congenital transmission by protozoandownloads.hindawi.com/journals/specialissues/829362.pdf ·...

Congenital Transmission by ProtozoanGuest Editors: Ricardo E. Fretes, Ulrike Kemmerling, and Demba Sarr

Journal of Tropical Medicine

Congenital Transmission by Protozoan

Journal of Tropical Medicine


Guest Editors: Ricardo E. Fretes, Ulrike Kemmerling,and Demba Sarr

Copyright © 2012 Hindawi Publishing Corporation. All rights reserved.

This is a special issue published in “Journal of Tropical Medicine.” All articles are open access articles distributed under the CreativeCommons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the originalwork is properly cited.

Editorial Board

Hans Peter Beck, SwitzerlandSukla Biswas, IndiaJoseph Blaney, USAIb Christian Bygbjerg, DenmarkMaureen Coetzee, South AfricaCarlos E. P. Corbett, BrazilRodrigo Correa-Oliveira, BrazilLuis E. Cuevas, UKAditya Prasad Dash, IndiaIbrahim M. Elhassan, Saudi ArabiaBlaise Genton, Switzerland

Jean-Paul Gonzalez, FranceJoseph Hamburger, IsraelLukasz Kedzierski, AustraliaSusana A. Laucella, ArgentinaPeter Leggat, AustraliaSylvie Manguin, FranceWilbur Milhous, USALouis H. Miller, USAMathieu Nacher, French GuianaChristian F. Ockenhouse, USAKenneth E. Olson, USA

John Henry Ouma, KenyaGerd Pluschke, SwitzerlandSasithon Pukrittayakamee, ThailandR. G. Ridley, USAGeorges Snounou, FrancePradya Somboon, ThailandShyam Sundar, IndiaMarcel Tanner, SwitzerlandThomas R. Unnasch, USADina Vlachou, UK

Contents

Congenital Transmission by Protozoan, Ricardo E. Fretes, Ulrike Kemmerling, and Demba SarrVolume 2012, Article ID 173437, 2 pages

Mechanism of Trypanosoma cruzi Placenta Invasion and Infection: The Use of Human Chorionic VilliExplants, Ricardo E. Fretes and Ulrike KemmerlingVolume 2012, Article ID 614820, 7 pages

Transplacental Transmission of Plasmodium falciparum in a Highly Malaria Endemic Area of BurkinaFaso, Alphonse Ouedraogo, Alfred B. Tiono, Amidou Diarra, Edith C. Christiane Bougouma, Issa Nebie,Amadou T. Konate, and Sodiomon B. SirimaVolume 2012, Article ID 109705, 7 pages

Reorganization of Extracellular Matrix in Placentas from Women with Asymptomatic Chagas Disease:Mechanism of Parasite Invasion or Local Placental Defense?, Juan Duaso, Erika Yanez, Christian Castillo,Norbel Galanti, Gonzalo Cabrera, Gabriela Corral, Juan Diego Maya, Ines Zulantay, Werner Apt,and Ulrike KemmerlingVolume 2012, Article ID 758357, 8 pages

Prevention of Congenital Transmission of Malaria in Sub-Saharan African Countries: Challenges andImplications for Health System Strengthening, Kayode O. Osungbade and Olubunmi O. OladunjoyeVolume 2012, Article ID 648456, 6 pages

In Vitro Infection of Trypanosoma cruzi Causes Decrease in Glucose Transporter Protein-1 (GLUT1)Expression in Explants of Human Placental Villi Cultured under Normal and High GlucoseConcentrations, Luciana Mezzano, Gaston Repossi, Ricardo E. Fretes, Susana Lin, Marıa Jose Sartori,and Sofıa G. Parisi de FabroVolume 2012, Article ID 969243, 8 pages

Prevalence of Congenital Malaria in Minna, North Central Nigeria,Innocent Chukwuemeka James Omalu, Charles Mgbemena, Amaka Mgbemena, Victoria Ayanwale,Israel Kayode Olayemi, Adeniran Lateef, and Victoria I. ChukwuemekaVolume 2012, Article ID 274142, 5 pages

Hindawi Publishing CorporationJournal of Tropical MedicineVolume 2012, Article ID 173437, 2 pagesdoi:10.1155/2012/173437

Editorial


Ricardo E. Fretes,1, 2 Ulrike Kemmerling,3 and Demba Sarr4

1 Cell Biology, Histology, and Embryology Department, Medicine School, Universidad Nacional de Cordoba, 5000 Cordoba, Argentina2 Universidad Nacional de La Rioja, 5300 La Rioja, Argentina3 Program of Anatomy and Developmental Biology, Institute for Biomedical Sciences, Faculty of Medicine, University of Chile,8380453 Santiago, Chile

4 Department of Infectious Diseases and Center for Tropical and Emerging Global Diseases, University of Georgia, Athens,GA 30602, USA

Correspondence should be addressed to Ricardo E. Fretes, [email protected]

Received 12 September 2012; Accepted 12 September 2012

Copyright © 2012 Ricardo E. Fretes et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

Congenital transmission by protozoan parasites is a world-wide important public health problem. Congenital infectionsaffects the mother and the fetus or newborn. It is stillsurprising that despite the abundant immunoepidemiolog-ical knowledge of congenital transmission of protozoanparasite, no definite etiology or predictive diagnostic testshave been identified. Understanding the mechanisms bywhich host/parasites infection and interaction occurs is oneof the most important topics that will help find specificbiomarkers of infection, prevent congenital transmission,maintain the health of the newborn, and develop safeand efficient treatments. In addition, understanding of thebiological mechanisms of host/parasite interactions willfacilitate protection of mothers and their families and reducecosts in health services. Moreover, this will lead to gaininsight into epidemiological aspects and association withother pathologies and preserve the wellbeing of the newborn.In this special issue we have invited a few papers that addresssuch issues.

Congenital malaria is underestimated and usually asso-ciated with low-density cord parasitemia. However, it isincreasingly recognized as a potentially serious complicationof maternal malaria during pregnancy in Sub-SaharianAfrica. Earliest epidemiological studies have reported preva-lence varying widely in malaria-endemic areas from 0% to33%. At birth, infections are usually asymptomatic with lowparasitemia and the diagnosis by microscopy is often missed.Infection may occur by transplacental passage of parasitesduring disruption of the placental barrier at the time ofdelivery, with subsequent clinical illness in the newborn baby.

A paper of this issue assessed the prevalence of congenitalmalaria during the dry season (period of low-mosquitodensity and low-malaria incidence) using peripheral andcord blood smears of new born babies in association withperipheral and placental blood smears of, respectively, near-term and term pregnant women. This study is interestingbecause the authors found that congenital malaria is not rarein their study area. Another paper is a literature review onthe prevalence, burden, diagnosis, prevention, and controlof congenital malaria in Sub-Saharian Africa. This review isof interest to individuals working in clinics and laboratorydiagnostic of malaria but also to national governmentsand partners in Sub-Saharian Africa. The authors highlightthe challenges in parasitological and clinical diagnosis, thechallenges in prevention with the use of intermittent pre-ventive treatment (IPT) and insecticide-treated nets (ITNs)and provides recommendation on how to strengthen thehealth system in Africa. Another paper reports the prevalenceof transplacental malaria in Burkina Faso, a West Africancountry, and determines the real burden of transplacentaltransmission. The authors show associations between levelsof parasite in the maternal, placental, and umbilical cordblood. All papers about malaria published in this specialissue show the interest of pursuing investigations for a betterunderstanding of vertical transmission by malarial parasites.

During congenital Chagas transmission, the parasitereaches the fetus by crossing the placental barrier. In the pastfew years congenital transmission of T. cruzi has increasinglybecome more important, and partly responsible for the“globalization of Chagas’ disease,” constituting a public

2 Journal of Tropical Medicine

health problem of increasing relevance. The fact that onlya percentage of the infected mothers transmit parasitesto their fetuses raises the question of the ability of theplacenta as well as the immunological status of motherand fetus/newborn to impair the parasite transmission.Therefore, it is thought that congenital Chagas disease isthe product of a complex interaction between the parasite,the maternal, and fetus/newborn immune responses, andplacental factors. Additionally, a paper of this special issueconstitutes a morphological analysis by immunohistochem-ical and histochemical methods of placentas from womenwith chronic Chagas’ disease. T. cruzi-infected placentaspresent destruction of the syncytiotrophoblast and villousstroma, selective disorganization of the basal lamina, anddisorganization of collagen I in villous stroma. Changes inthe extracellular matrix of placental tissues, together with theimmunological status of mother and fetus, and parasite loadmay determine the probability of congenital transmissionof T. cruzi. Another paper assayed the effect of T. cruzion glucose transporter protein-1 (GLUT1), which is themain isoform involved in transplacental glucose transport.High glucose media as well as T. cruzi infection reduceGLUT1 expression. The effect of T. cruzi infection on GLUT1expression may explain some of the clinical manifestationof congenital Chagas disease. Finally, a paper analyzing thecongenital transmission of Chagas disease is a review aboutthe role of possible local placental factors that contributeto the vertical transmission of the parasite. Additionally,in that review different available methods for studying thecongenital transmission of T. cruzi are analyzed. In thatcontext, the ex vivo infection of human placental chorionicvilli by T. cruzi trypomastigotes constitutes an excellent toolfor studying parasite infection strategies as well as possiblelocal antiparasitic mechanisms.

Ricardo E. FretesUlrike Kemmerling

Demba Sarr


Review Article

Mechanism of Trypanosoma cruzi Placenta Invasion andInfection: The Use of Human Chorionic Villi Explants

Ricardo E. Fretes1, 2 and Ulrike Kemmerling3

1 Department of Histology and Embryology, Faculty of Medicine, Universidad Nacional Cordoba, 5000 Cordoba, Argentina2 IICSHUM and Cathedra of Histology, Embryology and Genetic, Health Department, Universidad Nacional La Rioja,5300 La Rioja, Argentina

3 Program of Anatomy and Developmental Biology, Institute for Biomedical Sciences, Faculty of Medicine, University of Chile,8380453 Santiago, Chile

Correspondence should be addressed to Ricardo E. Fretes, [email protected] Ulrike Kemmerling, [email protected]

Received 28 November 2011; Revised 6 February 2012; Accepted 14 February 2012

Academic Editor: Demba Sarr

Copyright © 2012 R. E. Fretes and U. Kemmerling. This is an open access article distributed under the Creative CommonsAttribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work isproperly cited.

Congenital Chagas disease, a neglected tropical disease, endemic in Latin America, is associated with premature labor andmiscarriage. During vertical transmission the parasite Trypanosoma cruzi (T. cruzi) crosses the placental barrier. However, theexact mechanism of the placental infection remains unclear. We review the congenital transmission of T. cruzi, particularly therole of possible local placental factors that contribute to the vertical transmission of the parasite. Additionally, we analyze thedifferent methods available for studying the congenital transmission of the parasite. In that context, the ex vivo infection with T.cruzi trypomastigotes of human placental chorionic villi constitutes an excellent tool for studying parasite infection strategies aswell as possible local antiparasitic mechanisms.

1. Introduction

Chagas disease was first described by the Brazilian physician,Carlos Chagas, in 1909. He identified the causal agent,the protozoan Trypanosoma cruzi (T. cruzi), the vectorialtransmission and insect reservoirs as well as the clinical signsand symptoms. In other words, he described the completecycle of the disease and suggested the possibility of congenitaltransmission [1].

In the human villous hemochorial placenta, fetal andmaternal tissues are separated by a fetal epithelium (thetrophoblast). Within the villous placenta, a single multin-ucleated cell layer (syncytiotrophoblast) contacts maternalblood within the intervillous space. Beneath the syncytiotro-phoblast reside replicating progenitors (cytotrophoblast)that are separated by a basal lamina from the connec-tive tissue of villous stroma containing vascular endothe-lium, fibroblasts, and macrophages. The syncytiotrophoblastforms a surface of about 12 m2 that contacts maternal

blood. Therefore, in case of women with Chagas disease, theparasite has the opportunity to interact with a large cellularsurface.

The mechanism of T. cruzi congenital transmission canbe studied in different ways. One of the possible ways isthrough the dual placental perfusion system. The dual per-fusion that simulates the maternal and fetal circulation couldbe an excellent method to study the mechanism of the trans-mission of the parasite [2]. However, this method requirescomplex equipment and very experienced users. Anotherpossibility is the ex vivo infection of human chorionic villiexplants, in which samples of placental tissue (chorionic villi)can be challenged in vitro with T. cruzi or other pathogens.This system is preferred to analyze the process of infection ofthe placental barrier, although the immune system does notparticipate. The analysis of infection in animal models is nottreated in this paper because their placentas are not similarto human placenta. Also, pathological findings of placentasare not deeply described because they represent the final


stage of the placental infection process, and so, it is hard tounderstand an ongoing process.

The aim of the present paper is to review the congenitaltransmission of T. cruzi, particularly the role of possiblelocal placental factors which contribute to the verticaltransmission of the parasite and can be studied in the ex vivoinfection model of human chorionic villi explants.

2. Chagas Disease

Chagas disease develops in three phases. The acute phase isfirst developed, immediately after infection, with high levelsof parasitemia and symptoms in only some patients (regionallymph node enlargement, bipalpebral unilateral edema, orRomana’s sign, and characteristic electrocardiogram alter-ations). In most cases, acute infection is not accompaniedby clinical findings, thus moving onto the latent phase thatcan last for months or years. The chronic phase, present in30% of infected individuals, is associated with mega colon,mega esophagus, degeneration of the autonomous nervoussystem, arrhythmias, and abnormal growth of the heartwith progressive insufficiency, with evident negative impacton the patient’s health. In this phase the disease can behandicapping, and can either be the concurrent or the directcause of death. The course of the disease depends on diversefactors: parasite load at the site of inoculation, both theparasite’s genetic group and strain, whether it is an infectionde novo or reinfection, the host’s immunologic status, and thetype of vector (triatomid) [3, 4].

