premature lysis of the erythrocytes (114).

The treatment of P. berghei infected mice with amphotericin B significantly decreased the

development of in vivo parasitemia (Fig. 3. 10 & 3. 11). The effect is presumably due to

premature phosphatidylserine exposure of infected cells. In vitro, phoshpatidylserine

exposure does not lead to sequestration and clearance of erythrocytes and thus does not

significantly modify the increase of parasitemia. Amphotericin B treated

phosphatidylserine exposing erythrocytes are rapidly cleared from circulating blood. In

nucleated cells amphotericin B has been demonstrated to enhance the cation conductance

in the plasma membrane (221) with subsequent Na+ entry and increase in the intracellular

Ca2+ concentration (222). It has further been shown to trigger a signaling cascade involving

TLR-2, Btk, PLC, PKC, c-Src and NF-kappaB (223, 224).

A next series of experiments explored, whether the enhanced phosphatidylserine exposure

accelerates clearance of erythrocytes from circulating blood in vivo. For this purpose mice

75 

 

were either left untreated or treated with 1.4 mg/Kg b. w. amphotericin B., their

erythrocytes were labelled with the dye CFSE and re-injected into the tail vein. As

illustrated in Fig. 3. 12 A, the clearance of erythrocytes in amphotericin B treated mice was

slightly, but significantly faster than erythrocyte clearance in untreated mice. Most

importantly, the decline was particularly fast in infected CFSE-labelled erythrocytes (Fig.

3. 12 A). On the other hand, the majority of cleared labelled cells were accumulated in the

spleen (Fig. 3. 12 B).

Decreased in vivo parasitemia following amphotericin B treatment may be expected to

affect the survival of infected animals. Similar to previous report (212), none of the

untreated mice survived the infection with P. berghei (Fig. 3. 13). In contrast, 80 % of the

amphotericin B treated mice were still alive 28 days after the infection illustrating that

amphotericin B treatment indeed favourably modifies the course of malaria.

4. 2 Influence of butyrate and gum Arabic on the course of malaria

In the other part of this study butyrate in different concentrations inhibited the proliferation

of P. falciparum in a dosage dependant manner (Fig. 3. 14). This effect is most probably

due to the activity of butyrate as histone deacetylase inhibitor reported by Davie

2003.(225). Andrews et al reported the antimalarial activity of histone deacetylase

inhibitors and cyto-differentiating agents in vitro and in vivo (226). Histones are nuclear

core proteins act as spools around which DNA winds, they play an important role in

transcriptional regulation through the continuous acetylation/deacetylation of the έ-amino

group of specific lysine residues (225). Evert et al (2006) reported that butyrate alter gene

expression and arrest cell proliferation by inhibition of the chromatin-remodelling activity

of histone deacetylases (HDAC) (227). Interestingly, butyrate inhibitory action of HDAC

activity was found to affect the expression of only 2% of genes (225). The findings of this

study were in parallel with that, as butyrate exerted very mild effects on P. falciparum

DNA amplification only at highest concentrations (10 mM) (Fig. 3. 15). Butyrate farther

increased fluo-3 fluorescence intensity in a dose dependant manner (Fig. 3. 16). Increased

fluo 3 fluorescence intensity was found to be directly proportional to cytosolic Ca2+

concentration (207).

76 

 

On the other hand, the presence of butyrate significantly increased the percentage of

annexin V binding erythrocytes. The effect was particularly prominent in infected

erythrocytes (Fig. 3. 17). Thus, butyrate preferably triggered phosphatidylserine exposure

of infected cells. The specific binding of annexin V to phosphatidylserine has greatly

facilitated the measurement of the exposure of small amounts of this phospholipid on the

outer membrane of cells. Annexin V can be labelled with a fluorescent compound; this will

make it possible to use a flow cytometer to estimate the percentage of erythrocytes with

phosphatidylserine exposed on their outer surface. Several lines of evidence reported by a

number of investigators indicated that exposure of phosphatidylserine on the cell surface is

a general feature of apoptosis and suicidal erythrocyte death (eryptosis) (228, 229). As

shown earlier (179, 230), eryptosis is further stimulated by P. falciparum infection.

Intraerythrocytic Plasmodia impose oxidative stress to the host cell, which in turn activates

the Ca2+ permeable cation channels (231). The following Ca2+ entry leads to stimulation of

cell membrane scrambling (232).

Butyrate further decreased forward scatter pointing to cell shrinkage (Fig. 3. 18), which

was presumably due to increase of cytosolic Ca2+

activity, activation of Ca2+

sensitive K+

(GARDOS) channels (144), K+

exit, hyperpolarization of the erythrocyte membrane

potential and Cl- exit (143). The exit of KCl is followed by osmotically obliged water and

thus leads to cell shrinkage (170). The volume regulatory cation channels are not only

expressed in erythrocytes but in several nucleated cells (233). As an increase of cytosolic

Ca2+ could similarly induce apoptotic cell death in nucleated cells, activation of the volume

regulated cation channels could similarly participate in the triggering of apoptosis in

anucleated cells exposed to an osmotic shock (234, 235).

The present study reveals a completely novel effect of gum Arabic on the course of

malaria. Following infection with P. berghei the increase of parasitemia was significantly

less rapid in gum Arabic treated mice than in untreated mice (Fig. 3. 19). Moreover, gum

Arabic significantly increased the survival of P. berghei infected mice. Similar to what has

been shown earlier (210), untreated mice all die from infection with P. berghei at day 26.

In contrast, 70% of the gum Arabic treated mice survived up to day 26 following infection

(Fig. 3. 20). Thus, gum Arabic treatment favourably modifies the course of malaria.

77 

 

The present observations provide some hints to mechanisms underlying the beneficial

effect of gum Arabic treatment. It is well-known that gum Arabic is fermented by intestinal

bacteria, leading to the formation of various degradation products such as short-chain fatty

acids as butyrate which is important (236). In a recent study, serum butyrate concentrations

were increased after treatment with gum Arabic in healthy subjects (237). Moreover,

butyrate compounds have been shown to up regulate the formation of fetal haemoglobin

(238), which may in turn confer some protection against a severe course of malaria.

Specifically, fetal haemoglobin has been shown to delay the haemoglobin degradation and

thus to impede the intraerythrocyte growth of Plasmodium (239).

Gum Arabic could have affected parasitemia and host survival by increasing butyrate

levels, which is known to delay the intraerythrocyte growth of the parasite by its is effects

as inhibitor of histone deacetylase activity, or alternatively by increasing the erythrocyte

content of fetal haemoglobin, which is also known to delay the intraerythrocyte growth of

the parasite (239). Gum Arabic, butyrate and/or fetal hemoglobin may affect parasitemia

and host survival by accelerating the eryptosis of infected erythrocytes (240).

Phosphatidylserine-exposing erythrocytes are phagocytosed and thus rapidly cleared from

circulating blood (241).

One of the important results of this study is the prominent effects of amphotericin B and

butyrate/or gum Arabic in increasing erythrocytes cytosolic Ca2+. Increased cytosolic Ca2+

is crucial factor for stimulation of eryptosis (175). Eryptosis is triggered by redox-

sensitive, Ca2+ permeable, non-selective cation channels, activation of the Gardos K+

channel and cell shrinkage.

Infection with P. falciparum is expected to increase the cytosolic Ca2+ activity and thus to

induce eryptosis as a result of non selective cation channels stimulation. Most probably,

the parasite utilizes these Ca2+ permeable non-selective cation channels to adapt the ionic

composition of the host cytosol to its needs(175). However, most of the Ca2+ entering the

parasitized erythrocyte is either extruded across the host cell membrane or taken up by the

pathogen ‘‘buffering’’ so that cytosolic Ca2+ activity remains low. Buffering of the

cytosolic Ca2+ by the parasite probably prevents Gardos K+ channel activation and

substantial host cell shrinkage during early parasite development. Hence, in addition to its

78 

 

effect on parasite growth, the infection-induced Ca2+-permeable NSC conductance

mediates at least in part the reported transiently elevated free Ca2+ concentration in the host

cytosol Ca2+ (127, 242). Possibly in synergy with oxidative stress, the elevated cytosolic

Ca2+ in turn triggers breakdown of the phospholipid asymmetry of the host cell membrane

at mature parasite stages (243). Phosphatidylserine scrambling occurs especially under

conditions of high parasitemia, which is associated with increased oxidative stress and

ATP depletion (244).

Phosphatidylserine is present only in small amounts in the outer leaflet of fresh

erythrocytes (245). This is consistent with the low annexin-binding, i.e. in principle

phosphatidylserine exposure, observed in non-infected control co-cultured erythrocytes in

the present study. Phosphatidylserine exposure increases progressively on erythrocytes

when treated with the Ca2+ ionophore ionomycin (122). This is consistent with the present

measurements: erythrocytes treated with the Ca2+ ionophore ionomycin showed a high

degree of phosphatidylserine exposing erythrocytes. The phosphatidylserine exposure

observed in ionomycin-treated erythrocytes might be the result of the synergistic action of

the two known eryptosis signaling pathways: the increase in cytosolic Ca2+ per se activates

the scramblase and inhibits the translocase. Moreover, the Gardos effect activates the

sphingomyelinase that results in ceramide formation, which further stimulates the

scramblase.

Early the exposure of phosphatidylserine at the outer surface of the cell membrane is

presumably followed by binding to phosphatidylserine receptors on macrophages and

subsequent phagocytosis of the affected erythrocyte. The lysosomal degradation may

eventually eliminate the pathogen (246). Macrophages that phagocytose ring-stage infected

erythrocytes are less intoxicated by hemozoin as opposed to those phagocytose late-stage

infected erythrocytes. Plasmodium parasites detoxify the free heme of digested

hemoglobin as polymerized ß-hematin (the pigment hemozoin) in their food vacuoles.

Ingestion of late-trophozoite infected erythrocytes with a high amount of hemozoin by

phagocytes alters the function of macrophages, monocytes and dendritic cells (247).

Moreover, hemozoin-macrophage interactions impair migration of macrophages and

contribute to the cytokine-mediated pathology of malaria (248, 249).

79 

 

Erythrocytes with haemoglobinopathies or G6PD-deficiency are highly prone to enter

eryptosis (158) and pharmacological induction of phosphatidylserine exposure of ring

stage-infected erythrocytes reportedly accelerates their clearance. Thus, manoeuvres

accelerating eryptosis by administration of amphotericin B or gum Arabic may result in

premature clearance of the intraerythrocytic parasite. As a matter of fact, iron deficiency,

lead, chlorpromazine, inhibition of NO synthase by L-NAME, azathiopring, amiodarone,

and aurothiomalate (179, 180, 200, 250, 251), decrease parasitemia and partially enhance

the survival of Plasmodium berghei-infected mice eventually by accelerating erythrocyte

death. Thus, several conditions are known, which are associated with enhanced

susceptibility to eryptosis and at the same time with a milder course of malaria. Among

those, sickle cell trait is an example of a condition which is not associated with the

problem of developing resistance of the pathogen.

Conclusion

In conclusion amphotericin B and gum Arabic stimulate the erythrocytic machinery

responsible for the eryptosis following infection with Plasmodium. The acceleration of

eryptosis precedes the full intraerythrocytic maturation of the pathogen, thus prevents the

further lethal course of the disease and fosters host survival during malaria. The revelations

defend that the stimulation of eryptosis in infected erythrocytes is a host dependent

mechanism to combat against infection. The experimental results not only justify that the

stimulation of eryptosis in infected erythrocytes is not only a host dependent defence

mechanism but also a novel approach to prevent the chances of resistance in plasmodia.