3. Congenital Chagas Disease

Congenital T. cruzi infection is associated with prematurelabor, low birth weight, and stillbirths [5–7]. Serologicprevalence among pregnant women may reach 80%, andrates of congenital infection vary from 1–21% [8–11]. InArgentina the transmission rate is estimated between 2–12%[12]. In Chile, in two of the endemic regions (IV and Vregions), the congenital transmission rate of the parasiteis 8.4% [13]. According to WHO/PAHO the number ofinfected women at fertile age is approximately 1.8 millionand it is estimated that 14,400 neonates are being infectedeach year [14], this is another reason why this form oftransmission becomes epidemiologically more important.Additionally, congenital transmission is partly responsiblefor the “globalization of Chagas’s disease” [10, 15].

Congenital Chagas transmission implies a T. cruziseropositive mother and a postpartum detection of parasitesin the newborn. Congenital Chagas disease is diagnosed bydirect microscopic examination of blood samples, PCR, orstandard serological assays. The latter can be carried out ininfants when IgG antibodies transferred from the motherhave been eliminated (8-9 months after birth).

During congenital transmission, the parasite reaches thefetus by crossing the placental barrier [16–22]. The factthat only a percentage of the infected mothers transmitparasites to their fetuses raises the question of the ability ofthe placenta as well as the immunological status of motherand fetus/newborn to impair the parasite transmission.

Therefore, it is thought that congenital Chagas disease isthe product of a complex interaction between the parasite,the maternal and fetus/newborn immune responses, andplacental factors [9, 18].

4. The Parasite

T. cruzi is a haemoflagellated protozoan of the Kinetoplastidaorder and Trypanosomatidae family [22]. The parasite’sbiological cycle includes three cellular forms characterizedby the relative positions of the flagellum, kinetoplast, andnucleus [23]: (1) trypomastigotes: approximately 20 µmin length and sub terminal kinetoplast. They constitutethe nonreplicative, mammalian infecting cellular form thatis found in the blood and in the posterior intestine oftriatomids. In mammals, this is the cellular form thatdisseminates infection through blood. (2) Epimastigotes:also 20 µm in length with a kinetoplast anterior to thenucleus. They represent the multiplying parasite form in thetriatomid intestine. (3) Amastigotes: approximately 2 µm indiameter, rounded, with no emergent flagellum. It multiplieswithin the mammalian host cells, forming nests, until theyrupture after several cell divisions. Before their release fromthe host cells, amastigotes differentiate into trypomastigoteswhich once released, invade the blood stream; they maythen enter any other nucleated cell. Epimastigotes can begrown in axenic cultures while amastigotes are grown incultured mammalian cells, releasing trypomastigotes thatcan be harvested to perform in vitro assays.

T. cruzi displays great biological, biochemical, andgenetic diversity; therefore different strains of the parasitehave been identified and classified into six discrete typingunits (DTUs) [24, 25]. Strains of T. cruzi have been involvedin different clinical forms of Chagas disease [26, 27],thus implicating a different genetic population in tissuetropism, replication, and virulence, and in consequence indisease outcome. T. cruzi strains corresponding to differentDTUs might have relevant consequences on congenitaltransmission and fetal/neonatal pathology, even thoughVirreira et al. [28] and Burgos et al. [9] concluded thatcongenital transmission of T. cruzi is not associated withgenetic polymorphism of T. cruzi. However, Solana et al.[29] and Triquell et al. [21] described biological differencesamong subpopulations of T. cruzi in experimental verticaltransmission and placental infection.

5. Mother and Fetus/NewbornImmune Response

The immune system is fundamental to protect the motheragainst the environment, and to prevent damage to the fetus.During pregnancy the maternal immune system is charac-terized by a reinforced network of cellular and molecularrecognition, communication, trafficking and repair; it raisesthe alarm to maintain the wellbeing of the mother and thefetus. On the other hand, the fetus provides a developingactive immune system that will modify the way the motherresponds to the environment, providing a uniqueness ofthe immune system responses during pregnancy [30].

Journal of Tropical Medicine 3

A crucial factor to stop, limit, or permit the developmentof fetal/neonatal infection relates to the capacity of themother and fetus/newborn to mount innate and/or specificimmune response(s) against pathogens. Clinical studies haveshown a strong association between intrauterine infectionsand pregnancy disorders such as abortion, preterm labor,intrauterine growth retardation, and preeclampsia [31]. Asdescribed above, congenital T. cruzi infection is associatedwith some of these pathologies [5–7, 15]. Production ofproinflammatory cytokines can be observed in uninfectednewborn from infected mothers [15]. Contrarily, the levelsof inflammation markers and activation of NK cells arerather low in congenitally infected newborns [32]. This datahighly suggests a protective role of such innate defensesin an uninfected newborn from infected mothers. On theother hand, maternal T. cruzi-specific IgG antibodies playprotective roles in mothers and in fetuses when antibodiesare transferred through the placenta [33] and also maycontribute to a reduction in parasitaemia [15].

6. Placenta

The placenta is the principal site for the exchange of nutrientsand gases between the mother and fetus. This organ plays animportant role in hormone, peptide, and steroid synthesisnecessary for a successful pregnancy [34]. The humanplacenta is classified as a hemochorial villous placenta inwhich the free chorionic villi, formed by the trophoblast andthe villous stroma, are the functional units. The trophoblastcontacts maternal blood in the intervillous space, and it isseparated by a basal lamina from the villous stroma, whichis connective tissue containing the vascular endothelium,fibroblasts, and macrophages (Figure 1) [35]. Trophoblast,basal lamina, and villous stroma with the endothelium offetal capillaries form the placental barrier that must becrossed by different pathogens, including T. cruzi, to infectthe fetus during vertical transmission [16–22, 36–41].

Placentas from mother with acute Chagas disease (highparasitaemia) show severe histopathological changes, such asextensive necrosis, inflammatory infiltrate, and amastigotenests [5]. Contrarily, placentas from mother with chronicChagas disease do not present necrotic foci and inflamma-tory infiltrate. Although parasite antigens can be visualizedin the villous stroma, the typical amastigote nests are notpresent [22]. In accordance with these results, in ex vivoinfected placental explants, though parasite antigens andDNA can be detected [17, 18, 42], amastigote nests are notobserved. Only few individual parasites can be detected. Thisevidence suggests that antiparasite mechanisms may existin the placental tissue of women suffering chronic Chagasdisease.

7. Possible Antiparasitic Mechanisms ofthe Placenta

We updated the importance of the presence of the causalagent of Chagas disease in the intervillous space of humanplacentas, the viability of the parasite in this environment,

Figure 1: Electron micrograph of a chorionic villous humanplacenta. Picture depicts the intervillous space (1) and the placentalbarrier formed by the syncytiotrophoblast (2), a discontinuouscytotrophoblast (3), basal laminae (asterix), conective tissue, andfetal vessels (4).

and the process of infection of the placental tissue, mainly byex vivo and in vitro studies.

(1) Clearance of T. cruzi from the intervillous space isassociated with the risk of congenital transmission.Thus, a high parasitaemia, as in acute infection,correlates with a higher transmission rate [15, 18, 43,44]. Thus the amount of parasites could be an impor-tant risk factor for mother to fetus transmission ofT. cruzi. There are only few publications analyzingthe survival of T. cruzi in the placental environment.Triquell et al. [21], employing chorionic villi exvivo and in vitro infection model cocultured withtrypomastigotes from two different strains of T.cruzi, observed that one of the strains presents abetter survival rate than the other in the placentalenvironment. Furthermore, the two strains of T.cruzi respond differently when they were treatedwith placental media. Therefore, the great biological,biochemical, and genetic diversity of the parasite maydetermine, at least partially, the capacity of placentalinfection. These results open a new concept, thatplacenta might exert a clearance of the parasite fromthe intervillous space, and that different populationsof T. cruzi have different survival capacities in thatenvironment.

(2) Contact time between T. cruzi and the trophoblastin the intervillous space: the time that the parasiteremains in the intervillous space in contact withthe syncytiotrophoblast is poorly known. Placentalbarrier is constituted by the trophoblast tissue, thatcomprises a continuous multinucleated, nonreplicat-ing cell layer, the syncytiotrophoblast, a replicating


Cardiac output in woman 4250 mL/min

In pregnancy: 20% increase in circulating blood volume and 40% increase in cardiac output

Cardiac output in pregnant woman 5950 mL/min

10% of cardiac output reaches the uterus and 80% of this volume reaches the placenta

475 mL/min of blood reaches the placenta

Low parasitemia (0.1–1 parasite/mL): 68544–685440 parasites circulate through

the placenta in 24 hours

High parasitemia (over 40 parasite/mL): more than 27 million parasites circulate through the placenta in 24 hours

Figure 2: Estimation of T. cruzi contact with the placenta in infected mothers.

layer of cytotrophoblasts that fuses with the STB,a basal lamina, and an underlying villous stromaor connective tissue, that includes vascular endothe-lium [35]. The placental barrier must be crossedby the parasites, therefore the time that T. cruzitrypomastigotes stay in the intervillous space andinteract with the syncytiotrophoblast is of outmostimportance. Shippey et al. [45] in a dual perfusionsystem of placental cotyledons observed T. cruzi DNAin the maternal effluents at 30 min, 60 min, and90 min after injection an only one bolus of T. cruzitrypomastigotes through the maternal perfusate.There was no parasite DNA in the fetal effluent,indicating there was no passage to fetal circulationdespite the great concentration of parasites injected.However, the perfusion time was only 120 min inthese experiments. Contrarily, in the ex vivo infectionof chorionic villi explants, a reproducible infectionis obtained after 24 hours of coincubation with theparasite [17]. However, the perfusion experimentsindicate that T. cruzi is present in the intervillousspace at least for an hour and half. Despite this longtime of interaction, T. cruzi was not able to invade orsurvive in the placental barrier, indicating a defensemechanism of the placental barrier against the causalparasite of Chagas.

(3) Placental infection: the ex vivo and in vitro infectionof human chorionic villi explants from human termplacenta with the parasite is an excellent and easyway to study the mechanism of cellular and tissueinvasion mechanisms. The explants can be kept inculture for several days [46], where constituent cellsand tissues are in a more physiological conditionthan their isolated counterparts in monolayer cellculture models. Another advantage of this model isthat the cells retain physical contact with the basallamina and continue to receive paracrine growthfactor signals from the underlying villous stroma.

In our laboratories, we have established the optimalconditions for the ex vivo infection of chorionicvilli explants with T. cruzi [17, 18, 20, 21, 36–41,47]. The coincubation of 105 or 106 trypomastigotesproduces a reproducible infection of the chorionicvilli [17]. This parasite’s concentration may seem tobe extremely high, but if we consider the amountof blood that circulates through the placenta everyday, and then calculate the number of parasitesthat reaches the placenta, the parasite concentrationrecommended for ex vivo infection is not high.Therefore, if we consider that the cardiac output inwomen is 4250 mL/min, and that during pregnancythe circulating blood volume increases in 20% andthe cardiac output in 40%. Then the cardiac output inpregnant women is 5950 mL/min. From this output,10% reaches the pregnant uterus and 80% of thisvolume reaches the placenta. Taking into accountall the data, a volume of 475 mL/minute of bloodreaches the placenta [48]. Considering a parasitemiaas low as 0,1 to 1 parasite/mL, a total of 68544 to685440 parasites circulate through the placenta in24 hours (Figure 2). On the other hand, in pregnantwomen with acute Chagas disease, Torrico et al.[11] have reported parasitemias over 40 parasites/mL;therefore, in this condition a total of 27 millionparasites circulate in 24 hours in the placenta. If weconsider all these data, our experimental conditionsare not far from in vivo conditions.

The trophoblast, the first tissue that is in contact withthe parasite in the intervillous space, constitutes a potentialbarrier to T. cruzi. We observed that the most notable tissuedamage induced by T. cruzi in the chorionic villi explantsis the trophoblast detachment and destruction. Additionally,the parasite induces selective disorganization of basal lamina,collagen I destruction [17], and apoptosis (especially in thetrophoblast) in infected chorionic villi explants [49]. Inaccordance, we detected similar histopathological changes in


placentas from women with chronic Chagas disease ([22]manuscript in this number of J. of Tropical Medicine).Therefore, the similar histopathological changes observedin chagasic mothers and in ex vivo infected chorionic villi,validates the latter model. The extracellular matrix alterationproduced by T. cruzi not only promotes its motility in tissuesand its entrance into cells, but also alters the presence ofcytokines and chemokines, which in turn permits T. cruzito modulate and evade both the inflammatory and immuneresponses [16, 50, 51]. Alternatively, these changes in ECMfunction may be part of a local placental defense mechanism,which could explain why only very few parasites can bedetected in the placenta. Similar effects can be observed inthe chorionic villi explants during ex vivo infection, sinceplacental explants do not allow a sustained infection byT. cruzi [19]. Thus, the placenta controls the productiveinfection of T. cruzi in chorionic villous and exerts aprotective function to fetus.

In order to understand the mechanism by which T.cruzi fuses with trophoblast plasma membrane, Calderonand Fabro [52] studied the interaction between syncytialplasma membranes from the human placenta and from theparasites, founding modifications of membrane lipids andproteins of the syncytiotrophoblast. Additionally, modifica-tions of enzyme activities in the chorionic villi have beendescribed [37–39]. For instance, placental alkaline phos-phatase (PLAP) is a glycosylphosphatidylinositol anchoredplasma membrane protein present in the trophoblast thatdecreases its activity in chagasic women and is related tocongenital transmission [53]. In ex vivo infected chorionicvilli, pretreatment of the placental tissue with phospholipaseC prevents the parasite-induced decrease of PLAP activityand significantly reduces the infectivity of T. cruzi. Theseresults are consistent with a pathogenetic role for placentalalkaline phosphatase in congenital Chagas disease [36, 54].Additionally, T. cruzi induces in the ex vivo infection modelan increase of lysosomal vesicles in the trophoblast, which arefundamental during cell invasion of the parasite [36–39].

Analyzing the process of trophoblast infection by T.cruzi in an in vitro system culturing monolayer trophoblastscells in interaction with infective trypomastigotes, it wasshown that two types of chorionic villi trophoblasts, syn-cytiotrophoblast, and cytotrophoblast have a differentialsusceptibility to infection by the causal agent of congenitalChagas disease [55]. The reduced infection in the syncy-tiotrophoblast was associated to fewer viable parasites in theculture medium and increased levels of nitric oxide. Theseresults emphasize the importance of the integrity of the firstplacental barrier, the syncytiotrophoblast, in order to avoida T. cruzi infection of the inner trophoblasts or stromalchorionic villi cells. As it was described above, structuraltrophoblast alteration is a common sign of miscarriagesand premature births in placentas of chagasic women andstrongly associated to the congenital transmission of T.cruzi. In these clinical situations, the detachment of thefirst placental barrier is a common sign which is alsoassociated to parasitism of the placental tissue [9]. Thus,differential infection between the first placental barrier withrespect to the inner trophoblast or stromal cells could

represent a mechanism of invasion of the human placenta byT. cruzi.

8. Conclusion

Congenital Chagas transmission constitutes an increasingpublic health problem, and it is responsible for the urban-ization and spreading of the disease to nonendemic areas ofLatin America, United States of America, and Europe [18,21]. The fact that the ex vivo infection of the chorionic villiexplants with the parasite reproduces the in vivo infectionin terms of cellular changes and infectivity makes it anexcellent tool for studying parasite infection strategies as wellas possible local antiparasitic mechanisms.

Acknowledgments

This work was supported by Secretary of Science andTechnology, Universidad Nacional de La Rioja; Ministryof Science and Technology Cordoba-PID 48; Secretary ofScience and Technology-Universidad Nacional de Cordoba;PME, Secretary of Science and Technology, National Uni-versity of Cordoba [3042/09 and 3416/11] (to REF); Grants11080166, 1120230 from FONDECYT and CONICYT-PBCTAnillo ACT 112, Chile (to UK); and Grant “Programa deCooperacion Cientıfica Internacional” CONICYT/MINCYT2011-595-CH/11/08 (to REF and UK).

References

[1] J. Clayton, “Chagas disease 101,” Nature, vol. 465, no. 7301,pp. S4–S5, 2010.

[2] H. Schneider and J. Dancis, “In vitro perfusion of humanplacental tissue,” Contributions to Gynecology and Obstetrics,vol. 13, pp. 5–176, 1984.

[3] F. Kierszenbaum, “Mechanisms of pathogenesis in Chagasdisease,” Acta Parasitologica, vol. 52, no. 1, pp. 1–12, 2007.

[4] A. R. L. Teixeira, R. J. Nascimento, and N. R. Sturm, “Evolu-tion and pathology in Chagas disease: a review,” Memorias doInstituto Oswaldo Cruz, vol. 101, no. 5, pp. 463–491, 2006.

[5] A. M. Altemani, A. L. Bittencourt, and A. M. A. Lana,“Immunohistochemical characterization of the inflammatoryinfiltrate in placental Chagas’ disease: a qualitative andquantitative analysis,” American Journal of Tropical Medicineand Hygiene, vol. 62, no. 2, pp. 319–324, 2000.

[6] A. L. Bittencourt, “Congenital Chagas disease,” AmericanJournal of Diseases of Children, vol. 130, no. 1, pp. 97–103,1976.

[7] A. L. Bittencourt, “Possible risk factors for vertical trans-mission of Chagas’ disease,” Revista do Instituto de MedicinaTropical de Sao Paulo, vol. 34, no. 5, pp. 403–408, 1992.

[8] S. B. Blanco, E. L. Segura, E. N. Cura et al., “Congenitaltransmission of Trypanosoma cruzi: an operational outlinefor detecting and treating infected infants in northwesternArgentina,” Tropical Medicine and International Health, vol. 5,no. 4, pp. 293–301, 2000.

[9] J. M. Burgos, J. Altcheh, M. Bisio et al., “Direct molecularprofiling of minicircle signatures and lineages of Trypanosomacruzi bloodstream populations causing congenital Chagasdisease,” International Journal for Parasitology, vol. 37, no. 12,pp. 1319–1327, 2007.


[10] L. V. Kirchhoff, “Chagas disease (American Trypanosomiasis):a tropical disease now emerging in the United States,” inEmerging Infections, W. M. Scheld, C. William, and J. M.Hughes, Eds., vol. 3, pp. 111–134, ASM Press, Washington,DC, USA, 1999.

[11] F. Torrico, M. Castro, M. Solano et al., “Effects of maternalinfection with Trypanosoma cruzi in pregnancy developmentand in the newborn infant,” Revista da Sociedade Brasileira deMedicina Tropical, vol. 38, pp. 73–76, 2005.

[12] R. E. Gurtler, E. L. Segura, and J. E. Cohen, “Congenitaltransmission of Trypanosoma cruzi infection in Argentina,”Emerging Infectious Diseases, vol. 9, no. 1, pp. 29–32, 2003.

[13] M. I. Jercic, R. Mercado, and R. Villarroel, “CongenitalTrypanosoma cruzi infection in neonates and infants from tworegions of Chile where Chagas’ disease is endemic,” Journal ofClinical Microbiology, vol. 48, no. 10, pp. 3824–3826, 2010.

[14] World Health Organization, “Estimacion cuantitativa dela enfermedad de Chagas en las Americas,” Tech. Rep.OPS/HDM/CD/425-06, World Health Organization, Geneva,Switzerland, 2006.

[15] R. C. Lambert, K. N. Kolivras, L. M. Resler, C. C. Brewster, andS. L. Paulson, “The potential for emergence of chagas diseasein the United States,” Geospatial Health, vol. 2, no. 2, pp. 227–239, 2008.

[16] Y. Carlier and C. Truyens, “Maternal-fetal transmission ofTrypanosoma cruzi,” in American Trypanosomiasis ChagasDisease One Hundred years of Research, J. Telleria and M.Tibayrenc, Eds., pp. 539–581, Elsevier, New York, NY, USA,2010.

[17] J. Duaso, G. Rojo, G. Cabrera et al., “Trypanosoma cruziinduces tissue disorganization and destruction of chorionicvilli in an ex vivo infection model of human placenta,”Placenta, vol. 31, no. 8, pp. 705–711, 2010.

[18] U. Kemmerling, C. Bosco, and N. Galanti, “Infection andinvasion mechanisms of Trypanosoma cruzi in the congen-ital transmission of chagas’ disease: a proposal,” BiologicalResearch, vol. 43, no. 3, pp. 307–316, 2010.

[19] A. G. Schijman, “Congenital chagas disease,” in Congenitaland Other Related Infectious Diseases of the Newborn, vol. 13,pp. 223–258, Perspectives in Medical Virology, New York, NY,USA, 2006.

[20] C. D. Lujan, M. F. Triquell, A. Sembaj, C. E. Guerrero, andR. E. Fretes, “Trypanosoma cruzi: productive infection is notallowed by chorionic villous explant from normal humanplacenta in vitro,” Experimental Parasitology, vol. 108, no. 3-4, pp. 176–181, 2004.

[21] M. F. Triquell, C. Dıaz-Lujan, H. Freilij, P. Paglini, andR. E. Fretes, “Placental infection by two subpopulations ofTrypanosoma cruzi is conditioned by differential survival of theparasite in a deleterious placental medium and not by tissuereproduction,” Transactions of the Royal Society of TropicalMedicine and Hygiene, vol. 103, no. 10, pp. 1011–1018, 2009.

[22] J. Duaso, E. Yanez, C. Castillo et al., “Reorganization ofextracellular matrix in placentas from women with asymp-tomatic chagas disease: mechanism of parasite invasion orlocal placental defense?” Journal of Tropical Medicine, vol.2012, Article ID 758357, 8 pages, 2012.

[23] A. Prata, “Clinical and epidemiological aspects of Chagasdisease,” Lancet Infectious Diseases, vol. 1, no. 2, pp. 92–100,2001.

[24] J. Telleria, D. G. Biron, J. P. Brizard et al., “Phylogenetic char-acter mapping of proteomic diversity shows high correlationwith subspecific phylogenetic diversity in Trypanosoma cruzi,”

Proceedings of the National Academy of Sciences of the UnitedStates of America, vol. 107, no. 47, pp. 20411–20416, 2010.

[25] B. Zingales, S. G. Andrade, M. R. S. Briones et al., “A newconsensus for Trypanosoma cruzi intraspecific nomenclature:second revision meeting recommends TcI to TcVI,” Memoriasdo Instituto Oswaldo Cruz, vol. 104, no. 7, pp. 1051–1054,2009.

[26] W. O. Dutra, M. O. C. Rocha, and M. M. Teixeira, “Theclinical immunology of human Chagas disease,” Trends inParasitology, vol. 21, no. 12, pp. 581–587, 2005.

[27] B. Zingales, B. S. Stolf, R. P. Souto, O. Fernandes, and M.R. S. Briones, “Epidemiology, biochemistry and evolutionof Trypanosoma cruzi lineages based on ribosomal RNAsequences,” Memorias do Instituto Oswaldo Cruz, vol. 94, no.1, pp. 159–164, 1999.

[28] M. Virreira, C. Alonso-Vega, M. Solano et al., “CongenitalChagas disease in Bolivia is not associated with DNA poly-morphism of Trypanosoma cruzi,” American Journal of TropicalMedicine and Hygiene, vol. 75, no. 5, pp. 871–879, 2006.

[29] M. E. Solana, A. M. Celentano, V. Tekiel, M. Jones, and S.M. Gonzalez Cappa, “Trypanosoma cruzi: effect of parasitesubpopulation on murine pregnancy outcome,” Journal ofParasitology, vol. 88, no. 1, pp. 102–106, 2002.

[30] G. Mor and I. Cardenas, “The immune system in preg-nancy: a unique complexity,” American Journal of ReproductiveImmunology, vol. 63, no. 6, pp. 425–433, 2010.

[31] K. Koga and G. Mor, “Toll-like receptors at the maternal-fetal interface in normal pregnancy and pregnancy disorders,”American Journal of Reproductive Immunology, vol. 63, no. 6,pp. 587–600, 2010.

[32] E. Hermann, A. Berthe, C. Truyens et al., “Killer cellimmunoglobulin-like receptor expression induction onneonatal CD8+ T cells in vitro and following congenitalinfection with Trypanosoma cruzi,” Immunology, vol. 129, no.3, pp. 418–426, 2010.

[33] S. F. Breniere, M. Bailly, R. Carrasco, and Y. Carlier, “Transmis-sion transplacentaire des anticorps anti-Trypanosoma cruzi,”Cahiers ORSTOM. Serie Entomologie Medicale et Parasitologie,vol. 21, no. 3, pp. 139–140, 1983.

[34] K. L. Moore and T. V. N. Perseaud, The Developing Human,Clinically Oriented Embryology, Elsevier, New York, NY, USA,7th edition, 2004.

[35] K. Berniscke, P. Kaufmann, and R. N. Baergen, Pathology of theHuman Placenta, Springer, New York, NY, USA, 2006.

[36] R. E. Fretes and S. P. de Fabro, “Trypanosoma cruzi: modifica-tion of alkaline phosphatase activity induced by trypomastig-otes in cultured human placental villi,” Revista do Instituto deMedicina Tropical de Sao Paulo, vol. 32, no. 6, pp. 403–408,1990.

[37] R. E. Fretes and S. P. de Fabro, “Human chagasic placentallocalization of enzymes associated to syncytiotrophoblastmembrane,” Comunicaciones Biologicas, vol. 9, pp. 51–59,1990.

[38] R. E. Fretes and S. P. de Fabro, “Increase of Hofbauer cells inhuman placentas cocultured in vitro with Trypanosoma cruzi,”Revista de la Facultad de Ciencias Medicas, vol. 52, no. 1, pp.7–12, 1994.

[39] R. E. Fretes and S. P. de Fabro, “In vivo and in vitro analysisof lysosomes and acid phosphatase activity in human chagasicplacentas,” Experimental and Molecular Pathology, vol. 63, no.3, pp. 153–160, 1995.

[40] L. Mezzano, G. Repossi, R. E. Fretes, S. Lin, M. J. Sartori, andS. P. de Fabro, “In vitro infection of Trypanosoma cruzi causesdecrease in glucose transporter protein-1 (GLUT1) expression


in explants of human Placental Villi cultured under normaland high glucose concentrations,” Journal of Tropical Medicine,vol. 2012, Article ID 969243, 8 pages, 2012.

[41] L. Mezzano, M. J. Sartori, S. Lin, G. Repossi, and S. P.de Fabro, “Placental alkaline phosphatase (PLAP) study indiabetic human placental villi infected with Trypanosomacruzi,” Placenta, vol. 26, no. 1, pp. 85–92, 2005.

[42] A. Al-Khan, I. L. Aye, I. Barsoum et al., “IFPA meeting2010 workshops report II: placental pathology; trophoblastinvasion; fetal sex; parasites and the placenta; decidua andembryonic or fetal loss; trophoblast differentiation and syncy-tialisation,” Placenta, vol. 32, supplement 2, pp. S90–S99, 2011.

[43] E. Moretti, B. Basso, I. Castro et al., “Chagas’ disease: study ofcongenital transmission in cases of acute maternal infection,”Revista da Sociedade Brasileira de Medicina Tropical, vol. 38,no. 1, pp. 53–55, 2005.