80 

 

Recommendation

1. This method of identifying new host mediated antimalarial agents may be

combined with the regular antimalarial agents with host directed drug therapy and

increase the efficacy in eliminating the invaded and invading pathogen, thus control

the resurgence of once contained disease.

2. Amphotericin B and gum Arabic was found to blunts the parasitaemia and leads to

a favourable course of malaria. Further studies are needed to explore the additional

mechanisms behind these beneficial antimalarial activity.

81 

 

Refrences

1. Haldar K, Murphy SC, Milner DA, Taylor TE. Malaria: mechanisms of erythrocytic infection and pathological correlates of severe disease. Annu Rev Pathol. 2007;2:217-49.

2. Baumeister S, Winterberg M, Przyborski JM, Lingelbach K. The malaria parasite Plasmodium falciparum: cell biological peculiarities and nutritional consequences. Protoplasma. 2010 Apr;240(1-4):3-12.

3. Breman JG, Alilio MS, Mills A. Conquering the intolerable burden of malaria: what's new, what's needed: a summary. Am J Trop Med Hyg. 2004 Aug;71(2 Suppl):1-15.

4. Diap G, Amuasi J, Boakye I, Sevcsik AM, Pecoul B. Anti-malarial market and policy surveys in sub-Saharan Africa. Malar J. 2010;9 Suppl 1:S1.

5. Cox FE. History of the discovery of the malaria parasites and their vectors. Parasit Vectors. 2010 Feb 1;3(1):5.

6. Hay SI, Okiro EA, Gething PW, Patil AP, Tatem AJ, Guerra CA, et al. Estimating the Global Clinical Burden of Plasmodium falciparum Malaria in 2007. PLoS Med. 2010 Jun 15;7(6):e1000290.

7. World Health Organization, Malaria [database on the Internet]. WHO Media centre. 2007 [cited. Available from: http://www.who.int/mediacentre/factsheets/fs094/en/.

8. Sherman IW. The life of Plasmodium: An Overview. 2005. p. 3-11. 9. Ross R, Smyth J. On some peculiar pigmented cells found in two mosquitoes fed on malarial

blood. 1897. Indian J Malariol. 1997 Jun;34(2):47-55. 10. Grmek MD. Diseases In The Ancient Greek World. Baltimore; 1989. 11. Gething PW SD, Patil AP, Tatem AJ, Snow RW, Hay SI. Climate change and the global

malaria recession. Nature. 2010 May 20;465(7296):342-5. 12. Greene JA, Moormann AM, Vulule J, Bockarie MJ, Zimmerman PA, Kazura JW. Toll-like

receptor polymorphisms in malaria-endemic populations. Malar J. 2009;8:50. 13. Le Port A, Cot M, Etard JF, Gaye O, Migot-Nabias F, Garcia A. Relation between

Plasmodium falciparum asymptomatic infection and malaria attacks in a cohort of Senegalese children. Malar J. 2008;7:193.

14. Abel L, Cot M, Mulder L, Carnevale P, Feingold J. Segregation analysis detects a major gene controlling blood infection levels in human malaria. Am J Hum Genet. 1992 Jun;50(6):1308-17.

15. Aguas R, White LJ, Snow RW, Gomes MG. Prospects for malaria eradication in sub-Saharan Africa. PLoS One. 2008;3(3):e1767.

16. Giha HA, Elbashir MI, IE AE, TM AG, ElGhazali G. Bimodal transmission of cerebral malaria and severe malarial anemia and reciprocal co-existence of sexual and asexual parasitemia in an area of seasonal malaria transmission. Parasitol Res. 2008 Jun;103(1):81-5.

17. Peterson AT. Shifting suitability for malaria vectors across Africa with warming climates. BMC Infect Dis. 2009;9:59.

18. Roper C, Richardson W, Elhassan IM, Giha H, Hviid L, Satti GM, et al. Seasonal changes in the Plasmodium falciparum population in individuals and their relationship to clinical malaria: a longitudinal study in a Sudanese village. Parasitology. 1998 Jun;116 ( Pt 6):501-10.

82 

 

19. Giha HA, Rosthoj S, Dodoo D, Hviid L, Satti GM, Scheike T, et al. The epidemiology of febrile malaria episodes in an area of unstable and seasonal transmission. Trans R Soc Trop Med Hyg. 2000 Nov-Dec;94(6):645-51.

20. Robert V, Molez JF, Trape JF. Short report: gametocytes, chloroquine pressure, and the relative parasite survival advantage of resistant strains of falciparum malaria in west Africa. Am J Trop Med Hyg. 1996 Sep;55(3):350-1.

21. James MC, & Stephen, L.H. Medical Microbiology. The University of Texas Medical Branch at Galveston; 1996 [cited 2010 Jun 23]. Available from: http://www.ncbi.nlm.nih.gov/bookshelf/br.fcgi?book=mmed&part=A4419.

22. Snow RW, Korenromp EL, Gouws E. Pediatric mortality in Africa: plasmodium falciparum malaria as a cause or risk? Am J Trop Med Hyg. 2004 Aug;71(2 Suppl):16-24.

23. Azrak B, Gleissner C, Willershausen B, Jadamus-Stocker J, Callaway A. Accuracy of a chair-side test for predicting caries risk compared with established methods: A pilot study. Schweiz Monatsschr Zahnmed. 2010;120(5):409-14.

24. Tuteja R. Malaria - an overview. FEBS J. 2007 Sep;274(18):4670-9. 25. Mota MM, Rodriguez A. Migration through host cells by apicomplexan parasites. Microbes

Infect. 2001 Nov;3(13):1123-8. 26. Yamauchi LM, Coppi A, Snounou G, Sinnis P. Plasmodium sporozoites trickle out of the

injection site. Cell Microbiol. 2007 May;9(5):1215-22. 27. Sturm A, Amino R, van de Sand C, Regen T, Retzlaff S, Rennenberg A, et al. Manipulation of

host hepatocytes by the malaria parasite for delivery into liver sinusoids. Science. 2006 Sep 1;313(5791):1287-90.

28. Miller LH, Baruch DI, Marsh K, Doumbo OK. The pathogenic basis of malaria. Nature. 2002 Feb 7;415(6872):673-9.

29. Kuehn A, Pradel G. The coming-out of malaria gametocytes. J Biomed Biotechnol. 2010;2010:976827.

30. Eksi S, Czesny B, van Gemert GJ, Sauerwein RW, Eling W, Williamson KC. Malaria transmission-blocking antigen, Pfs230, mediates human red blood cell binding to exflagellating male parasites and oocyst production. Mol Microbiol. 2006 Aug;61(4):991-8.

31. Carter R, Mendis K. Measuring malaria. Am J Trop Med Hyg. 2006 Feb;74(2):187-8. 32. Gupta S, Snow RW, Donnelly CA, Marsh K, Newbold C. Immunity to non-cerebral severe

malaria is acquired after one or two infections. Nat Med. 1999 Mar;5(3):340-3. 33. Ohteki T. The dynamics of dendritic cell: mediated innate immune regulation. Allergol Int.

2007 Sep;56(3):209-14. 34. Ghosh K. Evolution and selection of human leukocyte antigen alleles by Plasmodium

falciparum infection. Hum Immunol. 2008 Dec;69(12):856-60. 35. Farnert A, Tengstam K, Palme IB, Bronner U, Lebbad M, Swedberg G, et al. Polyclonal

Plasmodium falciparum malaria in travelers and selection of antifolate mutations after proguanil prophylaxis. Am J Trop Med Hyg. 2002 May;66(5):487-91.

36. Doolan DL, Dobano C, Baird JK. Acquired immunity to malaria. Clin Microbiol Rev. 2009 Jan;22(1):13-36, Table of Contents.

37. Tangpukdee N, Duangdee C, Wilairatana P, Krudsood S. Malaria diagnosis: a brief review. Korean J Parasitol. 2009 Jun;47(2):93-102.

38. Patouillard E, Hanson KG, Goodman CA. Retail sector distribution chains for malaria treatment in the developing world: a review of the literature. Malar J. 2010;9:50.

39. White NJ. The role of anti-malarial drugs in eliminating malaria. Malar J. 2008;7 Suppl 1:S8.

83 

 

40. Giha HA. Prospects of intermittent preventive treatment of adults against malaria in areas of seasonal and unstable malaria transmission, and a possible role for chloroquine. Expert Opin Pharmacother. 2010 Apr;11(6):871-6.

41. Farooq U, Mahajan RC. Drug resistance in malaria. J Vector Borne Dis. 2004 Sep-Dec;41(3-4):45-53.

42. Winstanley P, Ward S, Snow R, Breckenridge A. Therapy of falciparum malaria in sub-saharan Africa: from molecule to policy. Clin Microbiol Rev. 2004 Jul;17(3):612-37, table of contents.

43. Skinner-Adams TS, Lawrie PM, Hawthorne PL, Gardiner DL, Trenholme KR. Comparison of Plasmodium falciparum transfection methods. Malar J. 2003 Jun 27;2:19.

44. Noiriel A, Naponelli V, Gregory JF, 3rd, Hanson AD. Pterin and folate salvage. Plants and Escherichia coli lack capacity to reduce oxidized pterins. Plant Physiol. 2007 Mar;143(3):1101-9.

45. The efficacy of antimalarial monotherapies, sulphadoxine-pyrimethamine and amodiaquine in East Africa: implications for sub-regional policy. Trop Med Int Health. 2003 Oct;8(10):860-7.

46. Whegang SY, Tahar R, Foumane VN, Soula G, Gwet H, Thalabard JC, et al. Efficacy of non-artemisinin- and artemisinin-based combination therapies for uncomplicated falciparum malaria in Cameroon. Malar J. 2010;9:56.

47. McIntosh HM, Olliaro P. Artemisinin derivatives for treating severe malaria. Cochrane Database Syst Rev. 2000(2):CD000527.

48. Nosten F, White NJ. Artemisinin-based combination treatment of falciparum malaria. Am J Trop Med Hyg. 2007 Dec;77(6 Suppl):181-92.

49. Hyde JE. Mechanisms of resistance of Plasmodium falciparum to antimalarial drugs. Microbes Infect. 2002 Feb;4(2):165-74.

50. Norbury CJ, Hickson ID. Cellular responses to DNA damage. Annu Rev Pharmacol Toxicol. 2001;41:367-401.

51. Hacker G. The morphology of apoptosis. Cell Tissue Res. 2000 Jul;301(1):5-17. 52. Panzarini E, Tenuzzo B, Dini L. Photodynamic therapy-induced apoptosis of HeLa cells. Ann

N Y Acad Sci. 2009 Aug;1171:617-26. 53. Savill J, Fadok V. Corpse clearance defines the meaning of cell death. Nature. 2000 Oct

12;407(6805):784-8. 54. Kurosaka K, Takahashi M, Watanabe N, Kobayashi Y. Silent cleanup of very early apoptotic

cells by macrophages. J Immunol. 2003 Nov 1;171(9):4672-9. 55. Forcet C, Ye X, Granger L, Corset V, Shin H, Bredesen DE, et al. The dependence receptor

DCC (deleted in colorectal cancer) defines an alternative mechanism for caspase activation. Proc Natl Acad Sci U S A. 2001 Mar 13;98(6):3416-21.