[44] H. Freilij and J. Altcheh, “Congenital Chagas’ disease: diagnos-tic and clinical aspects,” Clinical Infectious Diseases, vol. 21, no.3, pp. 551–555, 1995.

[45] S. H. Shippey, C. M. Zahn, M. M. Cisar, T. J. Wu, and A.J. Satin, “Use of the placental perfusion model to evaluatetransplacental passage of Trypanosoma cruzi,” American Jour-nal of Obstetrics and Gynecology, vol. 192, no. 2, pp. 586–591,2005.

[46] S. Drewlo, D. Baczyk, C. Dunk, and J. Kingdom, “Fusion assaysand models for the trophoblast,” Methods in Molecular Biology,vol. 475, pp. 363–382, 2008.

[47] F. Frank, M. J. Sartori, C. Asteggiano, S. Lin, S. P. De Fabro,and R. E. Fretes, “The effect of placental subfractions onTrypanosoma cruzi,” Experimental and Molecular Pathology,vol. 69, no. 2, pp. 144–151, 2000.

[48] P. Libby, R. O. Bonow, D. L. Mann, and D. P. Zipes, Braun-wald’s Heart Disease: A Textbook of Cardiovascular Medicine,Elsevier, New York, NY, USA, 8th edition, 2008.

[49] J. Duaso, G. Rojo, F. Jana et al., “Trypanosoma cruzi inducesapoptosis in ex vivo infected human chorionic villi,” Placenta,vol. 32, no. 5, pp. 356–361, 2011.

[50] T. C. Cardenas, C. A. Johnson, S. Pratap et al., “Regulation ofthe extracellular matrix interactome by Trypanosoma cruzi,”Open Parasitology Journal, vol. 4, pp. 72–76, 2010.

[51] A. P. M. P. Marino, A. A. Silva, R. T. Pinho, and J. Lannes-Vieira, “Trypanosoma cruzi infection: a continuous invader-host cell cross talk with participation of extracellular matrixand adhesion and chemoattractant molecules,” BrazilianJournal of Medical and Biological Research, vol. 36, no. 8, pp.1121–1133, 2003.

[52] R. O. Calderon and S. P. de Fabro, “Trypanosoma cruzi:fusogenic ability of membranes from cultured epimastigotesin interaction with human syncytiotrophoblast,” ExperimentalParasitology, vol. 56, no. 2, pp. 169–179, 1983.

[53] M. J. Sartori, S. Lin, R. E. Fretes, L. Ruiz Moreno, L.Goldemberg, and S. P. de Fabro, “Alkaline phosphatase activityin plasma of pregnant chagasic patients,” Revista de la Facultadde Ciencias Medicas, vol. 55, no. 1-2, pp. 5–8, 1997.

[54] M. J. Sartori, S. Lin, F. M. Frank, E. L. Malchiodi, andS. P. De Fabro, “Role of placental alkaline phosphatase inthe interaction between human placental trophoblast andTrypanosoma cruzi,” Experimental and Molecular Pathology,vol. 72, no. 1, pp. 84–90, 2002.

[55] C. Diaz-Lujan, M. F. Triquell, A. Schijman, P. Paglini, andR. E. Fretes, “Differential susceptibility of isolated humantrophoblasts to infection by Trypanosoma cruzi,” Placenta, vol.33, no. 4, pp. 264–270, 2012.


Research Article

Transplacental Transmission of Plasmodium falciparum ina Highly Malaria Endemic Area of Burkina Faso

Alphonse Ouedraogo,1 Alfred B. Tiono,1 Amidou Diarra,1 Edith C. Christiane Bougouma,1

Issa Nebie,1 Amadou T. Konate,1 and Sodiomon B. Sirima1, 2

1 Centre National de Recherche et de Formation sur le Paludisme, 1487 Avenue de l’Oubritenga, BP 2208,Ouagadougou 01, Burkina Faso

2 Groupe de Recherche Action en Sante, BP 10248, Secteur 25, Somgande, Rue 25.26, Porte 79, Ouagadougou 06, Burkina Faso

Correspondence should be addressed to Sodiomon B. Sirima, [email protected]

Received 1 June 2011; Revised 3 October 2011; Accepted 17 October 2011

Academic Editor: Ricardo E. Fretes

Copyright © 2012 Alphonse Ouedraogo et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

Malaria congenital infection constitutes a major risk in malaria endemic areas. In this study, we report the prevalence oftransplacental malaria in Burkina Faso. In labour and delivery units, thick and thin blood films were made from maternal,placental, and umbilical cord blood to determine malaria infection. A total of 1,309 mother/baby pairs were recruited. Eighteencord blood samples (1.4%) contained malaria parasites (Plasmodium falciparum). Out of the 369 (28.2%) women with peripheralpositive parasitemia, 211 (57.2%) had placental malaria and 14 (3.8%) had malaria parasites in their umbilical cord blood. Theumbilical cord parasitemia levels were statistically associated with the presence of maternal peripheral parasitemia (OR = 9.24,P � 0.001), placental parasitemia (OR = 10.74, P � 0.001), high-density peripheral parasitemia (OR = 9.62, P � 0.001), andhigh-density placental parasitemia (OR = 4.91, P = 0.03). In Burkina Faso, the mother-to-child transmission rate of malariaappears to be low.

1. Introduction

In malaria high-transmission areas, some population groupsare at considerably higher risk of infection with Plasmodiumfalciparum and development of malaria morbidity or mortal-ity than others. These include children less than five years ofage and pregnant women. Malaria contributes significantlyto perinatal disease burden in terms of pregnancy loss,prematurity due to preterm labor, and intrauterine growthretardation [1]. Malaria infection during pregnancy poses asubstantial risk to the mother, her fetus, and the neonate. Inareas of stable malaria transmission such as Burkina Faso,where adult women have considerable acquired immunity,Plasmodium falciparum infection during pregnancy doesnot cause symptomatic malaria, but may lead to maternalanemia as well as placental and cord blood malaria infection,especially among primigravidae and secundigravida [2–4].

Placental malaria is defined as the accumulation ofPlasmodium-infected erythrocytes in the intervillous space in

the placenta, causing histologic changes including leukocyte-induced damage to the trophoblastic basement membrane.The placental infection does not reflect the existence ofperipheral infection over a short period preceding the deliv-ery or whether it is related to infection during pregnancy.Susceptibility to this may be correlated to high exposureto malaria and repeated episodes of parasitemia during thepregnancy [5].

Vertical transmission of malaria from mother to foetusthrough the placenta and umbilical cord is defined as umbil-ical cord blood parasitemia. The transplacental transmissionof Plasmodium falciparum from mother to fetus has longbeen well-described [6, 7].

The direct burden of neonatal malaria infection in termsof prevalence is not well-described in malaria endemic areas.In fact, the method used to identify congenital transmissionis peripheral blood of newborns or umbilical cord blood.Malaria parasites have been detected only rarely in theperipheral blood of newborns, whether the blood specimen


is collected at the time of birth or hours later [8, 9]. In studiesin which both umbilical cord blood and infant peripheralblood were obtained at the time of birth, the parasite loadin the babies’ peripheral blood has always been lower thanthat in the umbilical cord blood [9–11].

Studies published so far have documented contradictorylevels of this burden. In countries without endemic malaria,congenital malaria has occurred in children born to womenwho have immigrated from malarious areas. However,transplacental transmission of Plasmodium falciparum hasbeen found to be rare in malaria-endemic areas, rangingfrom about 1 to 5% [12–15]. In contrast, data from recentstudies on the burden of congenital and neonatal malaria,while scarce and contradictory, have indicated a high burden(more than 15%) in parts of sub-Saharan Africa [13, 15–18].

In Burkina Faso, the real prevalence of neonatal malaria isunknown but is estimated to be even higher. This assessmentis based on presumptive malaria diagnosis. The present studywas designed to determine the real burden of transplacentaltransmission, the risk factors associated with transplacentaltransmission, and the prevalence of cord blood and placentalmalaria parasitemia in malaria holoendemic area of BurkinaFaso.

These results represent a pooled analysis of studies onmalaria prevention in pregnant women [4, 19, 20] examiningumbilical cord blood to determine the frequency of transpla-cental transmission of Plasmodium falciparum.

2. Materials and Methods

2.1. Study Site. The first study took place within six deliveryunits (DUs) of the Koupela health district, which is locatedaround 120 km east of Ouagadougou. The second study wasconducted in one DU of the Health District of Bousse. Bothstudy sites have been extensively described elsewhere [4, 19,20]. Malaria transmission is stable, with marked seasonalityin both sites. Transmission is intense during the rainy season(June to October). The main malaria vectors are Anophelesgambiae, Anopheles arabiensis, and Anopheles funestus. Theannual entomological inoculation rates range from 10 to500 infective bites per individual. Plasmodium falciparum isresponsible for more than 90% of malaria infections.

2.2. Study Population. The study participants were pregnantwomen who voluntarily consented to participate in trials ofmalaria prophylaxis during pregnancy. They were encour-aged to deliver at the health facility where the study sampleswere collected for processing.

2.3. Clinical Procedures. Women delivering at the healthfacility, after giving informed consent, were asked a standardseries of questions focused on sociodemographic charac-teristics, history of fever, antimalarial drug use, and theuse of antimalarial chemoprophylaxis and bednets. Capillaryblood was obtained by fingerstick for malaria blood filmpreparation. Placental blood films were prepared by identi-fying the maternal side of the placenta, wiping away excessblood, cutting into the surface, and placing pooled blood

onto a slide. Umbilical cord blood samples were obtained bywiping away excess blood from a clamped cord, piercing itwith a lancet, and placing a drop of expressed blood on aslide.

A detailed clinical examination was done on each new-born infant within 24 hours of delivery. Neonates wereweighted with an electronic digital scale (±10 grams) (TanitaCorporation, Tokyo, Japan). The Dubowitz scoring systemwas used to estimate gestational age, using findings fromphysical and neurologic examinations. Scoring by APGARindex was performed at delivery but was not recorded in thisstudy.

2.4. Lab Methods. All blood films (maternal, placental,and cord) were stained with Giemsa and examined forparasites at the “Centre National de Recherche et de For-mation sur le Paludisme” immunoparasitology laboratoryin Ouagadougou. For thick films, parasites and leukocyteswere counted in the same fields until 500 leukocytes werecounted. Parasite densities were estimated using an assumedleukocyte count of 8,000 leukocytes/µL. Parasite densitieswere calculated according to the following formula: numberof parasites counted × 8000/number of leucocytes counted.Thin films were then used to determine species when thickfilms were positive. All slides were double-read by twoindependent microscopists. If the ratio of densities from thefirst two readings was greater than 1.5 or less than 0.67, orif fewer than 30 parasites were counted with a differenceof more than 10 parasites between the two readings, theslide was evaluated a third time. The geometric mean ofthe parasite density of the two most concordant results ofthe three readings was taken as the final result. When thediscordance was only in terms of positivity, the slide was alsoevaluated a third time and the definitive result was based onthe majority verdict for positivity.

2.5. Definitions. Blood film results were considered to bepositive if any asexual-stage parasites were identified andnegative if no parasites were seen in 100 high-power fields.

Prematurity was defined as a gestational age <37 weeks asestimated by Dubowitz examination.

We defined low birth weight as a birth weight of <2,500 gwith a gestational age ≥37 weeks.

2.6. Statistical Analysis. Data collected in the study ques-tionnaire were verified, then double-entered, and validatedwith EpiInfo, version 6.04 fr (Centers for Diseases Controland Prevention, Atlanta, USA). Data were analyzed usingEpi-info and Stata 7.0. The analysis included data frombirths of all enrolled participants who delivered at the studycentre clinic for whom data were available. Continuous,normally distributed data were described by mean andstandard deviation and nonnormally distributed data by themedian or geometrical mean and range. Proportions werecompared using the Chi-square test and normally distributedcontinuous variables were compared using Student’s t-test.Statistical results were considered significant, when the two-sided P value was <0.05.


Table 1: Characteristics of enrolled women.

Gravidity

Characteristics 1 2 >2 All

Age, median (years) 19.24± 2.25 21.68± 2.60 28.71± 5.38 23.90± 5.89

Mean gestational age (weeks) 38.16± 4.24 38.79± 3.79 39.02± 3.59 38.66± 3.89

Mossi ethnic group (%) 95.8 96.2 94.7 95.5

Sleeps under bednets (%) 22.8 28.1 19.4 22.4

Mother death (N) 2 0 0 0

Pregnancy outcome

Stillbirth (N) 19 8 23 50

Miscarriage (N) 8 3 7 18

Weight (g) 2, 719± 430 2, 893± 390 2, 995± 440 2, 875± 450

Number of live babies (N) 460 270 579 1309

LBW∗ (N/%) 111 (24.1%) 39 (14.4%) 57 (9.8%) 207 (15.8%)

LBW∗: low birth weight.

2.7. Ethical Considerations. The study was discussed withhealth authorities and local leaders to obtain their assent.The study was reviewed and approved by the Burkina FasoMinistry of Health ethical committee. Informed consent (bysignature or thumbprint) was obtained after the consentdocument was read to the women in the local language. Forilliterate mothers, the informed consent discussion processwas witnessed by an impartial individual.

3. Results

3.1. Characteristics of Women at the Delivery Unit. In total,1,374 women delivered at the health unit; 1,309 womendelivered singleton infants and provided data in the formof peripheral blood smear, placental blood, and umbilicalcord blood. The profile of enrolled women is summarized inTable 1. Most of the women spoke Moore, a language indige-nous to this region of Burkina Faso. The mean age (SD) ofthe women was 23.90 ± 5.89 years. The self-report indicatedthat ownership and use of insecticide-impregnated materials(bed nets and/or curtains) was reported by approximatelyone-third of the women. The rate of use of insecticide-impregnated materials (bed nets and/or curtains) was verylow in the study population. All of the women used drugprophylaxis during their pregnancy (IPTp/SP or weeklyprophylaxis with chloroquine).

Primigravidae comprised 35% of the total sample size,and primigravidae and secundigravida together made up56% of the total sample size. The mean duration of gestationwas 38.66 ± 3.89 weeks. The mode of delivery of babies wasspontaneous for 97.5% of the mothers.

Two women died during delivery as a consequence ofeclampsia. Other complications during the delivery includedperipartum haemorrhage, placental retention, and retropla-cental haemorrhage. Fifty stillbirths (3.6%) and 18 (1.3%)miscarriages occurred at the DUs.

The weight range of babies was 700 g to 4,700 g with amean weight of 2, 875 ± 450 g. Sixteen percent of the babiespresented low birth weight at delivery (Table 1).

3.2. Maternal Peripheral Parasitemia: Placental and UmbilicalCord Blood Parasitemia. Over one-third of the women wereparasitaemic. Indeed, of the 1,309 pregnant women, Plas-modium falciparum was detected in peripheral blood from369 (28.2%) of the women at the time of delivery. Overall, thegeometric mean of the maternal peripheral parasite densitywas 1,106.47 (95% CI 871.58–1404.67) parasites/µL. Table 2shows the prevalence of peripheral, placental, and umbilicalcord parasitemia. Plasmodium falciparum was detected in theplacentas of 255 (19.5%) women; 28.5% of these belonged toprimigravidae and 11.4% to multigravidae. The difference ininfection between the primigravidae and the multigravidaewas statistically significant (P < 0.0001). All of the womenreceived malaria prophylaxis during pregnancy.

Umbilical cord blood parasitemia with Plasmodium falci-parum was detected in 18 (1.4%) babies, and the geometricmean umbilical cord blood parasitemia was 315.69 (95% CI87.02–1145.20) parasites/µL. Of these, 2.4% (11) belongedto primigravidae and 0.5% (3) to multigravidae. Of the18 babies with Plasmodium-falciparum-positive umbilicalcord blood parasitemia, 17 were positive for trophozoitesand 1 was positive for schizonts only. Both schizonts andtrophozoites were present in seven babies, and one newbornhad a triple-positive association (trophozoite + schizont +gametocyte). Five newborns (27.8%) out of the 18 infectedwith Plasmodium falciparum had low birth weight.

Out of 369 (28.2%) women with peripheral positive par-asitemia, 211 (57.2%) had positive placental malaria and 14(3.8%) had malaria parasites in the umbilical cord blood.Eleven (5.2%) of the 211 (57.2%) women with positive pla-cental smears for malaria had malaria parasites in the umbil-ical cord blood. Of the 18 babies infected with Plasmodiumfalciparum, 14 (77.8%) were born from mothers with


Table 2: The proportion of newborns with Plasmodium falciparum parasitemia in umbilical cord blood, placental parasitemia, and maternalparasitemia, along with parasite density.

Gravidity

1 2 >2 All

Peripheral parasitemia (%) 34.3 35.2 20.3 28.2

Geometric mean (parasites/µL) (CI 95%)2,143.18

(1,552.86–2,957.92)894.71

(548.60–1,459.19)495.09

(318.44–769.74)1,106.47

(871.58–1404.67)

Negative peripheral parasitemia (%) 32.4 18.8 48.8 71.7

Peripheral parasitemia 1–999 (%) 30.9 29.5 39.6 11.4

Peripheral parasitemia 1,000–4,999 (%) 53.3 19.6 27.2 7.1

Peripheral parasitemia 5,000–9,999 (%) 52.5 22.5 25.0 3.1

Peripheral parasitemia ≥10,000 (%) 58.8 25.0 16.2 5.2

Placental parasitemia (%) 28.5 21.9 11.4 19.5

Geometric mean (parasites/µL) (CI 95%)1384.55

(880.97–2,176.00)396

(223.30–703.17)591.71

(289.60–1,209.00)830.52

(599.66–1,150.27)

Negative placental parasitemia (%) 30.9 20.0 49.1 19.9

Placental parasitemia 1–999 (%) 46.5 26.4 27.1 10.1

Placental parasitemia 1,000–4,999 (%) 51.2 26.8 22.0 3.2

Placental parasitemia 5,000–9,999 (%) 50.0 29.2 20.8 1.9

Placental parasitemia ≥10,000 (%) 69.7 6.1 24.2 2.6

Umbilical cord parasitemia (%) 2.4 1.5 0.5 1.4

Geometric mean (parasites/µL) (CI 95%)263.74

(40.85–1,702.50)644.08

(5.85–70,803.72)222.16

(0.76–64,635.57)315.69

(87.02–1,145.20)

Negative umbilical cord parasitemia (%) 34.9 20.5 44.7 98.6

Umbilical cord parasitemia 1–999 (%) 61.5 15.4 23.1 1.0

Umbilical cord parasitemia 1,000–4,999 (%) 0.0 100 0.0 0.1

Umbilical cord parasitemia 5,000–9,999 (%) 0.0 0.0 0.0 0.0

Umbilical cord parasitemia ≥10,000 (%) 66.7 33.3 0.0 0.2

positive peripheral parasitemia. For two newborns withPlasmodium falciparum infection, no peripheral or placentalinfection was found in the mother. None of the babies withcord blood positive for malaria had fever within the first 24 hof life.

The number of pregnant women with peripheral, placen-tal, and umbilical infection and high peripheral parasitemia(>10,000 parasites/µL) decreased with increasing gravidity(Table 2).

The combination of being born to a mother withboth peripheral and placental infection conferred a greaterlikelihood of infection of the baby (61%). Being born withonly maternal peripheral infection reduced the likelihoodof having umbilical cord parasitemia (17%). There wasan association between umbilical cord blood infection andmaternal peripheral infection (P < 0.001). Being born withonly placental infection reduced the rate of umbilical cordparasitemia (11%). There was also an association betweenumbilical cord blood infection and placental infection(P < 0.001) (Table 3).

Being born to a mother with maternal peripheral parasitedensity ≥5,000 parasites/µL or placental parasite density

≥10,000 parasites/µL conferred a greater likelihood of umbil-ical cord parasitemia (Table 3).

3.3. Risk Factors. We further examined the correlationbetween risk factors and the presence or absence of umbilicalcord parasitemia to determine possible factors that mightaffect infection with malaria parasites in umbilical cordblood. Univariate logistic regression analysis indicated thatbeing born with maternal parasitemia, being born withmaternal parasite density ≥ 5,000 parasites/µL, being bornwith placental parasitemia, being born with placental par-asite density between 1,000–4,999 parasites/µL, being bornwith placental parasite density ≥10,000 parasites/µL, andbeing born in a first pregnancy were all associated withthe presence of umbilical cord blood parasitemia. Lowbirth weight was not associated with umbilical cord bloodparasitemia (Table 3).

Low maternal peripheral and placental parasitemia,prematurity, use of impregnated bednets, low birth weight,and female sex were independent risk factors for infection inthe babies.


Table 3: Univariate logistic regression of risk factors associated with Plasmodium falciparum parasitemia in umbilical cord blood.

Proportion of newborns withumbilical cord blood parasitemia

and the risk factor specifiedOdds ratio 95% confidence intervals P

Peripheral positive parasitemia 77.8% 9.24 3.02–28.28 <0.001

Peripheral parasitemia 1–999 22.2% 2.23 0.75–7.20 0.13

Peripheral parasitemia 1,000–4,999 11.1% 1.64 0.37–7.28 0.50

Peripheral parasitemia 5,000–9,999 22.2% 9.62 3.01–30.69 <0.001

Peripheral parasitemia ≥10,000 22.2% 5.29 1.69–16.52 0.004

Placental positive parasitemia 72.2% 10.74 3.79–30.42 <0.001

Placental parasitemia 1–999 16.7% 1.78 0.50–6.24 0.36

Placental parasitemia 1,000–4,999 22.2% 9.62 3.01–30.69 <0.001

Placental parasitemia 5,000–9,999 5.6% 3.13 0.40–24.56 0.27

Placental parasitemia ≥10,000 11.1% 4.91 1.08–22.28 0.03

Primigravid 61.1% 2.96 1.13–7.69 0.02

Prematurity (<37 weeks) 33.3% 1.58 0.52–4.25 0.36

ITN 33.3% 1.69 0.63–4.56 0.29

LBW∗ 27.8% 2.10 0.74–5.97 0.16

Female sex of infant 61.1% 1.78 0.68–4.63 0.33

LBW∗: low birth weight.

Table 4: Multivariate logistic regression of risk factors, associated with Plasmodium falciparum parasitemia in umbilical cord blood.

Odds ratio 95% confidence intervals P

Peripheral positive parasitemia 2.43 0.50–11.66 0.26

Peripheral parasitemia 5,000–9,999 3.71 0.92–14.98 0.06

Peripheral parasitemia ≥10,000 1.60 0.38–6.66 0.51

Placental positive parasitemia 3.30 0.73–14.87 0.11

Placental parasitemia 1,000–4,999 2.27 0.58–8.77 0.23

Placental parasitemia ≥10,000 0.95 0.15–5.69 0.95

Primigravid 1.91 0.70–5.21 0.20

On multivariate logistic regression, none of the sevenfactors that were significant on univariate logistic regressionremained significant (Table 4).

4. Discussion

4.1. Low Levels of Umbilical Cord Parasitemia. This studyshows the burden of malaria in pregnant women at thedelivery unit in a malaria-endemic area of Burkina Faso. Thisinvestigation demonstrated the burden of malaria duringpregnancy. The maternal peripheral parasitemia rate foundin this study was 28.2%. This is similar to the rate reportedin other countries in sub-Saharan Africa [2, 21].

The prevalence of placental parasites was 19.5%. Previ-ously reported prevalence rates of placental parasitemia inAfrica have been highly variable, ranging from 17.2% to 57%[11, 22–24]. The placenta is a site for Plasmodium falciparumsequestration. Many hypotheses, based on a systemic orlocal failure of the immunological response to malaria, havebeen proposed to explain the preference of the parasites forreplication in the placenta [25].

The malaria rate in umbilical cord blood was low in thestudy population, occurring in 1.4% of all newborns, in 5.2%of babies born from mothers with peripheral and placentalmalaria infection, and in 5.1% of babies born from womenwith placental malaria infection only. Previously reportedrates from different parts of Africa are highly variable,ranging from 0% to 54% [11, 22, 24, 26, 27]. The low ratein the present study, which is comparable to rates reported insome previous manuscripts, could be explained by effectiveprevention of malaria during pregnancy. In fact, it has nowbeen proven that malaria parasites identified in cord bloodare acquired antenatally by transplacental transmission ofinfected erythrocytes. The mechanisms underlying congen-ital transmission, according to an earlier report, includematernal transfusion into the fetal circulation either at thetime of delivery or during pregnancy, through direct pen-etration through the chorionic villi, or through prematureseparation of the placenta [28, 29]. Pregnant women, whohave received antimalarial treatment, should be able toclear parasitemia to avoid umbilical cord transmission.Immunity in the mother could also provide an explanation.Transplacental transmission of malaria appears to occur


infrequently; it may be possible that after transplacentaltransmission, some elements of immunity, acquired fromthe mother, protect the infants from becoming infected[24]. This resistance may reflect, among other things, thephysical barrier of the placenta to infected red cells, thepassive transfer of maternal antibodies, and/or the poorenvironment afforded by fetal erythrocytes for plasmodialreplication, due to their fetal hemoglobin composition andlow free-oxygen tension [28].

Some studies have found high rates of transplacentalmalaria infection. The difference between those studies andour report could be explained by several factors. First, thehigh efficacy of malaria prophylaxis during this study mayhave allowed the women to clear their parasitaemia andtherefore prevent placental and transplacental transmissionof the infection to their babies. Second, our sole use of mi-croscopy may have led to underdiagnosis, since this methodhas lower sensitivity than the newer molecular methods, suchas real-time PCR, used in other studies [22, 29]. Finally, itmight be that our numbers reflect the true prevalence ofinfection, as compared with data collected during routineservices, in which lack of quality control of the reading ofsmears could lead to an overestimation of the infection rate.

4.2. Risk Factors. Univariate analysis of the association ofrisk factors with the presence or absence of umbilical cordparasitemia revealed seven risk factors that was associatedwith Plasmodium falciparum parasitemia in umbilical cordblood. The presence of maternal peripheral parasitemia,placental parasitemia, high-density peripheral parasitemia,high-density placental parasitemia, and primigravid statuswas associated with an increased risk of malaria parasitemiain cord blood. However, in multivariate analysis, theseassociations failed to reach statistical significance.

Maternal peripheral blood parasitemia was associatedwith umbilical cord blood parasitemia (OR = 9.24, P <0.001). Four cases of umbilical cord blood parasitemia oc-curred in pregnancies not complicated by maternal periph-eral malaria infection.

There were also associations between placental and cordblood parasitemia (OR = 10.74, P < 0.001), and betweencord blood/placental and maternal peripheral parasitemia.Five cases of umbilical cord blood parasitemia occurred inpregnancies not complicated by placental malaria infection.Only two newborns had mothers with neither peripheralmalaria infection nor placental malaria infection. Earlierstudies of transplacental malaria reported these associations[11, 18, 24, 30]. A strong association between placentalmalaria and umbilical cord blood parasitemia has beenreported, and this was suggested to be responsible forcongenital malaria. However, whether the presence of Plas-modium falciparum malaria parasites in umbilical cord blooddenotes infection acquired antenatally or contaminationwith infected maternal blood at delivery is not clear [15].These associations suggest that clearing maternal and placen-tal parasitemia with effective antimalarial drugs before deliv-ery could prevent transplacental transmission of parasitemia.