56. Castro-Obregon S, Del Rio G, Chen SF, Swanson RA, Frankowski H, Rao RV, et al. A ligand-receptor pair that triggers a non-apoptotic form of programmed cell death. Cell Death Differ. 2002 Aug;9(8):807-17.

57. Van Cruchten S, Van Den Broeck W. Morphological and biochemical aspects of apoptosis, oncosis and necrosis. Anat Histol Embryol. 2002 Aug;31(4):214-23.

58. Zeiss CJ. The apoptosis-necrosis continuum: insights from genetically altered mice. Vet Pathol. 2003 Sep;40(5):481-95.

59. Fiers W, Beyaert R, Declercq W, Vandenabeele P. More than one way to die: apoptosis, necrosis and reactive oxygen damage. Oncogene. 1999 Dec 16;18(54):7719-30.

84 

 

60. Patel VA, Lee DJ, Longacre-Antoni A, Feng L, Lieberthal W, Rauch J, et al. Apoptotic and necrotic cells as sentinels of local tissue stress and inflammation: response pathways initiated in nearby viable cells. Autoimmunity. 2009 May;42(4):317-21.

61. Igney FH, Krammer PH. Death and anti-death: tumour resistance to apoptosis. Nat Rev Cancer. 2002 Apr;2(4):277-88.

62. Martinvalet D, Zhu P, Lieberman J. Granzyme A induces caspase-independent mitochondrial damage, a required first step for apoptosis. Immunity. 2005 Mar;22(3):355-70.

63. Hengartner MO. The biochemistry of apoptosis. Nature. 2000 Oct 12;407(6805):770-6. 64. Yaginuma H, Sato N, Homma S, Oppenheim RW. Roles of caspases in the programmed cell

death of motoneurons in vivo. Arch Histol Cytol. 2001 Dec;64(5):461-74. 65. Rai NK, Tripathi K, Sharma D, Shukla VK. Apoptosis: a basic physiologic process in wound

healing. Int J Low Extrem Wounds. 2005 Sep;4(3):138-44. 66. Nakagawa T, Zhu H, Morishima N, Li E, Xu J, Yankner BA, et al. Caspase-12 mediates

endoplasmic-reticulum-specific apoptosis and cytotoxicity by amyloid-beta. Nature. 2000 Jan 6;403(6765):98-103.

67. Koenig U, Eckhart L, Tschachler E. Evidence that caspase-13 is not a human but a bovine gene. Biochem Biophys Res Commun. 2001 Aug 3;285(5):1150-4.

68. Kang SJ, Wang S, Kuida K, Yuan J. Distinct downstream pathways of caspase-11 in regulating apoptosis and cytokine maturation during septic shock response. Cell Death Differ. 2002 Oct;9(10):1115-25.

69. Elmore S. Apoptosis: a review of programmed cell death. Toxicol Pathol. 2007;35(4):495-516.

70. Borges HL, Linden R, Wang JY. DNA damage-induced cell death: lessons from the central nervous system. Cell Res. 2008 Jan;18(1):17-26.

71. Bratton DL, Fadok VA, Richter DA, Kailey JM, Guthrie LA, Henson PM. Appearance of phosphatidylserine on apoptotic cells requires calcium-mediated nonspecific flip-flop and is enhanced by loss of the aminophospholipid translocase. J Biol Chem. 1997 Oct 17;272(42):26159-65.

72. Gardai SJ, McPhillips KA, Frasch SC, Janssen WJ, Starefeldt A, Murphy-Ullrich JE, et al. Cell-surface calreticulin initiates clearance of viable or apoptotic cells through trans-activation of LRP on the phagocyte. Cell. 2005 Oct 21;123(2):321-34.

73. Arur S, Uche UE, Rezaul K, Fong M, Scranton V, Cowan AE, et al. Annexin I is an endogenous ligand that mediates apoptotic cell engulfment. Dev Cell. 2003 Apr;4(4):587-98.

74. Locksley RM, Killeen N, Lenardo MJ. The TNF and TNF receptor superfamilies: integrating mammalian biology. Cell. 2001 Feb 23;104(4):487-501.

75. Mathew SJ, Haubert D, Kronke M, Leptin M. Looking beyond death: a morphogenetic role for the TNF signalling pathway. J Cell Sci. 2009 Jun 15;122(Pt 12):1939-46.

76. Bentele M, Lavrik I, Ulrich M, Stosser S, Heermann DW, Kalthoff H, et al. Mathematical modeling reveals threshold mechanism in CD95-induced apoptosis. J Cell Biol. 2004 Sep 13;166(6):839-51.

77. Suliman A, Lam A, Datta R, Srivastava RK. Intracellular mechanisms of TRAIL: apoptosis through mitochondrial-dependent and -independent pathways. Oncogene. 2001 Apr 19;20(17):2122-33.

78. Rubio-Moscardo F, Blesa D, Mestre C, Siebert R, Balasas T, Benito A, et al. Characterization of 8p21.3 chromosomal deletions in B-cell lymphoma: TRAIL-R1 and TRAIL-R2 as candidate dosage-dependent tumor suppressor genes. Blood. 2005 Nov 1;106(9):3214-22.

85 

 

79. Hyer ML, Samuel T, Reed JC. The FLIP-side of Fas signaling. Clin Cancer Res. 2006 Oct 15;12(20 Pt 1):5929-31.

80. Debatin KM. Growth control of normal and malignant lymphocytes--cell death research from basic concepts to signal pathways and translation into the clinic. Klin Padiatr. 2009 Nov-Dec;221(6):332-8.

81. Kuhla A, Eipel C, Abshagen K, Siebert N, Menger MD, Vollmar B. Role of the perforin/granzyme cell death pathway in D-Gal/LPS-induced inflammatory liver injury. Am J Physiol Gastrointest Liver Physiol. 2009 May;296(5):G1069-76.

82. Pardo J, Bosque A, Brehm R, Wallich R, Naval J, Mullbacher A, et al. Apoptotic pathways are selectively activated by granzyme A and/or granzyme B in CTL-mediated target cell lysis. J Cell Biol. 2004 Nov 8;167(3):457-68.

83. Trisciuoglio L, Bianchi ME. Several nuclear events during apoptosis depend on caspase-3 activation but do not constitute a common pathway. PLoS One. 2009;4(7):e6234.

84. Cagigi A, Nilsson A, Levitsky V, Sabri F. Cytotoxic T-lymphocytes secrete soluble factors that induce caspase-mediated apoptosis in glioblastoma cell lines. J Neuroimmunol. 2010 May 16.

85. Fan Z, Beresford PJ, Oh DY, Zhang D, Lieberman J. Tumor suppressor NM23-H1 is a granzyme A-activated DNase during CTL-mediated apoptosis, and the nucleosome assembly protein SET is its inhibitor. Cell. 2003 Mar 7;112(5):659-72.

86. Lieberman J, Fan Z. Nuclear war: the granzyme A-bomb. Curr Opin Immunol. 2003 Oct;15(5):553-9.

87. Saelens X, Festjens N, Vande Walle L, van Gurp M, van Loo G, Vandenabeele P. Toxic proteins released from mitochondria in cell death. Oncogene. 2004 Apr 12;23(16):2861-74.

88. Garrido C, Galluzzi L, Brunet M, Puig PE, Didelot C, Kroemer G. Mechanisms of cytochrome c release from mitochondria. Cell Death Differ. 2006 Sep;13(9):1423-33.

89. Ledgerwood EC, Morison IM. Targeting the apoptosome for cancer therapy. Clin Cancer Res. 2009 Jan 15;15(2):420-4.

90. Hill MM, Adrain C, Duriez PJ, Creagh EM, Martin SJ. Analysis of the composition, assembly kinetics and activity of native Apaf-1 apoptosomes. EMBO J. 2004 May 19;23(10):2134-45.

91. Joza N, Susin SA, Daugas E, Stanford WL, Cho SK, Li CY, et al. Essential role of the mitochondrial apoptosis-inducing factor in programmed cell death. Nature. 2001 Mar 29;410(6828):549-54.

92. Cory S, Adams JM. The Bcl2 family: regulators of the cellular life-or-death switch. Nat Rev Cancer. 2002 Sep;2(9):647-56.

93. Chipuk JE, Green DR. Dissecting p53-dependent apoptosis. Cell Death Differ. 2006 Jun;13(6):994-1002.

94. Walsh JG, Cullen SP, Sheridan C, Luthi AU, Gerner C, Martin SJ. Executioner caspase-3 and caspase-7 are functionally distinct proteases. Proc Natl Acad Sci U S A. 2008 Sep 2;105(35):12815-9.

95. Conradt B, Xue D. Programmed cell death. WormBook. 2005:1-13. 96. Carter R, Sykes V, Lanning D. Scarless fetal mouse wound healing may initiate apoptosis

through caspase 7 and cleavage of PARP. J Surg Res. 2009 Sep;156(1):74-9. 97. Matsuda-Minehata F, Inoue N, Goto Y, Manabe N. The regulation of ovarian granulosa cell

death by pro- and anti-apoptotic molecules. J Reprod Dev. 2006 Dec;52(6):695-705. 98. Quaglino A, Salierno M, Pellegrotti J, Rubinstein N, Kordon EC. Mechanical strain induces

involution-associated events in mammary epithelial cells. BMC Cell Biol. 2009;10:55.

86 

 

99. Trinei M, Berniakovich I, Beltrami E, Migliaccio E, Fassina A, Pelicci P, et al. P66Shc signals to age. Aging (Albany NY). 2009 Jun;1(6):503-10.

100. Larsson NG. Somatic mitochondrial DNA mutations in mammalian aging. Annu Rev Biochem. 2010;79:683-706.

101. Kerr JF. History of the events leading to the formulation of the apoptosis concept. Toxicology. 2002 Dec 27;181-182:471-4.

102. Lopez-Huertas MR, Callejas S, Abia D, Mateos E, Dopazo A, Alcami J, et al. Modifications in host cell cytoskeleton structure and function mediated by intracellular HIV-1 Tat protein are greatly dependent on the second coding exon. Nucleic Acids Res. 2010 Jun;38(10):3287-307.

103. Nicholson DW. From bench to clinic with apoptosis-based therapeutic agents. Nature. 2000 Oct 12;407(6805):810-6.

104. Schimmer AD. Inhibitor of apoptosis proteins: translating basic knowledge into clinical practice. Cancer Res. 2004 Oct 15;64(20):7183-90.

105. Ward TH, Cummings J, Dean E, Greystoke A, Hou JM, Backen A, et al. Biomarkers of apoptosis. Br J Cancer. 2008 Sep 16;99(6):841-6.

106. Ward TH, Cummings J, Dean E, Greystoke A, Hou JM, Backen A, et al. Biomarkers of apoptosis. Br J Cancer. 2008 Aug 26.

107. Enomoto R, Sugahara C, Suzuki C, Nagase I, Takamura Y, Yoshikawa A, et al. Wogonin prevents glucocorticoid-induced thymocyte apoptosis without diminishing its anti-inflammatory action. J Pharmacol Sci. 2007 Aug;104(4):355-65.

108. Bossy-Wetzel E, Green DR. Detection of apoptosis by annexin V labeling. Methods Enzymol. 2000;322:15-8.

109. Zucker RM, Hunter ES, 3rd, Rogers JM. Confocal laser scanning microscopy of morphology and apoptosis in organogenesis-stage mouse embryos. Methods Mol Biol. 2000;135:191-202.

110. Holme AL, Yadav SK, Pervaiz S. Automated laser scanning cytometry: a powerful tool for multi-parameter analysis of drug-induced apoptosis. Cytometry A. 2007 Feb;71(2):80-6.