The density of maternal peripheral parasitemia (>5,000parasites/µL) was an important determinant of the likelihood

of umbilical cord blood parasitemia (OR = 5.29, P = 0.004).The density of placental parasitemia (>10,000 parasites/µL)was also an important determinant of the likelihood ofumbilical cord parasitemia (OR = 4.91, P = 0.03).High density parasitemia seems to facilitate umbilical cordinfection. Conversely, previous reports, showing low ratesof transplacental transmission of malaria, suggest that theplacental barrier is very effective in the case of very lowmalaria parasite density [22].

Finally, univariate analysis showed that being born froma first pregnancy was also linked to umbilical cord bloodparasitemia (OR = 2.96, P = 0.02).

5. Conclusions

Our data indicate that the rate of mother-to-child trans-mission of malaria, defined as positive umbilical cord bloodparasitemia, appears to be low. Clinicians are thereforeadvised to investigate other aetiologies of fever in neonates.The low level of parasitaemia in cord blood suggests thatcontamination probably occurs during the labour period.Maternal, placental, and high density parasitemia were allassociated with umbilical cord parasitemia. Prevention ofmalaria during pregnancy with effective antimalarial drugsshould reduce the risk of infection for newborns.

Conflict of Interests

The authors declare that they have no conflict of interests.

Acknowledgments

The authors express our gratitude to the pregnant womenwho participated in the study for their kind cooperation.Special thanks are due to all the staff of the health districtsof Bousse and Koupela. The authors also thank the lab staffof the Centre National de Recherche et de Formation sur laPaludisme, Mr. Ouedraogo Z. Amidou for data managementand Convelbo Nathalie for organizing the logistics of thestudy (Ouagadougou, Burkina Faso).

References

[1] J. M. McDermott, J. J. Wirima, R. W. Steketee, J. G. Breman,and D. L. Heymann, “The effect of placental malaria infectionon perinatal mortality in rural Malawi,” American Journal ofTropical Medicine and Hygiene, vol. 55, no. 1, supplement, pp.61–65, 1996.

[2] S. B. Sirima, A. H. Cotte, A. Konate et al., “Malaria preventionduring pregnancy: assessing the disease burden one year afterimplementing a program of intermittent preventive treatmentin Koupela District, Burkina Faso,” American Journal ofTropical Medicine and Hygiene, vol. 75, no. 2, pp. 205–211,2006.

[3] R. W. Steketee, B. L. Nahlen, M. E. Parise, and C. Menendez,“The burden of malaria in pregnancy in malaria-endemicareas,” American Journal of Tropical Medicine and Hygiene, vol.64, no. 1-2, supplement, pp. 28–35, 2001.


[4] A. B. Tiono, A. Ouedraogo, E. C. Bougouma et al., “Placentalmalaria and low birth weight in pregnant women living in arural area of Burkina Faso following the use of three preventivetreatment regimens,” Malaria Journal, vol. 8, p. 224, 2009.

[5] C. J. Uneke, “Impact of placental Plasmodium falciparummalaria on pregnancy and perinatal outcome in sub-SaharanAfrica: I: introduction to placental malaria,” Yale Journal ofBiology and Medicine, vol. 80, no. 2, pp. 39–50, 2007.

[6] J. Schwetz and M. Peel, “Congenital malaria and placentalinfections amongst the negroes of central Africa,” Transactionsof the Royal Society of Tropical Medicine and Hygiene, vol. 28,no. 2, pp. 167–174, 1934.

[7] WHO, The African Malaria Report, World Health Organiza-tion, Geneva, Switzerland, 2003.

[8] L. J. Bruce-Chwatt, “Malaria in African infants and children inSouthern Nigeria,” Annals of Tropical Medicine and Parasitol-ogy, vol. 46, no. 2, pp. 173–200, 1952.

[9] D. E. Marshall, “The transplacental passage of malaria para-sites in the Solomon Islands,” Transactions of the Royal Societyof Tropical Medicine and Hygiene, vol. 77, no. 4, pp. 470–473,1983.

[10] P. J. Lehner and C. J. Andrews, “Congenital malaria in PapuaNew Guinea,” Transactions of the Royal Society of TropicalMedicine and Hygiene, vol. 82, no. 6, pp. 822–826, 1988.

[11] P. O. Obiajunwa, J. A. Owa, and O. O. Adeodu, “Prevalenceof congenital malaria in Ile-Ife, Nigeria,” Journal of TropicalPediatrics, vol. 51, no. 4, pp. 219–222, 2005.

[12] F. E. Lesi, M. Y. Mukhtar, E. U. Iroha, and M. T. Egri-Okwaji, “Clinical presentation of congenital malaria at thelagos university teaching hospital,” Nigerian Journal of ClinicalPractice, vol. 13, no. 2, pp. 134–138, 2010.

[13] M. K. Mwaniki, A. W. Talbert, F. N. Mturi et al., “Congenitaland neonatal malaria in a rural Kenyan district hospital: aneight-year analysis,” Malaria Journal, vol. 9, no. 1, article 313,2010.

[14] T. C. Quinn, R. F. Jacobs, and G. J. Mertz, “Congenital malaria:a report of four cases and a review,” Journal of Pediatrics, vol.101, no. 2, pp. 229–232, 1982.

[15] C. J. Uneke, “Congenital Plasmodium falciparum malaria insub-Saharan Africa: a rarity or frequent occurrence?” Parasi-tology Research, vol. 101, no. 4, pp. 835–842, 2007.

[16] J. A. Akindele, A. Sowunmi, and A. E. Abohweyere, “Con-genital malaria in a hyperendemic area: a preliminary study,”Annals of Tropical Paediatrics, vol. 13, no. 3, pp. 273–276, 1993.

[17] P. R. Fischer, “Congenital malaria: an African survey,” ClinicalPediatrics, vol. 36, no. 7, pp. 411–413, 1997.

[18] M. Y. Mukhtar, F. E. Lesi, E. U. Iroha, M. T. Egri-Okwaji, andA. G. Mafe, “Congenital malaria among inborn babies at atertiary centre in Lagos, Nigeria,” Journal of Tropical Pediatrics,vol. 52, no. 1, pp. 19–23, 2006.

[19] A. Ouedraogo, E. C. Bougouma, A. Diarra et al., “Impact com-paratif de trois schemas de prevention du paludisme pendantla grossesse sur l’anemie maternelle, associee a l’infectionpalustre au Burkina Faso,” Medecine et Maladies Infectieuses,vol. 38, no. 4, pp. 180–186, 2008.

[20] S. B. Sirima, R. Sawadogo, A. C. Moran et al., “Failure ofa chloroquine chemoprophylaxis program to adequately pre-vent malaria during pregnancy in Koupela District, BurkinaFaso,” Clinical Infectious Diseases, vol. 36, no. 11, pp. 1374–1382, 2003.

[21] S. Gies, S. O. Coulibaly, F. T. Ouattara, C. Ky, B. J. Brabin,and U. D’Alessandro, “A community effectiveness trial of

strategies promoting intermittent preventive treatment withsulphadoxine-pyrimethamine in pregnant women in ruralBurkina Faso,” Malaria Journal, vol. 7, article 180, 2008.

[22] S. D. Perrault, J. Hajek, K. Zhong et al., “Human immunodefi-ciency virus co-infection increases placental parasite densityand transplacental malaria transmission in western Kenya,”American Journal of Tropical Medicine and Hygiene, vol. 80, no.1, pp. 119–125, 2009.

[23] J. Y. Le Hesran, M. Cot, P. Personne et al., “Maternalpiacental infection with Plasmodium falciparum and malariamorbidity during the first 2 years of life,” American Journal ofEpidemiology, vol. 146, no. 10, pp. 826–831, 1997.

[24] S. C. Redd, J. J. Wirima, R. W. Steketee, J. G. Breman, andD. L. Heymann, “Transplacental transmission of plasmodiumfalciparum in rural Malawi,” American Journal of TropicalMedicine and Hygiene, vol. 55, no. 1, supplement, pp. 57–60,1996.

[25] A. Matteelli, S. Caligaris, F. Castelli, and G. Carosi, “Theplacenta and malaria,” Annals of Tropical Medicine and Para-sitology, vol. 91, no. 7, pp. 803–810, 1997.

[26] M. Adachi, K. Manji, R. Ichimi et al., “Detection of congenitalmalaria by polymerase-chain-reaction methodology in Dar esSalaam, Tanzania,” Parasitology Research, vol. 86, no. 8, pp.615–618, 2000.

[27] O. T. Lamikanra, “A study of malaria parasitaemia in pregnantwomen, placentae, cord blood and newborn babies in Lagos,Nigeria,” West African Journal of Medicine, vol. 12, no. 4, pp.213–217, 1993.

[28] D. H. De Silva, K. N. Mendis, U. N. Premaratne, S. M.Jayatilleke, and P. E. Soyza, “Congenital malaria due to Plas-modium vivax: a case report from Sri Lanka,” Transactions ofthe Royal Society of Tropical Medicine and Hygiene, vol. 76, no.1, pp. 33–35, 1982.

[29] I. Malhotra, P. Mungai, E. Muchiri, J. J. Kwiek, S. R.Meshnick, and C. L. King, “Umbilical cord-blood infectionswith Plasmodium falciparum malaria are acquired antenatallyin Kenya,” Journal of Infectious Diseases, vol. 194, no. 2, pp.176–183, 2006.

[30] O. A. Oduwole, G. C. Ejezie, F. A. Odey et al., “Congenitalmalaria in Calabar, Nigeria: the molecular perspective,” Amer-ican Journal of Tropical Medicine and Hygiene, vol. 84, no. 3,pp. 386–389, 2011.


Research Article

Reorganization of Extracellular Matrix in Placentas fromWomen with Asymptomatic Chagas Disease: Mechanism ofParasite Invasion or Local Placental Defense?

Juan Duaso,1 Erika Yanez,1 Christian Castillo,1 Norbel Galanti,2 Gonzalo Cabrera,2

Gabriela Corral,3 Juan Diego Maya,4 Ines Zulantay,2 Werner Apt,2 and Ulrike Kemmerling1, 5

1 Programa de Anatomıa y Biologıa del Desarrollo, Instituto de Ciencias Biomedicas, Facultad de Medicina, Universidad de Chile,Independencia, Region Metropolitana, 1027 Santiago de Chile, Chile

2 Programa de Biologıa Celular y Molecular, Instituto de Ciencias Biomedicas, Facultad de Medicina, Universidad de Chile,Independencia, Region Metropolitana, 1027 Santiago de Chile, Chile

3 Servicio de Obstetricia y Ginecologıa, Hospital de Illapel, Independencia, IV Region, 0512 Illapel, Chile4 Programa de Farmacologıa Molecular y Clınica, Instituto de Ciencias Biomedicas, Facultad de Medicina, Universidad de Chile,Independencia, Region Metropolitana, 1027 Santiago de Chile, Chile

5 Departamento de Estomatologıa, Facultad de Ciencias de la Salud, Universidad de Talca, Avenida Lircay s/n, VII Region,3460000 Talca, Chile

Correspondence should be addressed to Ulrike Kemmerling, [email protected]

Received 27 April 2011; Accepted 8 August 2011

Academic Editor: Ricardo E. Fretes

Copyright © 2012 Juan Duaso et al. This is an open access article distributed under the Creative Commons Attribution License,which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Chagas disease, produced by the protozoan Trypanosoma cruzi (T. cruzi), is one of the most frequent endemic diseases in LatinAmerica. In spite the fact that in the past few years T. cruzi congenital transmission has become of epidemiological importance,studies about this mechanism of infection are scarce. In order to explore some morphological aspects of this infection in theplacenta, we analyzed placentas from T. cruzi-infected mothers by immunohistochemical and histochemical methods. Infectionin mothers, newborns, and placentas was confirmed by PCR and by immunofluorescence in the placenta. T. cruzi-infected pla-centas present destruction of the syncytiotrophoblast and villous stroma, selective disorganization of the basal lamina, and disor-ganization of collagen I in villous stroma. Our results suggest that the parasite induces reorganization of this tissue component andin this way may regulate both inflammatory and immune responses in the host. Changes in the ECM of placental tissues, togetherwith the immunological status of mother and fetus, and parasite load may determine the probability of congenital transmission ofT. cruzi.

1. Introduction

American Trypanosomiasis, or Chagas disease, is a zoonosiscaused by Trypanosoma cruzi (T. cruzi). Currently, 10 millionpeople in the Americas, from Mexico in the north toArgentina and Chile in the south, are estimated to be infected[1]. For thousands of years, Chagas disease was known onlyin the Region of the Americas, mainly in Latin America,where it has been endemic [2]. In past decades, it has beenincreasingly detected in other non-endemic countries inthe American (Canada and the United States), the WesternPacific (mainly Australia and Japan) and the European conti-nents. The presence of Chagas disease outside Latin America

is the result of population mobility, notably migration, butcases have been reported among travelers returning fromLatin America and even in adopted children [3]. Subsequenttransmission occurs through transfusion, vertical, and trans-plantation routes [3].

The vertical transmission of T. cruzi cannot be prevented,but early detection and treatment of congenital infectionachieves cure rates close to 100 per cent [4].

Fetal and maternal tissues are separated by a fetal epithe-lium (the trophoblast), the greatest area of which is in thevillous placenta, the site of nutrient and gas exchange [5].The human placenta is classified as a hemochorial villous pla-centa in which the free chorionic villi are the functional units.


These chorionic villi are formed by the trophoblast and thevillous stroma. The trophoblast is formed by a single mul-tinucleated cell layer (syncytiotrophoblast) which contactsmaternal blood in the intervillous space, and by the cytotro-phoblast which contains replicating progenitor cells. Thetrophoblast is separated by a basal lamina from the villousstroma, which is connective tissue containing vascular endo-thelium, fibroblasts, and macrophages [5]. Trophoblast, basallaminae, and villous stroma with endothelium of fetal cap-illaries form the placental barrier that must be crossed bydifferent pathogens, including T. cruzi, in order to infect thefetus during vertical transmission [6, 7].