111. Bratosin D, Estaquier J, Petit F, Arnoult D, Quatannens B, Tissier JP, et al. Programmed cell death in mature erythrocytes: a model for investigating death effector pathways operating in the absence of mitochondria. Cell Death Differ. 2001 Dec;8(12):1143-56.

112. Lang F, Foller M, Lang KS, Lang PA, Ritter M, Gulbins E, et al. Ion channels in cell proliferation and apoptotic cell death. J Membr Biol. 2005 Jun;205(3):147-57.

113. Khandelwal S, Saxena RK. A role of phosphatidylserine externalization in clearance of erythrocytes exposed to stress but not in eliminating aging populations of erythrocyte in mice. Exp Gerontol. 2008 Aug;43(8):764-70.

114. Lang KS, Lang PA, Bauer C, Duranton C, Wieder T, Huber SM, et al. Mechanisms of suicidal erythrocyte death. Cell Physiol Biochem. 2005;15(5):195-202.

115. Lang F, Lang KS, Wieder T, Myssina S, Birka C, Lang PA, et al. Cation channels, cell volume and the death of an erythrocyte. Pflugers Arch. 2003 Nov;447(2):121-5.

116. Lang KS, Duranton C, Poehlmann H, Myssina S, Bauer C, Lang F, et al. Cation channels trigger apoptotic death of erythrocytes. Cell Death Differ. 2003 Feb;10(2):249-56.

117. Lang KS, Weigert C, Braedel S, Fillon S, Palmada M, Schleicher E, et al. Inhibition of interferon-gamma expression by osmotic shrinkage of peripheral blood lymphocytes. Am J Physiol Cell Physiol. 2003 Jan;284(1):C200-8.

118. Peng Z, Asaro RJ, Zhu Q. Multiscale simulation of erythrocyte membranes. Phys Rev E Stat Nonlin Soft Matter Phys. 2010 Mar;81(3 Pt 1):031904.

87 

 

119. Walensky LD, Mohandas N, Lux SE. In: Blood, Principles and Practice of Hematology. 2 ed. Handin RI LS, Stossel TP, editor. Philadelphia, PA: Lippincott Williams & Wilkins; 2003.

120. Gummadi SN, Kumar KS. The mystery of phospholipid flip-flop in biogenic membranes. Cell Mol Biol Lett. 2005;10(1):101-21.

121. Daleke DL. Regulation of phospholipid asymmetry in the erythrocyte membrane. Curr Opin Hematol. 2008 May;15(3):191-5.

122. Lang F, Gulbins E, Lang PA, Zappulla D, Foller M. Ceramide in suicidal death of erythrocytes. Cell Physiol Biochem. 2010;26(1):21-8.

123. Mohandas N, Gallagher PG. Red cell membrane: past, present, and future. Blood. 2008 Nov 15;112(10):3939-48.

124. Yasin Z, Witting S, Palascak MB, Joiner CH, Rucknagel DL, Franco RS. Phosphatidylserine externalization in sickle red blood cells: associations with cell age, density, and hemoglobin F. Blood. 2003 Jul 1;102(1):365-70.

125. Lew VL, Bookchin RM. Ion transport pathology in the mechanism of sickle cell dehydration. Physiol Rev. 2005 Jan;85(1):179-200.

126. Lew VL, Tiffert T, Etzion Z, Perdomo D, Daw N, Macdonald L, et al. Distribution of dehydration rates generated by maximal Gardos-channel activation in normal and sickle red blood cells. Blood. 2005 Jan 1;105(1):361-7.

127. Mauritz JM, Esposito A, Ginsburg H, Kaminski CF, Tiffert T, Lew VL. The homeostasis of Plasmodium falciparum-infected red blood cells. PLoS Comput Biol. 2009 Apr;5(4):e1000339.

128. Huber SM, Duranton C, Lang F. Patch-clamp analysis of the "new permeability pathways" in malaria-infected erythrocytes. Int Rev Cytol. 2005;246:59-134.

129. Bennekou P, Kristensen BI, Christophersen P. The human red cell voltage-regulated cation channel. The interplay with the chloride conductance, the Ca(2+)-activated K(+) channel and the Ca(2+) pump. J Membr Biol. 2003 Sep 1;195(1):1-8.

130. Rodighiero S, De Simoni A, Formenti A. The voltage-dependent nonselective cation current in human red blood cells studied by means of whole-cell and nystatin-perforated patch-clamp techniques. Biochim Biophys Acta. 2004 Jan 28;1660(1-2):164-70.

131. Barksmann TL, Kristensen BI, Christophersen P, Bennekou P. Pharmacology of the human red cell voltage-dependent cation channel; Part I. Activation by clotrimazole and analogues. Blood Cells Mol Dis. 2004 May-Jun;32(3):384-8.

132. Bruce LJ. Red cell membrane transport abnormalities. Curr Opin Hematol. 2008 May;15(3):184-90.

133. Bennekou P, Barksmann TL, Jensen LR, Kristensen BI, Christophersen P. Voltage activation and hysteresis of the non-selective voltage-dependent channel in the intact human red cell. Bioelectrochemistry. 2004 May;62(2):181-5.

134. Duranton C, Huber SM, Lang F. Oxidation induces a Cl(-)-dependent cation conductance in human red blood cells. J Physiol. 2002 Mar 15;539(Pt 3):847-55.

135. Huber SM, Uhlemann AC, Gamper NL, Duranton C, Kremsner PG, Lang F. Plasmodium falciparum activates endogenous Cl(-) channels of human erythrocytes by membrane oxidation. EMBO J. 2002 Jan 15;21(1-2):22-30.

136. Gardos G. The function of calcium in the potassium permeability of human erythrocytes. Biochim Biophys Acta. 1958 Dec;30(3):653-4.

137. Hoffman JF, Joiner W, Nehrke K, Potapova O, Foye K, Wickrema A. The hSK4 (KCNN4) isoform is the Ca2+-activated K+ channel (Gardos channel) in human red blood cells. Proc Natl Acad Sci U S A. 2003 Jun 10;100(12):7366-71.

88 

 

138. Dyrda A, Cytlak U, Ciuraszkiewicz A, Lipinska A, Cueff A, Bouyer G, et al. Local membrane deformations activate Ca2+-dependent K+ and anionic currents in intact human red blood cells. PLoS One. 2010;5(2):e9447.

139. Del Carlo B, Pellegrini M, Pellegrino M. Calmodulin antagonists do not inhibit IK(Ca) channels of human erythrocytes. Biochim Biophys Acta. 2002 Feb 1;1558(2):133-41.

140. Cheng K, Haspel HC, Vallano ML, Osotimehin B, Sonenberg M. Measurement of membrane potentials (psi) of erythrocytes and white adipocytes by the accumulation of triphenylmethylphosphonium cation. J Membr Biol. 1980 Oct 31;56(3):191-201.

141. Lang PA, Kempe DS, Myssina S, Tanneur V, Birka C, Laufer S, et al. PGE(2) in the regulation of programmed erythrocyte death. Cell Death Differ. 2005 May;12(5):415-28.

142. Lang F, Lang KS, Lang PA, Huber SM, Wieder T. Osmotic shock-induced suicidal death of erythrocytes. Acta Physiol (Oxf). 2006 May-Jun;187(1-2):191-8.

143. Myssina S, Lang PA, Kempe DS, Kaiser S, Huber SM, Wieder T, et al. Cl- channel blockers NPPB and niflumic acid blunt Ca(2+)-induced erythrocyte 'apoptosis'. Cell Physiol Biochem. 2004;14(4-6):241-8.

144. Lang PA, Kaiser S, Myssina S, Wieder T, Lang F, Huber SM. Role of Ca2+-activated K+ channels in human erythrocyte apoptosis. Am J Physiol Cell Physiol. 2003 Dec;285(6):C1553-60.

145. De Franceschi L, Rivera A, Fleming MD, Honczarenko M, Peters LL, Gascard P, et al. Evidence for a protective role of the Gardos channel against hemolysis in murine spherocytosis. Blood. 2005 Aug 15;106(4):1454-9.

146. Zhou Q, Zhao J, Wiedmer T, Sims PJ. Normal hemostasis but defective hematopoietic response to growth factors in mice deficient in phospholipid scramblase 1. Blood. 2002 Jun 1;99(11):4030-8.

147. Bookchin RM, Ortiz OE, Lew VL. Activation of calcium-dependent potassium channels in deoxygenated sickled red cells. Prog Clin Biol Res. 1987;240:193-200.

148. Lang PA, Warskulat U, Heller-Stilb B, Huang DY, Grenz A, Myssina S, et al. Blunted apoptosis of erythrocytes from taurine transporter deficient mice. Cell Physiol Biochem. 2003;13(6):337-46.

149. Shimizu T, Maeno E, Okada Y. Prerequisite role of persistent cell shrinkage in apoptosis of human epithelial cells. Sheng Li Xue Bao. 2007 Aug 25;59(4):512-6.

150. Klarl BA, Lang PA, Kempe DS, Niemoeller OM, Akel A, Sobiesiak M, et al. Protein kinase C mediates erythrocyte "programmed cell death" following glucose depletion. Am J Physiol Cell Physiol. 2006 Jan;290(1):C244-53.

151. Sopjani M, Foller M, Haendeler J, Gotz F, Lang F. Silver ion-induced suicidal erythrocyte death. J Appl Toxicol. 2009 Aug;29(6):531-6.

152. Bourikas D, Kaloyianni M, Bougoulia M, Zolota Z, Koliakos G. Modulation of the Na(+)-H(+) antiport activity by adrenaline on erythrocytes from normal and obese individuals. Mol Cell Endocrinol. 2003 Jul 31;205(1-2):141-50.

153. Barvitenko NN, Adragna NC, Weber RE. Erythrocyte signal transduction pathways, their oxygenation dependence and functional significance. Cell Physiol Biochem. 2005;15(1-4):1-18.

154. Kanias T, Acker JP. Biopreservation of red blood cells--the struggle with hemoglobin oxidation. FEBS J. 2010 Jan;277(2):343-56.

155. Matarrese P, Straface E, Pietraforte D, Gambardella L, Vona R, Maccaglia A, et al. Peroxynitrite induces senescence and apoptosis of red blood cells through the activation of aspartyl and cysteinyl proteases. FASEB J. 2005 Mar;19(3):416-8.

89 

 

156. Mandal D, Baudin-Creuza V, Bhattacharyya A, Pathak S, Delaunay J, Kundu M, et al. Caspase 3-mediated proteolysis of the N-terminal cytoplasmic domain of the human erythroid anion exchanger 1 (band 3). J Biol Chem. 2003 Dec 26;278(52):52551-8.

157. Foller M, Huber SM, Lang F. Erythrocyte programmed cell death. IUBMB Life. 2008 Oct;60(10):661-8.

158. Bilmen S, Aksu TA, Gumuslu S, Korgun DK, Canatan D. Antioxidant capacity of G-6-PD-deficient erythrocytes. Clin Chim Acta. 2001 Jan;303(1-2):83-6.

159. Tanneur V, Duranton C, Brand VB, Sandu CD, Akkaya C, Kasinathan RS, et al. Purinoceptors are involved in the induction of an osmolyte permeability in malaria-infected and oxidized human erythrocytes. FASEB J. 2006 Jan;20(1):133-5.