Damage to the villous placenta is almost always accom-panied by inflammation, either in the intervillous space orwithin the fetal villi, and in severe cases is accompanied byloss of the protective trophoblast. Extensive placental damagemay lead to fetal loss, intrauterine growth retardation [5],and clinical manifestations that can be observed in acute con-genital Chagas disease [8, 9].

We have previously demonstrated that T. cruzi inducessyncytiotrophoblast destruction and detachment in chori-onic villi explants, and selective disorganization of basal lam-ina and disorganization of collagen I in the connective tissueof villous stroma [6]. However, no studies observing similarhistopathological alterations in the different tissue compart-ments of chorionic villi from placentas of mothers withChagas disease have been reported.

The aim of the present study was to determine whetherthe tissue disorganization and destruction of chorionic villi,described previously in ex vivo infected chorionic villi ex-plants, can also be observed in placentas from Chagasicwomen. The diagnosis of Chagas disease in the women stud-ied was performed by standard serological testing and PCR.Additionally, the parasite was detected in the placenta byimmunofluorescence and PCR. Histopathological analysis,immunohistochemical studies of basal lamina, and histo-chemical studies of carbohydrate-rich molecules and colla-gen I in villous stroma of these placentas were performed.

2. Material and Methods

2.1. Patients and Diagnosis of T. cruzi Infection. Three preg-nant women with asymptomatic Chagas disease were en-rolled from the hospitals of Illapel, Servicio de Salud deCoquimbo, IV Region of Chile. Informed consent for theexperimental use of the placentas was given by each patientas stipulated by the Code of Ethics of the Faculty of Medicineof the University of Chile. Maternal T. cruzi infectionwas assessed by standard serological techniques and PCR(Figure 1(a) and Table 1). T. cruzi infection in neonates wasdiagnosed by detection of parasites in umbilical cord or inperipheral blood by microscopic examination using hep-arinized microhematocrite tubes and PCR (Figure 1(a) andTable 1). All of the neonates were negative for the presenceof the parasite by direct parasitological examination. Twomothers and their respective newborns were positive forparasite DNA detection by PCR (Figure 1 lines 5 & 7 and8 & 10), the third mother and her newborn were negativefor parasite DNA detection by PCR (Figure 1 lines 11 & 13).

MW 1 2 3 4 5 6 7 8 9 10 11 12 13

400 pd300 pd

Figure 1: Detection of T. cruzi in mother, newborn, and placenta byPCR: A 330-base pair fragment of the T. cruzi satellite DNA wasamplified as described in Material and Methods. MW: molecularmarker, lane 1: Negative control without DNA; lane 2: DNA from T.cruzi epimastigotes; lane 3: noninfected human blood DNA; lane4 non-infected human placenta DNA; lanes 5–7: parasite DNAdetected in peripheral blood (lane 5), in placenta (lane 6), andrespective umbilical chord from neonate (lane 7) of mother 1;lanes 8–10: parasite DNA detected in peripheral blood (lane 8), inplacenta (lane 9), and respective umbilical cord from neonate (lane10) of mother 2; lanes 11–13: parasite DNA detected in peripheralblood (lane 11), in placenta (lane 12) and respective umbilical chordfrom neonate (lane 13) of mother 3. Note that in mother 3 parasiteDNA can only be detected in the placenta. The PCR product wassubjected to electrophoresis in 1.6% agarose gels and stained withethidium bromide. PCR markers from Promega were employed asmolecular weight standards.

Nevertheless, this last mother was positive using standard se-rological techniques. Gestational age was determined by clin-ical and ultrasound analysis. Clinical data of the mothers andnewborns are shown in Table 1.

For parasite DNA detection by PCR, a 330-base pairfragment of the T. cruzi satellite DNA was amplified asdescribed previously [10]. The sequence of the oligonu-cleotides is the following: forward (5-AAATAATGTACG-G(T/G)GAGATGCATGA-3, specific primer 121) and back-ward (5-GGTTCGATTGGGGTTGGTGTAATATA-3 specificprimer122). The PCR product was subjected to electrophore-sis in 1.6% agarose gels and stained with ethidium bromide.PCR markers from Promega were employed as molecularweight standards [10].

2.2. Placentas. We used three placentas from asymptomaticchagasic women. Placentas from control healthy women wereobtained from vaginal or caesarean deliveries of uncompli-cated pregnancies from the Hospital San Jose, Servicio deSalud Norte; Region Metropolitana, Santiago, Chile. Exclu-sion criteria for the control placentas were the following:major fetal abnormalities, placental tumor, intrauterine in-fection, obstetric pathology, and any other maternal disease.Villous tissue was obtained from the central part of cotyle-dons and immediately fixed in 10% formaldehyde in 0.1 Mphosphate buffer (pH 7.3) for 24 hours.

2.3. Histological and Histochemical Techniques. The fixed tis-sues were dehydrated in alcohol, clarified in xylene, embed-ded in paraffin, and sectioned at 5 μm. Paraffin-embeddedhistological sections that were stained with hematoxylin-eosin for routine histological analysis, Picrosirius red-hematoxylin [6] and Arteta [11] for collagen histochemistry,and periodic acid-Schiff (PAS) for carbohydrate containingtissue elements (Sigma-Aldrich kit 395B) were applied.


Table 1: Characterization of clinical features of mothers and neonates.

Mother 1 Mother 2 Mother 3

Age mother (years) 43 32 39

Gestational age (weeks) 38 36 40

Type of delivery Vaginal Cesarean Vaginal

Birth weight (g) 2880 2690 3550

APGAR 5 min 9.9 9.9 9.9

Chagas’ disease diagnosis of the mother ELISA/IFI + ELISA/IFI + ELISA/IFI +

Detection of the parasite in the neonate by direct parasitological examination Negative Negative Negative

Detection of the parasite in the mother by PCR Positive Positive Negative

Detection of the parasite in the neonate by PCR Positive Positive Negative

Detection of the parasite in the placenta by PCR Positive Positive Positive

As a control of PAS method, 5 μm-thick sections wereincubated with a 4 μg/mL solution of α-amylase (NutritionalBiochemical Corporation) in PBS pH 6.0, for 30 minutes at37◦C prior to the PAS reaction (data not shown). A decreasein the intensity of the PAS stain reaction after the enzymetreatment was considered as evidence for the presence ofglycosylated molecules [11].

2.4. Immunohistochemistry. The placental chorionic villiwere fixed, dehydrated, and embedded in paraffin as de-scribed above. Standard immunoperoxidase techniques wereused to show collagen IV, heparan sulphate and fibronectin.The primary antibodies were applied individually to eachsection at 4◦C overnight (anti-collagen IV Novocastra NCL-COLL-IV, dilution 1 : 100 v/v; anti-heparan sulphate Novo-castra NCL-CD44-2, dilution 1 : 40 v/v; anti-fibronectin ABRMA1-83176, dilution 1 : 50 v/v). Immunostaining was per-formed using a horseradish peroxidase-labelled streptavidinbiotin kit (RTU-Vectastain kit) following the manufac-turer’s directions using diaminobenzidine as the chromogen.Sections were counterstained with Mayer’s haematoxylin(DAKO) and mounted with Entellan (Merck). Immunohis-tochemical controls were performed by replacing the pri-mary antibodies with phosphate buffered saline. All con-trols were negative. All sections were examined by lightmicroscopy (Leitz Orthoplan), and images were capturedwith a Canon 1256 camera.

2.5. Detection of T. cruzi in Placental Tissue. T. cruzi invasionwas observed by immunofluorescence and by parasite DNAdetection using the polymerase chain reaction (PCR).

Immunofluorescence. The placental chorionic villi were fixed,dehydrated, and embedded in paraffin as described above. Amonoclonal antibody (mAb 25, dilution 1 : 100 v/v) specificfor the T. cruzi flagellar calcium-binding protein (a gift fromDr. Schenkman, Universidade de Sao Paulo, Sao Paulo,Brasil) was applied to each section overnight at 4◦C. Thepreparations were washed with PBS and incubated with anti-mouse IgG conjugated with fluorescein (ScyTek, ACA) inthe presence of 1 μg/mL of 4′6′diamidino-2-phenylindole(DAPI). Afterwards, sections were mounted in VectaShield(ScyTek, ACA) and observed in a Nikon Eclipse E400 micro-scope (Tokio, Japan).

PCR. Genomic DNA that was extracted from the paraf-fin included placental tissue with the NucleoSpin Tissuekit (Machery-Nagel) according manufacturer’s instructions.The same satellite DNA fragment of T. cruzi, used for parasitedetection in blood of mother and child (see above), wasamplified.

3. Results

3.1. T. cruzi Detection in Mothers, Newborns, and Placentas.The diagnosis of Chagas disease of the three mothers enrolledin our study was performed by standard serological tech-niques. After delivery, blood samples from the mothers andumbilical cords from respective neonates were processed forparasite DNA detection by PCR (Figure 1 lines 5 & 7 (mother& neonate 1), 8 & 10 (mother & neonate 2), 11 & 13 (mother& neonate 3)). DNA from the placentas was obtained fromformalin-fixed placental tissue samples (Figure 1 lines 6, 9and 12 show the results from placentas from mother 1, 2,and 3, resp.). Interestingly, one of the mothers (mother 3)and her newborn were negative for PCR (Figure 1 lines 11and 13) but the placenta was positive (Figure 1 line 12).The negative result for parasitic DNA detection by PCRin this mother and newborn may be due to a very lowparasitemia, characteristic in the asymptomatic and chronicphases of Chagas disease. In these phases, the sensitivityof the techniques depends on the varying concentration ofparasites in blood [1, 10]. For this reason, negative resultsdo not necessarily indicate a lack of parasites in the bloodor absence of infection. Serological diagnosis of T. cruziinfection is more sensitive than PCR [10, 12]; all threemothers analyzed by us had positive serology.

Figure 2 shows the detection of the parasite by immun-ofluorescence. The parasite can be visualized in connectivetissue cells of the villous stroma (Figures 2(e)–2(h)). Pla-centas from healthy, non-T. cruzi-infected, women did notshow evidence of the presence of the parasite (Figures 2(a)–2(d)). Interestingly, parasites do not form amastigote nests(the obligate intracellular replicative form of the parasite) asseen in other tissues like the myocardium [13, 14]. In theplacentas of T. cruzi-infected women, only few parasites canbe visualized. The limited presence of the parasite maybe dueto a very low parasitemia present in the asymptomatic orchronic phases of the disease. Another explanation for this


Phase contrast

(a)

DAPI

(b)

T.cruzi

(c)

Merged

(d)

(e) (f) (g) (h)

Figure 2: Detection of T. cruzi in human placental chorionic villi from infected mothers. The presence of parasites in villous stroma of chorionicvilli from T. cruzi infected placentas was shown by immunofluorescence, detected using mAb 25 antibodies. In (a–d), noninfected chorionicvilli (controls) are shown. Panels (a–d) and (e–h) show images of the same fields. Panels (a) and (e): phase contrast; (b) and (f): nuclearstaining with DAPI; (c) and (g): detection of the parasite by immunofluorescence; (d) and (h): merged images of (b-c) and (f-g), respectively.Inserts show an amplified region of the detection of the parasite by immunofluorescence (g-h). Bar scale (a–d): 50 μm; (e–h): 25 μm.

(a) (b)

Figure 3: Histopathological analysis of placentas from mothers infected with T. cruzi. Chorionic villi from T. cruzi-infected placentas(b) show severe tissue damage compared to control villi from healthy noninfected woman (a). Detachment and disorganization of thesyncytiotrophoblast (arrow) as well as fetal connective tissue destruction of the villous stroma (arrowhead) is observed. Chorionic villi wereprocessed for routine histological techniques and stained with hematoxylin-eosin. Bar scale: 25 μm.

fact may be the presence of a local placental antiparasiticmechanism, like the increase of nitric oxide which has anti-trypanosomal activity [15]. We observed the same limitedpresence of the parasite in the ex vivo infected explants ofchorionic villi [6].

3.2. Histopathological Analysis of Placentas from Women Infec-ted with T. cruzi. Figure 3(b) shows severe tissue disorgani-zation and destruction of the chorionic villi from T. cruzi-infected mothers, detachment of the syncytiotrophoblast(arrows), and destruction of villous stroma (arrowhead). Noinflammatory infiltrate is observed in placentas from infectedwomen or control placentas. The control placentas fromhealthy women show an intact structure of the chorionic villi(Figure 3(a)).

3.3. Placental Chorionic Villi from Women Infected withT. cruzi Present Disorganization of the Extracellular Matrix.The extracellular matrix (ECM) is the noncellular compo-nent present in all tissues and organs and not only providesessential physical scaffolding for the cellular constituents,but also initiates crucial biochemical and biomechanical cuesthat are required for different tissue functions [16]. The ECMis composed of collagens, elastin, proteoglycans (includinghyaluronan), and noncollagenous glycoproteins forming acomplex, three-dimensional network among the cells of dif-ferent tissues [17]. The non-collagenous proteins and pro-teoglycans are highly glycosylated and especially abundantin basal lamina [6, 18]. Several lines of evidence haveshown that T. cruzi interacts with host ECM componentsproducing breakdown products that play an important role


(a) (b) (c)

(d) (e) (f)

Figure 4: Placentas from mothers infected with T. cruzi present severe disorganization of chorionic villi ECM. Placental tissue from healthynoninfected (a–c) and infected (d–f) women were processed for routine histochemical methods and stained with the PAS method for thedetection of glycosylated components (a, d) with the Arteta trichromic (b, e), and picrosirius red-hematoxylin (c, f) methods for collagenhistochemistry. Placental chorionic villi from infected mothers show reduced staining of glycosylated molecules (d), especially in basal lamina(arrows) compared to controls (a). Samples from infected mothers also show reduced histochemical reaction for collagen in basal lamina (e)and in villous stroma (f) compared to controls (b, c). Bar scale: 20 μm.

in parasite mobilization and infectivity [6, 19]. We used PASreagent to analyze the non-collagenous component of theECM. The placentas from Chagasic women (Figure 4(d))show a significant decrease in PAS reactivity compared tocontrol tissue (Figure 4(a)). This reduced PAS reactivity isparticularly evident in basal lamina (Figure 4(d) arrows).