160. Gokkusu C, Mostafazadeh T. Changes of oxidative stress in various tissues by long-term administration of vitamin E in hypercholesterolemic rats. Clin Chim Acta. 2003 Feb;328(1-2):155-61.

161. Kobayashi S, Moriya H, Aso K, Ohtake T. Vitamin E-bonded hemodialyzer improves atherosclerosis associated with a rheological improvement of circulating red blood cells. Kidney Int. 2003 May;63(5):1881-7.

162. Hess JR. Red cell storage. J Proteomics. 2010 Jan 3;73(3):368-73. 163. de Franceschi L, Turrini F, Honczarenko M, Ayi K, Rivera A, Fleming MD, et al. In vivo

reduction of erythrocyte oxidant stress in a murine model of beta-thalassemia. Haematologica. 2004 Nov;89(11):1287-98.

164. Lang PA, Kempe DS, Tanneur V, Eisele K, Klarl BA, Myssina S, et al. Stimulation of erythrocyte ceramide formation by platelet-activating factor. J Cell Sci. 2005 Mar 15;118(Pt 6):1233-43.

165. Lang KS, Myssina S, Brand V, Sandu C, Lang PA, Berchtold S, et al. Involvement of ceramide in hyperosmotic shock-induced death of erythrocytes. Cell Death Differ. 2004 Feb;11(2):231-43.

166. Rivera A. Reduced sickle erythrocyte dehydration in vivo by endothelin-1 receptor antagonists. Am J Physiol Cell Physiol. 2007 Sep;293(3):C960-6.

167. Lang F, Lang KS, Lang PA, Huber SM, Wieder T. Mechanisms and significance of eryptosis. Antioxid Redox Signal. 2006 Jul-Aug;8(7-8):1183-92.

168. Lang KS, Roll B, Myssina S, Schittenhelm M, Scheel-Walter HG, Kanz L, et al. Enhanced erythrocyte apoptosis in sickle cell anemia, thalassemia and glucose-6-phosphate dehydrogenase deficiency. Cell Physiol Biochem. 2002;12(5-6):365-72.

169. Kempe DS, Lang PA, Duranton C, Akel A, Lang KS, Huber SM, et al. Enhanced programmed cell death of iron-deficient erythrocytes. FASEB J. 2006 Feb;20(2):368-70.

170. Lang F, Busch GL, Ritter M, Volkl H, Waldegger S, Gulbins E, et al. Functional significance of cell volume regulatory mechanisms. Physiol Rev. 1998 Jan;78(1):247-306.

171. Gulbins E, Lang F. Pathogens, host-cell invasion and disease. Am Scientist. [Web Article]. 2001;89(5):406-41.

172. Martin RE, Ginsburg H, Kirk K. Membrane transport proteins of the malaria parasite. Mol Microbiol. 2009 Nov;74(3):519-28.

173. Lew VL, Tiffert T, Ginsburg H. Excess hemoglobin digestion and the osmotic stability of Plasmodium falciparum-infected red blood cells. Blood. 2003 May 15;101(10):4189-94.

174. Vaid A, Thomas DC, Sharma P. Role of Ca2+/calmodulin-PfPKB signaling pathway in erythrocyte invasion by Plasmodium falciparum. J Biol Chem. 2008 Feb 29;283(9):5589-97.

175. Lang F, Lang PA, Lang KS, Brand V, Tanneur V, Duranton C, et al. Channel-induced apoptosis of infected host cells-the case of malaria. Pflugers Arch. 2004 Jun;448(3):319-24.

90 

 

176. Gazarini ML, Thomas AP, Pozzan T, Garcia CR. Calcium signaling in a low calcium environment: how the intracellular malaria parasite solves the problem. J Cell Biol. 2003 Apr 14;161(1):103-10.

177. Sherman IW, Eda S, Winograd E. Cytoadherence and sequestration in Plasmodium falciparum: defining the ties that bind. Microbes Infect. 2003 Aug;5(10):897-909.

178. Foller M, Bobbala D, Koka S, Huber SM, Gulbins E, Lang F. Suicide for survival--death of infected erythrocytes as a host mechanism to survive malaria. Cell Physiol Biochem. 2009;24(3-4):133-40.

179. Koka S, Foller M, Lamprecht G, Boini KM, Lang C, Huber SM, et al. Iron deficiency influences the course of malaria in Plasmodium berghei infected mice. Biochem Biophys Res Commun. 2007 Jun 8;357(3):608-14.

180. Koka S, Lang C, Niemoeller OM, Boini KM, Nicolay JP, Huber SM, et al. Influence of NO synthase inhibitor L-NAME on parasitemia and survival of Plasmodium berghei infected mice. Cell Physiol Biochem. 2008;21(5-6):481-8.

181. Sijwali PS, Rosenthal PJ. Functional evaluation of Plasmodium export signals in Plasmodium berghei suggests multiple modes of protein export. PLoS One. 2010;5(4):e10227.

182. Garcia CR, de Azevedo MF, Wunderlich G, Budu A, Young JA, Bannister L. Plasmodium in the postgenomic era: new insights into the molecular cell biology of malaria parasites. Int Rev Cell Mol Biol. 2008;266:85-156.

183. Coppens I, Sullivan DJ, Prigge ST. An update on the rapid advances in malaria parasite cell biology. Trends Parasitol. 2010 Jun;26(6):305-10.

184. Young JA, Johnson JR, Benner C, Yan SF, Chen K, Le Roch KG, et al. In silico discovery of transcription regulatory elements in Plasmodium falciparum. BMC Genomics. 2008;9:70.

185. Younes H, Garleb K, Behr S, Remesy C, Demigne C. Fermentable fibers or oligosaccharides reduce urinary nitrogen excretion by increasing urea disposal in the rat cecum. J Nutr. 1995 Apr;125(4):1010-6.

186. Tiss A, Carriere F, Verger R. Effects of gum arabic on lipase interfacial binding and activity. Anal Biochem. 2001 Jul 1;294(1):36-43.

187. Fauconnier ML, Blecker C, Groyne J, Razafindralambo H, Vanzeveren E, Marlier M, et al. Characterization of two Acacia gums and their fractions using a langmuir film balance. J Agric Food Chem. 2000 Jul;48(7):2709-12.

188. Islam AM, Phillips GO, Sljivo A, Snowden MJ, Williams PA. A review of recent developments on the regulatory, structural and functional aspects of gum arabic. Food Hydrocolloids; 1997 [updated 1997; cited]; 493–505]. Available from: http://www.fstadirect.com/GetRecord.aspx?AN=1998-02-Tb0091.

189. Phillips GO. Acacia gum (Gum Arabic): a nutritional fibre; metabolism and calorific value. Food Addit Contam. 1998 Apr;15(3):251-64.

190. Kishimoto A, Ushida K, Phillips GO, Ogasawara T, Sasaki Y. Identification of intestinal bacteria responsible for fermentation of gum arabic in pig model. Curr Microbiol. 2006 Sep;53(3):173-7.

191. Verbeken D, Dierckx S, Dewettinck K. Exudate gums: occurrence, production, and applications. Appl Microbiol Biotechnol. 2003 Nov;63(1):10-21.

192. Adedoyin A, Bernardo JF, Swenson CE, Bolsack LE, Horwith G, DeWit S, et al. Pharmacokinetic profile of ABELCET (amphotericin B lipid complex injection): combined experience from phase I and phase II studies. Antimicrob Agents Chemother. 1997 Oct;41(10):2201-8.

91 

 

193. Chandrasekar PH, Ito JI. Amphotericin B lipid complex in the management of invasive aspergillosis in immunocompromised patients. Clin Infect Dis. 2005 May 1;40 Suppl 6:S392-400.

194. Boucherit Z, Seksek O, Bolard J. Dormancy of Candida albicans cells in the presence of the polyene antibiotic amphotericin B: simple demonstration by flow cytometry. Med Mycol. 2007 Sep;45(6):525-33.

195. Perfect JR, Marr KA, Walsh TJ, Greenberg RN, DuPont B, de la Torre-Cisneros J, et al. Voriconazole treatment for less-common, emerging, or refractory fungal infections. Clin Infect Dis. 2003 May 1;36(9):1122-31.

196. Nolin TD, Himmelfarb J. Mechanisms of drug-induced nephrotoxicity. Handb Exp Pharmacol. 2010(196):111-30.

197. Binh VQ, Luty AJ, Kremsner PG. Differential effects of human serum and cells on the growth of Plasmodium falciparum adapted to serum-free in vitro culture conditions. Am J Trop Med Hyg. 1997 Nov;57(5):594-600.

198. Beaudoin RL, Strome CP, Tubergen TA, Mitchell F. Plasmodium berghei berghei: irradiated sporozoites of the ANKA strain as immunizing antigens in mice. Exp Parasitol. 1976 Jun;39(3):438-43.

199. Desruisseaux MS, Gulinello M, Smith DN, Lee SC, Tsuji M, Weiss LM, et al. Cognitive dysfunction in mice infected with Plasmodium berghei strain ANKA. J Infect Dis. 2008 Jun 1;197(11):1621-7.

200. Bobbala D, Alesutan I, Foller M, Tschan S, Huber SM, Lang F. Protective effect of amiodarone in malaria. Acta Trop. 2010 May 27.

201. Jensen JB, Trager W. Plasmodium falciparum in culture: establishment of additional strains. Am J Trop Med Hyg. 1978 Jul;27(4):743-6.

202. Trager W, Jensen JB. Human malaria parasites in continuous culture. Science. 1976 Aug 20;193(4254):673-5.

203. Ljungström I, Perlmann H, Schlichtherle M, Scherf A, Wahlgren M. Methods in malaria research. 4 ed. Manassas (Virginia): Malaria Research and Reference Reagent Resource Center; 2004. p. 248.

204. Ameri M. Laboratory diagnosis of malaria in nonhuman primates. Vet Clin Pathol. 2010 Mar;39(1):5-19.

205. Kucherenko YV, Bhavsar SK, Grischenko VI, Fischer UR, Huber SM, Lang F. Increased cation conductance in human erythrocytes artificially aged by glycation. J Membr Biol. 2010 Jun;235(3):177-89.

206. Andrews DA, Yang L, Low PS. Phorbol ester stimulates a protein kinase C-mediated agatoxin-TK-sensitive calcium permeability pathway in human red blood cells. Blood. 2002 Nov 1;100(9):3392-9.

207. Mahmud H, Mauro D, Qadri SM, Foller M, Lang F. Triggering of suicidal erythrocyte death by amphotericin B. Cell Physiol Biochem. 2009;24(3-4):263-70.

208. Bhavsar SK, Eberhard M, Bobbala D, Lang F. Monensin induced suicidal erythrocyte death. Cell Physiol Biochem. 2010;25(6):745-52.

209. Brand V, Koka S, Lang C, Jendrossek V, Huber SM, Gulbins E, et al. Influence of amitriptyline on eryptosis, parasitemia and survival of Plasmodium berghei-infected mice. Cell Physiol Biochem. 2008;22(5-6):405-12.

210. Prudencio M, Derbyshire ET, Marques CA, Krishna S, Mota MM, Staines HM. Plasmodium berghei-infection induces volume-regulated anion channel-like activity in human hepatoma cells. Cell Microbiol. 2009 Oct;11(10):1492-501.

92 

 

211. Duranton C, Huber SM, Tanneur V, Brand VB, Akkaya C, Shumilina EV, et al. Organic osmolyte permeabilities of the malaria-induced anion conductances in human erythrocytes. J Gen Physiol. 2004 Apr;123(4):417-26.

212. Wiehart UI, Rautenbach M, Hoppe HC. Selective lysis of erythrocytes infected with the trophozoite stage of Plasmodium falciparum by polyene macrolide antibiotics. Biochem Pharmacol. 2006 Mar 14;71(6):779-90.

213. Nath CE, McLachlan AJ, Shaw PJ, Coakley JC, Earl JW. Amphotericin B dose optimization in children with malignant diseases. Chemotherapy. 2007;53(2):142-7.

214. Wiederhold NP, Tam VH, Chi J, Prince RA, Kontoyiannis DP, Lewis RE. Pharmacodynamic activity of amphotericin B deoxycholate is associated with peak plasma concentrations in a neutropenic murine model of invasive pulmonary aspergillosis. Antimicrob Agents Chemother. 2006 Feb;50(2):469-73.

215. Jung SH, Lim DH, Lee JE, Jeong KS, Seong H, Shin BC. Amphotericin B-entrapping lipid nanoparticles and their in vitro and in vivo characteristics. Eur J Pharm Sci. 2009 Jun 28;37(3-4):313-20.

216. Bosman GJ, Lasonder E, Groenen-Dopp YA, Willekens FL, Werre JM, Novotny VM. Comparative proteomics of erythrocyte aging in vivo and in vitro. J Proteomics. 2010 Jan 3;73(3):396-402.

217. Birka C, Lang PA, Kempe DS, Hoefling L, Tanneur V, Duranton C, et al. Enhanced susceptibility to erythrocyte "apoptosis" following phosphate depletion. Pflugers Arch. 2004 Aug;448(5):471-7.

218. Myssina S, Huber SM, Birka C, Lang PA, Lang KS, Friedrich B, et al. Inhibition of erythrocyte cation channels by erythropoietin. J Am Soc Nephrol. 2003 Nov;14(11):2750-7.

219. Hagerstrand H, Holmstrom TH, Bobrowska-Hagerstrand M, Eriksson JE, Isomaa B. Amphiphile-induced phosphatidylserine exposure in human erythrocytes. Mol Membr Biol. 1998 Apr-Jun;15(2):89-95.

220. Lang F, Gulbins E, Szabo I, Lepple-Wienhues A, Huber SM, Duranton C, et al. Cell volume and the regulation of apoptotic cell death. J Mol Recognit. 2004 Sep-Oct;17(5):473-80.

221. Resat H, Baginski M. Ion passage pathways and thermodynamics of the amphotericin B membrane channel. Eur Biophys J. 2002 Jul;31(4):294-305.

222. Yano T, Itoh Y, Kawamura E, Maeda A, Egashira N, Nishida M, et al. Amphotericin B-induced renal tubular cell injury is mediated by Na+ Influx through ion-permeable pores and subsequent activation of mitogen-activated protein kinases and elevation of intracellular Ca2+ concentration. Antimicrob Agents Chemother. 2009 Apr;53(4):1420-6.

223. Matsuo K, Hotokezaka H, Ohara N, Fujimura Y, Yoshimura A, Okada Y, et al. Analysis of amphotericin B-induced cell signaling with chemical inhibitors of signaling molecules. Microbiol Immunol. 2006;50(4):337-47.

224. Delic D, Warskulat U, Borsch E, Al-Qahtani S, Al-Quraishi S, Haussinger D, et al. Loss of ability to self-heal malaria upon taurine transporter deletion. Infect Immun. 2010 Apr;78(4):1642-9.

225. Davie JR. Inhibition of histone deacetylase activity by butyrate. J Nutr. 2003 Jul;133(7 Suppl):2485S-93S.

226. Andrews KT, Walduck A, Kelso MJ, Fairlie DP, Saul A, Parsons PG. Anti-malarial effect of histone deacetylation inhibitors and mammalian tumour cytodifferentiating agents. Int J Parasitol. 2000 May;30(6):761-8.

93 

 

227. Evert BO, Araujo J, Vieira-Saecker AM, de Vos RA, Harendza S, Klockgether T, et al. Ataxin-3 represses transcription via chromatin binding, interaction with histone deacetylase 3, and histone deacetylation. J Neurosci. 2006 Nov 1;26(44):11474-86.

228. Sesti-Costa R, Baccan GC, Chedraoui-Silva S, Mantovani B. Effects of acute cold stress on phagocytosis of apoptotic cells: the role of corticosterone. Neuroimmunomodulation. 2010;17(2):79-87.

229. Wong K, Valdez PA, Tan C, Yeh S, Hongo JA, Ouyang W. Phosphatidylserine receptor Tim-4 is essential for the maintenance of the homeostatic state of resident peritoneal macrophages. Proc Natl Acad Sci U S A. 2010 May 11;107(19):8712-7.

230. Koka S, Huber SM, Boini KM, Lang C, Foller M, Lang F. Lead decreases parasitemia and enhances survival of Plasmodium berghei-infected mice. Biochem Biophys Res Commun. 2007 Nov 23;363(3):484-9.

231. Duranton C, Huber S, Tanneur V, Lang K, Brand V, Sandu C, et al. Electrophysiological properties of the Plasmodium Falciparum-induced cation conductance of human erythrocytes. Cell Physiol Biochem. 2003;13(4):189-98.

232. Tyurina YY, Tyurin VA, Zhao Q, Djukic M, Quinn PJ, Pitt BR, et al. Oxidation of phosphatidylserine: a mechanism for plasma membrane phospholipid scrambling during apoptosis? Biochem Biophys Res Commun. 2004 Nov 19;324(3):1059-64.

233. Numata T, Sato K, Okada Y, Wehner F. Hypertonicity-induced cation channels rescue cells from staurosporine-elicited apoptosis. Apoptosis. 2008 Jul;13(7):895-903.

234. Jessica Chen M, Sepramaniam S, Armugam A, Shyan Choy M, Manikandan J, Melendez AJ, et al. Water and ion channels: crucial in the initiation and progression of apoptosis in central nervous system? Curr Neuropharmacol. 2008 Jun;6(2):102-16.

235. Zhang Y, Bhavnani BR. Glutamate-induced apoptosis in primary cortical neurons is inhibited by equine estrogens via down-regulation of caspase-3 and prevention of mitochondrial cytochrome c release. BMC Neurosci. 2005;6:13.

236. Bliss DZ. Dietary fiber in conservative management of chronic renal failure. Pediatr Nephrol. 2004 Oct;19(10):1069-70.

237. Matsumoto N, Riley S, Fraser D, Al-Assaf S, Ishimura E, Wolever T, et al. Butyrate modulates TGF-beta1 generation and function: potential renal benefit for Acacia(sen) SUPERGUM (gum arabic)? Kidney Int. 2006 Jan;69(2):257-65.

238. Mankidy R, Faller DV, Mabaera R, Lowrey CH, Boosalis MS, White GL, et al. Short-chain fatty acids induce gamma-globin gene expression by displacement of a HDAC3-NCoR repressor complex. Blood. 2006 Nov 1;108(9):3179-86.

239. Larru B, Molyneux E, Ter Kuile FO, Taylor T, Molyneux M, Terlouw DJ. Malaria in infants below six months of age: retrospective surveillance of hospital admission records in Blantyre, Malawi. Malar J. 2009;8(1):310.

240. Geiger C, Foller M, Herrlinger KR, Lang F. Azathioprine-induced suicidal erythrocyte death. Inflamm Bowel Dis. 2008 Aug;14(8):1027-32.

241. Hirano K, Miki Y, Hirai Y, Sato R, Itoh T, Hayashi A, et al. A multifunctional shuttling protein nucleolin is a macrophage receptor for apoptotic cells. J Biol Chem. 2005 Nov 25;280(47):39284-93.

242. Rohrbach P, Friedrich O, Hentschel J, Plattner H, Fink RH, Lanzer M. Quantitative calcium measurements in subcellular compartments of Plasmodium falciparum-infected erythrocytes. J Biol Chem. 2005 Jul 29;280(30):27960-9.

94 

 

243. Lang KS, Myssina S, Tanneur V, Wieder T, Huber SM, Lang F, et al. Inhibition of erythrocyte cation channels and apoptosis by ethylisopropylamiloride. Naunyn Schmiedebergs Arch Pharmacol. 2003 Apr;367(4):391-6.

244. Bosman GJ, Werre JM, Willekens FL, Novotny VM. Erythrocyte ageing in vivo and in vitro: structural aspects and implications for transfusion. Transfus Med. 2008 Dec;18(6):335-47.

245. Smith C, Gibson DF, Tait JF. Transmembrane voltage regulates binding of annexin V and lactadherin to cells with exposed phosphatidylserine. BMC Biochem. 2009;10:5.

246. Barascuk N, Skjot-Arkil H, Register TC, Larsen L, Byrjalsen I, Christiansen C, et al. Human macrophage foam cells degrade atherosclerotic plaques through cathepsin K mediated processes. BMC Cardiovasc Disord. 2010;10:19.

247. Skorokhod OA, Schwarzer E, Ceretto M, Arese P. Malarial pigment haemozoin, IFN-gamma, TNF-alpha, IL-1beta and LPS do not stimulate expression of inducible nitric oxide synthase and production of nitric oxide in immuno-purified human monocytes. Malar J. 2007;6:73.

248. McDevitt MA, Xie J, Shanmugasundaram G, Griffith J, Liu A, McDonald C, et al. A critical role for the host mediator macrophage migration inhibitory factor in the pathogenesis of malarial anemia. J Exp Med. 2006 May 15;203(5):1185-96.

249. Awandare GA, Ouma Y, Ouma C, Were T, Otieno R, Keller CC, et al. Role of monocyte-acquired hemozoin in suppression of macrophage migration inhibitory factor in children with severe malarial anemia. Infect Immun. 2007 Jan;75(1):201-10.

250. Bobbala D, Koka S, Geiger C, Foller M, Huber SM, Lang F. Azathioprine favourably influences the course of malaria. Malar J. 2009;8:102.

251. Alesutan I, Bobbala D, Qadri SM, Estremera A, Foller M, Lang F. Beneficial effect of aurothiomalate on murine malaria. Malar J. 2010;9:118.

95 

 

Appendix I

Laboratory chemicals

Acetic acid (100%, glacial) Merck Eurolab GmbH

Adenine Sigma, Deisenhofen

Agarose Carl Roth, Karlsruhe

Azure B Sigma-Aldrich, Taufkirchen

Albumax II Gibco-Invitrogen, Karlsruhe

Annexin V-FLUOS Boehringer, Mannheim

Annexin V-568 Roche, Mannheim

Amphtericin B Sigma, Deisenhofen

t-butylhydroperoxide (8 M) Sigma, Deisenhofen

CaCl2 Sigma, Deisenhofen

Cardiolipin (from bovine heart) Sigma, Munich

Citrate-Phosphate-Dextrose-Stabiliser Baxter S.A. (Fenwal), Maurepas, France

DETAPAC (C14H23N3O10) Sigma (Fluka), Munich

Dextrose monohydrate (D (+) glucose) Sigma, Deisenhofen

DL-Dithiothreitol (DTT) minimum 99 % Sigma, Deisenhofen

2,4 –Dichloroisocoumarin Biomol GmbH, Hamburg

Dipotassium hydrogen phosphate Merck KGaA, Darmstadt

Dimethylsulfoxid (DMSO) Sigma, Deisenhofen

Disodium hydrogen phosphate anhydrous Merck KgaA, Darmstadt

EDTA Sigma, Steinheim

Ethanol Merck Eurolab, Darmstadt

Ethidium bromide (10 mg/ml) Carl Roth, Karlsruhe

Fluo-3/AM Calbiochem, Bad Soden

Gas mixture (90% N2 / 5% O2 / 5% CO2) Hoepfner, Reutlingen; Mast, Tuebingen Gentamicin sulphate Gibco InVitrogen, Karlsruhe

Glutamine Gibco InVitrogen, Karlsruhe

96 

 

Giemsa staining solution Sigma, Deisenhofen

Glycerol Sigma-Aldrich, Taufkirchen

Gum Arabic Dar Savana, Sudan

HCL Merck Eurolab, Darmstadt

HEPES Sigma, Deisenhofen

HEPES/NaOH 1M solution Sigma, Deisenhofen

Hypoxanthine Sigma, Deisenhofen

Ionomycin Sigma, Munich

Immersion oil Type A Cargille Laboratories, Cedar Grove, NJ, USA

Inosine Sigma-Aldrich, Taufkirchen

Instamed RPMI 1640 with Glutamine Gibco Invitrogen, Karlsruhe

Isopropanol Merck Eurolab, Darmstadt

KCl Sigma, Deisenhofen

Mannitol Sigma, Deisenhofen

Methanol Merck KgaA, Darmstadt

Methylene blue Sigma-Aldrich, Taufkirchen

Multi Twist Top Vials 1.7 ml Roth, Karlsruhe

Multi Twist Top Vial Caps Roth, Karlsruhe

NaCl Sigma, Deisenhofen

Na-gluconate Sigma, Deisenhofen

NaHCO3 Sigma, Deisenhofen

NaOH Sigma Diagnostics, St Louis, MO, USA

Na-vanadate Sigma-Deisenhofen

Nitrogen (Gas) Mast, Tuebingen

NMDG-Cl Sigma, Deisenhofen

n-octylglucopyranoside (C14H28O6) Sigma (Fluka), Munich

PBS Gibco BRL, Karlsruhe

Potassium dihydrogen phosphate Merck Eurolab, Darmstadt

97 

 

Propidiume iodide Sigma, Munich

SAG-Mannitol solution Baxter S.A. (Fenwal), Maurepas, France

Sodium butyrate Sigma, Deisenhofen

Spiroepoxide Alexis, Gruenberg

Sphingomyelinase (50 U) Biomol, Hamburg

1-Stearoyl-2-arachidonoyl-sn-glycerol Alexis, Lausen, Switzerland

Syto16 (1mM in DMSO) Invitrogen (Molecular Probes), Karlsruhe

Sorbitol Sigma, Deisenhofen

Stock materials 6-, 12-, 24- & 96-well plates Greiner Bio-One, Frickenhausen

Bottle-top filters for 0.125 l Millipore, Schwalbach

Bottle-top filters for 0.25 and 0.5 l Carl Roth, Karlsruhe

CryoTube Vials 2.0 ml Greiner Bio-One, Frickenhausen

EDTA tubes Monovette, Sarstedt

FACS tubes Greiner Bio-One, Frickenhausen

Falcon tubes Greiner Bio-One, Frickenhausen

Microcentrifuge Filters Ultrafree-MC,

NMWL 5.000 Dalton, PLCC

cellulosic membrane Sigma, Taufkirchen

MµlTI Twist Top Vials (1.7 ml;

10 x 45 mm polypropylene) Roth, Karlsruhe

MµlTI Twist Top Vial Caps Roth, Karlsruhe

Syringe sterile filter (0.22 µm) Millipore, Schwalbach

Scintillation vials PerkinElmer (LAS), Rodgau-Juegesheim

Tissue culture flasks Nunc, Wiesbaden

Transfer pipettes, polyethylene, extended

fine tip, large bulb, Bulb: 3 ml, sterile Sigma, Deisenhofen

Tissue culture flasks Greiner Bio-One, Frickenhausen

98 

 

Devices

Adapter for gas dosing unit EC

for 24 Pasteur pipettes (V826.612.000) VLM, Leopoldshöhe

Autoclave (with outlet air bacterial filter) Systec Labor Systemtechnik, Wettenberg

8-channel-multi-pipette Research pro Eppendorf, Hamburg

Bath sonicator (Transsonic 310) Elma, Singen

Biofuge pico Heraeus Kendro laboratory products, Hanau

β-scintillation counter Wallac 1409 PerkinElmer (LAS), Rodgau-Juegesheim

Concentrator 5301 Eppendorf, Hamburg

Dissolved Oxygen Meter (DO-5509) Lutron, Copersburg, PA, USA

Horizontal DMZ puller Zeitz, Augsburg

Evaporator EVA-EC1-24-S (V832.000.002) VLM, Leopoldshöhe

FACS Calibur Becton Dickinson, Heidelberg

Hamilton microliter syringe(100 µl, 250 µl) Carl Roth, Karlsruhe

Hera cell incubator 37 °C Kendro Laboratory Products, Langenselbold

Heraeus Sepatech Centrifuge Kendro laboratory products, Hanau

Light microscope Leica CM E Leica, Solms

Rotina35 centrifuge Hettich GmbH & Co KG, Tuttlingen

Safety cabinet class II (Hera Safe) Kendro Laboratory Products, Langenselbold

Scanning Spectrophotometer Thermo Electron, Bremen

Thermomixer 5436 Eppendorf, Hamburg

Media, buffers and solutions

Solutions for preparing human erythrocytes for storage

Citrate-Phosphate-Dextrose-Stabilizer (g/l aqua ad iniectabilia)

Citrate acide – monohydrate 3.27

Sodium citrate – monohydrate 26.30

Sodium dihydrogen phosphate – dehydrate 2.50

Dextrose monohydrate 25.50

99 

 

SAG-Mannitol solution (g/l)

NaCl 8.77

Dextrose monohydrate (D (+) – glucose) 9.00

Adenine 0.169

Mannitol 5.25

Citrate-Phosphate-Dextrose-Stabilizer (70 ml) and SAG-Mannitol (SAGM) solution (110

ml) were obtained sterile in pockets connected by thin tubes, separated by a leukocyte

filter, ready for a blood donation of 500 ml.

Sterile saline solution (in mM):

NaCl 150

KCl 5

CaCl2 1.4

MgCl2 1

HEPES/NaOH pH 7.4 10

Glucose 10

The saline solution was prepared according to Egee et al. {Egee, 2002 484 /id}. The pH

was adjusted to 7.4 and the osmolarity to 320 ± 5 mOsm (kg H2O)-1 before sterile filtering

the saline solution through a filter unit of 0.22 µm pore size.

Media and solutions for the maintenance of P. falciparum in vitro culture

Supplemented RPMI 1640 medium

HEPES/NaOH 25 mM

Gentamicin sulphate 20 µg /ml

Glutamine 2 mM

Hypoxanthine 200 µM

Albumax IITM 0.5 %

1 M sterile filtered CaCl2 (0.22 µm pore size) was used for preparing human serum from

plasma.

100 

 

Albumax stock solution 10 x (in g/l):

Instamed RPMI 1640 with Glutamine

NaHCO3 10.43

HEPES 5.96

D (+) glucose 2

Hypoxanthine 0.2

Albumax IITM 50

Gentamicin sulphate 0.01

The pH was adjusted to 7.1 - 7.4. The sterile filtered Albumax stock solution was stored in 50 ml aliquots at -20 or – 80 °C.

Freezing solution (in %) Glycerol 28

Sorbitol 3

NaCl 0.65

The freezing solution was sterile filtered (through 0.22 µm pore size) and stored at 4 °C.

Defreezing solution (in %)

NaCl 3.5

The sterile filtered defreezing solution (through 0.22 µm pore size) was stored at room temperature (RT). The solution was warmed up to 37 °C before use.

Solutions to analyze blood smears of P. falciparum infected RBCs

Dipotassium hydrogenphosphate 1 M

Potassium dihydrogenphosphate 1 M

Potassium phosphate buffer of pH 7.2 0.1 M was prepared according to Sambrook et al.

{Sambrook, 1989 1205 /id}. GIEMSA staining solution was freshly prepared from the

purchased GIEMSA staining solution by dilution 1:10 with 0.1 M potassium phosphate

buffer pH 7.2, filtered over a paper filter.

101 

 

Solutions for P. falciparum in vitro growth assays

All solutions were sterile filtered (through 0.22 µm pore size) and added to the

supplemented RPMI1640 media. Changes in media concentration resulting from the

addition of solutions were tolerated up to a concentration loss of 10%. In cases where the

addition of the stock solution to the RPMI medium would have diluted the supplemented

RPMI medium by more than 10%, the relevant chemical was added directly to the

supplemented medium, which was then sterile-filtered again.

For the test solutions the different stock solutions or chemicals were diluted with sterile distilled water to obtain the final concentrations shown below.

Stock solutions (1 M):

CaCl2

KCl

NaCl

Na-gluconate

NMDG-Cl titrated with HCl to pH 7.4

D (+) glucose

HEPES (titrated with TRIS (Trizma base) or with NaOH to pH 7.4)

Synchronization solution (in mM)

Sorbitol isosmotic (5%) 290

HEPES / NaOH pH 7.4 10

Glucose 5

When erythrocytes were to be used further, e.g. for western blots (membrane proteins) or for measuring 45Ca2+ uptake, the 5% sorbitol solution was supplemented with HEPES and glucose. This was done to prevent infected RBCs from being stressed or oxidized through glucose depletion.

Ca2+- Ringer (in mM)

NaCl 140

KCl 5

HEPES 10

MgCl2 1

CaCl2 1

102 

 

Glucose 5

Hyperosmolar Ca2+- Ringer (in mM)

NaCl 140

KCl 5

HEPES 10

MgCl2 1

CaCl2 1

Glucose 5

Sucrose 650

NaCl test solution (in mM)

NaCl 120

HEPES / TRIS (titrated with TRIS to pH 7.4) 30

KCl 5

Glucose 5

CaCl2 1.7

Solutions for fluorescence assisted cell sorting (FACS) analysis

Annexin binding buffer (in mM)

NaCl 140

HEPES / NaOH pH 7.4 10

CaCl2 5

PBS (in M)

NaCl 0.14

PO4 Buffer pH 7.4 0.01

KCl 0.0003

Each tablet of 5 g PBS had to be dissolved in 500 ml distilled water, giving a final pH 7.45.

Syto16 was diluted before use in a dark environment in PBS or annexin binding buffer to a

final concentration of 10 nM - 1 µM.

103 

 

Appendix II

Protocol (1): Preparation of human and mouse erythrocytes (Washing of RBCs)

1. Take fresh heparinised blood from the transfusion centre.

2. Transfer the heparinised blood into the 50 ml falcon tube and centrifuge it at

2600 rpm for 4 min without any break.

3. Separate the plasma from the cells in the falcon tube.

4. Now add double volume of RBC wash to the cells in the falcon tube and gently

mix it.

5. Again centrifuge the mixture at 2600 rpm for 4 min.

6. Repeat 4th and 5th steps for three times…

7. Finally store the washed RBC in the 4oC for further experimental procedures.

Protocol (2) Infection of washed RBCs with Plasmodium falciparum

Check the parasitemia of the culture flask in the FACS machine using Syto 16

1. Take into account what percent of parasitemia is needed for experimentation.

2. Split the culture into the new sterile flasks

I. Remove the supernatant from the old flask so that the dead RBC and the

metabolic wastes of the pathogen may be cleared off.

II. Gently tilt the flask and measure the pellet volume in the flask (M).

3. Follow the practical protoco

Calculations:

Then,

Amount of old culture to be added from the old culture flask…

= OB/500µl x M

104 

 

Procedure:

1. In the laminar flow hood, label sterile culture flasks clearly.

2. Add 25 ml RPMI complete into the labelled culture flasks.

3. Add calculated amount of fresh blood to each labelled flask.

4. Add the calculated amount of old blood from the old culture flasks into the new

culture flasks.

5. Gently tilt the flasks.

6. Gas the culture flasks with 5% CO2, 5% O2 and 90% N2 for 1 min.

7. Finally incubate the culture flasks at 37oC.

Protocol (3): Thawing parasite and culture

1. Take a 15 ml sterile falcon tube.

2. Adjust the water bath for 37oC.

3. Take one parasites cryotube from the liquid nitrogen and thaw at water bath for 5 min

4. Spray the disinfectant on the thawed tube.

5. Gently uncap the tube, with the help of a pipette collect the thawed blood and add it

into the 15ml falcon tube.

6. Discard the thawed tube.

7. Centrifuge the 15 ml falcon tube with thawed blood at 1800 rpm for 4 min.

8. Remove the supernatant from the falcon tube. (if there is no supernatant remove the

blood till there is at most 500µl of it left in falcon tube)

9. Add equal amount of sterile 3.5% sodium chloride solution drop/sec into the falcon

tube. (This kills all other cells but keeps alive the parasites.)

10. Mix gently and then centrifuge at 1800 rpm for 4 min.

11. Wash the pellet with RPMI normal twice to remove the traces of sodium chloride

and maintain the tonicity of the solution.

12. Estimate the pellet and remove the supernatant, now add almost excess double the

volume of RPMI normal to the volume of pellet and mix gently

13. Centrifuge the falcon at 1800 rpm for 4 min.

14. Remove the supernatant.

15. Suspend the pellet in 3 ml of RPMI complete medium.

105 

 

16. Transfer 12 ml of the RPMI complete medium into the cultured flasks.

17. Add 180µl of fresh blood to the culture flask.

18. Add the pellet to the cultured flask.

19. Gas the culture with 5% CO2, 5% O2 and 90% N2 for 1 min. and keep in the BOD

for incubation.

Protocol (4): Isosmotic sorbitol synchronization of P. falciparum infected human

erythrocytes for in vitro growth assay or DNA amplification and double staining

Select low parasitemia culture (3-5%) for in vitro growth assay or a high parasitemia

culture (20-30%) for DNA amplification and double staining techniques. (double staining

to detect PS exposure).

1. Remove the supernatant from the selected culture flask

2. Collect the blood in the sterile 50 ml falcon tube.

3. Centrifuge the falcon tube at 2000 rpm for 4 min.

4. Remove the supernatant an estimate the volume of the pellet.

5. Add 5ml of sorbitol to 500µl of pellet. (Synchronisation) only the ring stages and

non- infected RBC are left out.

6. Transfer the pellet sorbitol mixture in to 25 ml falcon tube.

7. Put on the Wave Tec machine for synchronisation up to 20 min.

8. Again centrifuge the falcons at 2000 rpm for 4 min.

9. Remove the supernatant and add 10 ml of normal RPMI-1640 medium to remove

the remaining sorbitol (the tube containing only ring stages and non-infected RBC).

10. Centrifuge again for 4 min at 2000 rpm.

11. Transfer the pellet into the small eppendorf tube and wash again the ring stages and

non-infected erythrocytes with normal RPMI-1640 medium and centrifuge at 2000

rpm for 2 min.

12. Remove the supernatants from the eppendoff tube and mix the pellet gently.

13. Keep the eppendoff tube in the refrigerator at 4oC.

14. Now the pellet is ready for DNA amplification and double staining.

106 

 

Protocol (5) In vitro growth assay

Selection of low parasitemia culture (3-5%) for in vitro growth assay technique.

1. Remove the supernatant from the selected culture flask

2. Collect the blood in the sterile 50 ml falcon tube.

3. Centrifuge the falcon tube at 2000 rpm for 4 min.

4. Remove the supernatant and estimate the volume of the pellet.

5. Add 5ml of sorbitol to 500µl of pellet. (Synchronisation) only the ring stages and

non- infected RBC are left out.

6. Transfer the pellet sorbitol mixture in to 25 ml falcon tube.

7. Put on the Wave Tec for synchronisation up to 20 min.

8. Again centrifuge the falcons at 2000 rpm for 4 min.

9. Remove the supernatant and add 10 ml of normal RPMI-1640 to the falcon and

wash the ring stage.

10. Centrifuge again for 4 min at 2000 rpm.

11. Transfer the pellet into the small eppendorf tube and wash again the erythrocytes

with normal RPMI-1640 and centrifuge at 2000 rpm for 2 min.

12. Remove the supernatants in the eppendoff tube.

13. Keep the eppendorf tube in the refrigerator at 4oC.

14. Now the pellet is ready for in vitro growth assay.

Procedure:

1. Collect the low parasitemia blood pellet and store in the refrigerator in 4oC.

2. Label the required 12ml PP-tubes according the drug concentrations.

3. Put 2 ml of complete RPMI into each labelled PP-tubes.

4. Add respective drug concentrations in to the RPMI containing labelled PP-tubes.

5. Vortex the falcons for homogenous mixture of the drug in the RPMI.

6. Add 20 µl of low parasitemia blood pellet to the above falcons and mix gently with

a pipette.

7. Take a sterile 96 well plate and label it.

8. Put 200µl of the RPMI+ high parasitemia blood pellet + drug into each well. Avoid

the bubble formation in the wells.

107 

 

9. Incubate for 48 hrs in the candle jar in the BOD.

10. After 48 hrs, take slowly the incubated 96 well plate and gently mix the culture in

each well.

11. Take 50µl of the culture from wells of each concentration and put in labelled

FACS falcon tubes.

12. Now add 200µl of PBS + EDTA with Syto-16 to each falcon tube and incubate

for 20 min.

13. Analyse the sample with FACS machine.

Protocol (6): Measurement of percentage parasitemia by Syto-16

1. Preparer the required amount of BPS containing Syto 16 by the ratio of 1000:1 (the

volume of BPS required for each tube is 250 µl.)

2. Label the FACS (Fluorescence Activated Cell Sorting) tubes.

3. Take small amount (~20 µl) of infected blood from the cultured flasks.

4. Add Syto-16 to PBS with Syto 16 to FACS tube containing infected blood and

vortex.

5. Incubate the FACS tubes in the BOD (Biochemical Oxygen Demand) 37oC for 30

min.

6. Estimate the percent parasitemia in each cultured flasks with the help of FACS

machine.

Protocol (7): Giems staining technique

1. Put a drop of sample on a clean and sterile slide with the help of transferring

pipette.

2. Make a thin film smear on the slide with the help of another slide

3. Air dry the smear on the slide.

4. fix the smear in methanol for 1 min.(by sprinkling few drops on the smear)

5. Dry again the slide and immerse the slide in Giems + May Grinwald (10+90

ml) mixture for 30 min.

6. wash the slide gently under running tap water and observe under the

microscope at 100X

108 

 

Protocol (8): Annexin binding experiments (Determination of phosphatidylserine

exposure by double staining technique)

1. Collect the high parasitemia blood pellet and store in the refrigerator in 4oC.

2. Label the required 12 ml PP-tubes according to the drug concentrations

required.

3. Put 2 ml of complete RPMI into each labelled falcon.

4. Add respective drug concentrations in to the RPMI containing labelled PP-

tubes.

5. Vortex the falcons for homogenous mixture of the drug in the RPMI.

6. Add 20µl of high parasitemia blood pellet to the above PP-tubes and mix gently

with a pipette.

7. Take a sterile 96 well plate and label it.

8. Put 200µl of the RPMI+ high parasitemia blood pellet+ drug into each well.

Avoid the bubble formation in the wells.

9. Incubate for 24 hrs in the candle jar in the BOD.

10. After 24 hrs, take slowly the incubated 96 well plate and gently mix the culture

in each well.

11. Take 50µl of the culture from any of the two wells from each concentration and

put in labelled eppendorf tubes.

12. Add 200µl of annexin wash buffer to each eppendorf tube and mix well.

13. Centrifuge at 2000 rpm for 5 min.

14. Remove the supernatant with help of a vacuum pump and retain the pellet.

15. Add 5µl of annexin APC to each pellet and mix.

16. Add 100 µl of annexin wash buffer with syto-16 to each eppendorf tube and

incubate for 20 min in the incubator.

17. Transfer the incubated mixture into small falcons and add 100µl of annexin

wash buffer to each small falcon before analysing with FACS.

18. Prepare some extra single stain samples for each syto-16 and annexin for

adjusting the settings for FACS.

109 

 

Protocol (9): Haemolysis test

1. Prepare 6 samples each containing 12.5 µl of pellet containing 5% of

hematocrit. (total pellet required for 6 samples is 75 µl)

2. Add 200 µl of ringer to each sample in sterile eppendorf.(total ringer required is

1200 µ)

3. Incubate the samples in the incubator for 24 hrs at 37 oC.

4. After incubation gently tilt the samples and centrifuge for 5 min at 2000 rpm.

5. Collect the supernatant and transfer it into cuvettes.

6. Add 550µl of transferring reagent to each sample.

7. Prepare a standard curve using control samples.

a. After removing the supernatant from the eppendorf add 200µl of

millipore water and gently mix.

b. Transfer it into another eppendorf and add 200 µl of millipore water.

c. Make the required concentrations to establish a standard curve.

8. Measure the absorbance of the samples in cuvettes at 546 nm using

spectrophotometer.

Protocol (10): Preparation of RPMI complete medium

Requirements:

Glutamine (2 mM) 5 ml

HEPES/ NaOH pH 7.4 (25 mM) 12.5 ml

Gentamycin (20µg/ml) 0.5ml

Albumax (0.5%) 50 ml

1. Take fresh RPMI-1640 from the cold room, falcon of 50 ml albumax 10X from

refrigerator and 5 ml of glutamine from -20oC, allow to worm in the water bath

to 37oC.

2. Take glutamine from the -20 oC refrigerator.

3. In the laminar flow hood, gently mix albumax and glutamine with the worm

RPMI-1640.

4. Then add 12.5 ml HEPES and 0.5 ml Gentamycin

5. Mix well.

6. Store now the RPMI complete medium in 4oC for further experimental

procedures.

110 

 

Protocol (11): Preparation of RBCs wash solution

Requirements:

150 mM NaCl (54.44) 8.166 g

5 mM KCl (74.55) 0.3727 g

1.4 mM CaCl2 (110.98) 0.15537 g

1 mM MgCl2 (95.21) 0.09521 g

10 mM HEPES (238.30) 2.38 g

10 mM Glucose (198.17) .9817 g

1. Weigh accurately all the required chemicals and put in a sterile 1000 ml bottle.

Then add millipore distilled water. (mix with magnetic stirrer to homogenise the

solution )

2. Adjust the pH of the solution upto7.4, initial pH will be up to 5.34 then add 1M

NaOH to make it 7.4.

3. Then check the osmolarity.

4. Calibrate the osmolity with 290 and 1000 then check the osmolity of the RBC

wash solution. It must be within 290-310. (If slightly above 310 add normal

distilled water to adjust.)

5. After preparation of the homogenous solution filter it with millipore (0.22µM)

using vacuum in sterile conditions.

6. Now store it in clean and dry place.