The principal fibrous components of chorionic villi con-nective tissue of the villous stroma are collagen fibers, partic-ularly collagen I [5]. Collagen fibers form the “basic skeleton”of the ECM three-dimensional network. We performedtwo histochemical stains to analyze the collagen molecules:arteta staining for collagen in basal laminae (Figures 4(b)and 4(e)) and Picrosirius red-hematoxylin for collagen I(Figures 4(c) and 4(f)). Histochemical reaction for collagenwas reduced in the placentas from mothers infected withthe parasite (Figures 4(e)–4(f)), indicating destruction ordisorganization of the collagen fibers present in basal laminaor villous stroma of the chorionic villi.

3.4. Placental Chorionic Villi from Women Infected withT. cruzi Present Selective Disorganization of Basal Lamina.The basal lamina is part of the placental barrier that parasiteshave to cross in order to invade the fetal connective tissue ofthe villous stroma containing fetal capillaries. PAS and Artetahistochemical stains showed alteration in this specializedstructure (Figures 4(d) and 4(e)). Collagen IV, heparan sul-phate and fibronectin are components of the basal lamina[18]. We have previously shown that the parasite affects the

immunoreactivity of heparan sulphate and collagen IV inex vivo infected chorionic villi explants in a parasite con-centration-dependent manner, and that fibronectin is notaffected [6]. Placentas from T. cruzi-infected women showsimilar results; the immunoreactivity for heparan sulphate isloose or markedly decreased (Figure 5(d)). The immunore-activity of collagen IV is decreased (Figure 5(e)) and nochange in this parameter can be observed for fibronectin(Figure 5(f)) with respect to placenta from healthy women.

4. Discussion

During congenital T. cruzi infection, the parasite reaches thefetus by crossing the placental barrier [6–8]. Congenital Cha-gas disease is a product of a complex interaction betweenthe maternal immune response, placental factors, and theparasite [20].

Congenitally infected newborns develop a parasite-spe-cific T-cell immune response comparable to that of adults[21], as well as phenotypic and functional modificationsof their NK cells [22]. On the other hand, newborns ofT. cruzi-infected mothers are prone to produce higher levelsof proinflammatory cytokines in comparison to those bornto non-infected mothers [23]. From these observations,some authors postulated that some newborns from T. cruzi-infected mothers might naturally autocure their congenitalinfection [24]. Interestingly, the placentas from women


Heparan sulphate

(a)

Collagen IV

(b)

Fibronectin

(c)

(d) (e) (f)

Figure 5: Placentas from mothers infected with T. cruzi present selective basal lamina disorganization in chorionic villi. Placental tissue fromcontrol non-infected (a–c) and infected (d–f) women were immunostained for heparan sulphate (a, d), collagen IV (b, e), and fibronectin(c, f). Placental chorionic villi from infected mothers completely lost their immunoreactivity for heparan sulphate (d) and showed reducedimmunoreactivity for collagen IV (e), compared to controls (a, b). No difference in immunoreactivity for fibronectin was observed betweencontrols and infected chorionic villi (c, f). Chorionic villi were processed for routine immunohistological techniques and counterstainedwith Mayer’s hematoxylin. Bar scale: 20 μm.

with asymptomatic Chagas disease analyzed by us do notpresent inflammatory infiltrate. This aspect might representan interesting aspect of the placental infection by T. cruzi,because of scarce invasion and/or destruction of the parasitewithin placenta. The lack of inflammation may be due tolow parasite load in blood and tissues, characteristic ofasymptomatic and chronic phases of the disease. Thus, otherfactors may also determine the capacity of the parasite toinfect the placenta or, alternatively, impair the congenitaltransmission.

Parasitemia is one factor associated with the risk of con-genital transmission of the parasite. A high parasitemia, ascan be detected in acute infection, correlates with a highertransmission rate of around 50%. In chronic infected pa-tients, with very low parasitemia, the transmission rate isbetween 1 and 21% [25].

Recently, increasing evidence of the presence of local pla-cental antiparasite factors has been reported. Among them,nitric oxide synthesis in placental tissue [15] and “heat-sensitive” agents [26] have been involved in local placentaldefense mechanisms.

The ECM is another factor that should be consideredduring tissue invasion and pathogen infection. The ECM is adynamic structure that interacts with cells and generates sig-nals through feedback loops to control the behavior of cells.

Thus, ECM macromolecules are bioactive and modulate cel-lular events such as adhesion, migration, proliferation, dif-ferentiation, and survival [27]. Additionally, ECM moleculesare strictly organized, and this organization determines thebioactivity of the ECM. Even minor alterations such as asingle amino acid substitution in a single ECM componentcan lead not only to altered physicochemical properties oftissues but also to changes in cellular phenotype and cell-matrix interactions [17]. It has been proposed that thesechanges in ECM structure and bioactivity in tissue functionultimately lead to development of disease [17]. It has beenproposed that ECM alterations produced by T. cruzi not onlypromote its motility in tissues and its entrance into cells, butalso alter the presence of cytokines and chemokines, whichin turn permit T. cruzi to modulate and escape both theinflammatory response and the immune response [6, 19].Alternatively, these changes in ECM function may be part oflocal placental defense mechanisms, which could explain whyonly very few parasites can be detected in the placenta.

During tissue invasion, T. cruzi is able to interact withdifferent elements of the ECM. Thus, the parasite presentssurface molecules, such as gp85 [19] and gp83 [28], glyco-proteins that bind to laminin and fibronectin [19, 28] as wellas to sulphated glycosaminoglicans such as heparan sulphate[29]. The parasite can induce the expression of ECM


molecules or decrease their presence [19]. The more obviousexplanation for the decrease of ECM is that the parasitedestroys the ECM by secretion of proteases like cruzipain[6, 30], which can degrade collagen I and IV [30], or byinduction of matrix metalloproteases [31]. However, anotherpossibility is that T. cruzi decreases the expression of thesemolecules by modulation of signal transduction pathways orcytoskeleton rearrangement. Probably, during tissue invasionT. cruzi first binds to molecules of the basal lamina, facilitat-ing its internalization to the different cells in the underlyingconnective tissue. T. cruzi infection decreases fibronectinexpression in cardiomyocyte culture. On the contrary, lam-inin protein expression levels do not change, but the distri-bution of this ECM component changes dramatically in thistype of cell culture [32]. We have shown previously that inex vivo infected human chorionic villi, the immunoreactivityfor heparansulphate decreases or disappears completely in aparasite-concentration-dependent manner. Fibronectin doesnot present changes in its immunoreactivity or in its distri-bution pattern [6]. Here we confirm these results in placentasfrom mothers with asymptomatic Chagas’ disease. In theex vivo infection of chorionic villi, the immunoractivity forcollagen IV decreases in the basal lamina between the tro-phoblast and villous stroma, but not around fetal capillaries[6]. In the placentas from the Chagasic women, collagen IVimmunoreactivity is decreased in both basal laminae. It isprobable that chronic exposure to the parasite during theentire pregnancy causes a more severe change in ECM than a24-hour challenge in ex vivo infected placental tissue.

5. Conclusion

The evidence that placentas from T. cruzi-infected womenpresent severe ECM alterations indicates that the parasiteinduces reorganization of this tissue component in such away that may regulate inflammatory and immune responsesin the host. If this is the case, changes in placental tissueECM, together with the immunological status of mother andfetus, and parasite load may determine the probability ofcongenital transmission of T. cruzi.

Acknowledgments

This study was supported by Grants 11080166 (to UlrikeKemmerling), 1090124 (to Norbel Galanti), 1090078 (to JuanDiego Maya), 1080445 (to Werner Apt), from FONDECYT,Chile and by Grant CONICYT-PBCT Anillo ACT 112, Chile.

References

[1] First WHO report on neglected tropical diseases: working toovercome the global impact of neglected tropical diseases,2010.

[2] A. Araujo, A. M. Jansen, K. Reinhard, and L. F. Ferreira, “Pale-oparasitology of chagas disease—a review,” Memorias do Insti-tuto Oswaldo Cruz, vol. 104, supplement 1, pp. 9–16, 2009.

[3] G. A. Schmunis, “Epidemiology of Chagas disease in non-endemic countries: the role of international migration,” Me-morias do Instituto Oswaldo Cruz, vol. 102, supplement 1, pp.75–85, 2007.

[4] Chagas disease control and prevention in Europe. Report ofa WHO Informal Consultation (jointly organized by WHOheadquarters and the WHO Regional Office for Europe),World Health Organization, Geneva, Switzerland, 2010.

[5] K. Berniscke, P. Kaufmann, and R. N. Baergen, Pathology of theHuman Placenta, Springer, New York, NY, USA, 2006.

[6] J. Duaso, G. Rojo, G. Cabrera et al., “Trypanosoma cruziinduces tissue disorganization and destruction of chorionicvilli in an ex vivo infection model of human placenta,” Pla-centa, vol. 31, no. 8, pp. 705–711, 2010.

[7] U. Kemmerling, C. Bosco, and N. Galanti, “Infection and inva-sion mechanisms of Trypanosoma cruzi in the congenitaltransmission of chagas’ disease: a proposal,” Biological Re-search, vol. 43, no. 3, pp. 307–316, 2010.

[8] A. L. Bittencourt, “Congenital chagas disease,” American Jour-nal of Diseases of Children, vol. 130, no. 1, pp. 97–103, 1976.

[9] A. M. Altemani, A. L. Bittencourt, and A. M. A. Lana,“Immunohistochemical characterization of the inflammatoryinfiltrate in placental Chagas’ disease: a qualitative and quan-titative analysis,” American Journal of Tropical Medicine andHygiene, vol. 62, no. 2, pp. 319–324, 2000.

[10] I. Zulantay, P. Honores, A. Solari et al., “Use of polymerasechain reaction (PCR) and hybridization assays to detectTrypanosoma cruzi in chronic chagasic patients treated withitraconazole or allopurinol,” Diagnostic Microbiology andInfectious Disease, vol. 48, no. 4, pp. 253–257, 2004.

[11] G. Cabrera, I. Espinoza, U. Kemmerling, and N. Galanti,“Mesocestoides corti: morphological features and glycogenmobilization during in vitro differentiation from larva to adultworm,” Parasitology, vol. 137, no. 3, pp. 373–384, 2010.

[12] G. E. B. Marcon, P. D. Andrade, D. M. De Albuquerque et al.,“Use of a nested polymerase chain reaction (N-PCR) to detectTrypanosoma cruzi in blood samples from chronic chagasicpatients and patients with doubtful serologies,” DiagnosticMicrobiology and Infectious Disease, vol. 43, no. 1, pp. 39–43,2002.

[13] M. Faundez, R. Lopez-Munoz, G. Torres et al., “Buthioninesulfoximine has anti-Trypanosoma cruzi activity in a murinemodel of acute Chagas’ disease and enhances the efficacy ofnifurtimox,” Antimicrobial Agents and Chemotherapy, vol. 52,no. 5, pp. 1837–1839, 2008.

[14] J. R. Coura, “Chagas disease: what is known and what isneeded—a background article,” Memorias do Instituto Os-waldo Cruz, vol. 102, no. 1, pp. 113–122, 2007.

[15] M. F. Triquell, C. Dıaz-Lujan, H. Freilij, P. Paglini, and R.E. Fretes, “Placental infection by two subpopulations of Try-panosoma cruzi is conditioned by differential survival of theparasite in a deleterious placental medium and not by tissuereproduction,” Transactions of the Royal Society of TropicalMedicine and Hygiene, vol. 103, no. 10, pp. 1011–1018, 2009.

[16] C. Frantz, K. M. Stewart, and V. M. Weaver, “The extracellularmatrix at a glance,” Journal of Cell Science, vol. 123, no. 24, pp.4195–4200, 2010.

[17] H. Jarvelainen, A. Sainio, M. Koulu, T. N. Wight, and R.Penttinen, “Extracellular matrix molecules: potential targets inpharmacotherapy,” Pharmacological Reviews, vol. 61, no. 2, pp.198–223, 2009.

[18] B. Alberts, A. Johnson, J. Lewis, M. Raff, K. Roberts, and P.Walter, Molecular Biology of the Cell, Garland Science, NewYork, NY, USA, 5th edition, 2008.

[19] A. P. M. P. Marino, A. A. Silva, R. T. Pinho, and J. Lannes-Vieira, “Trypanosoma cruzi infection: a continuous invader-host cell cross talk with participation of extracellular matrixand adhesion and chemoattractant molecules,” Brazilian


Journal of Medical and Biological Research, vol. 36, no. 8, pp.1121–1133, 2003.

[20] Y. Carlier, “Factors and mechanisms involved in the trans-mission and development of congenital infection with Try-panosoma cruzi,” Revista da Sociedade Brasileira de MedicinaTropical, vol. 38, pp. 105–107, 2005.

[21] E. Hermann, C. Truyens, C. Alonso-Vega et al., “Humanfetuses are able to mount an adultlike CD8 T-cell response,”Blood, vol. 100, no. 6, pp. 2153–2158, 2002.

[22] E. Hermann, C. Alonso-Vega, A. Berthe et al., “Human con-genital infection with Trypanosoma cruzi induces phenotypicand functional modifications of cord blood NK cells,” PediatricResearch, vol. 60, no. 1, pp. 38–43, 2006.

[23] J. Vekemans, C. Truyens, F. Torrico et al., “Maternal Try-panosoma cruzi infection upregulates capacity of uninfectedneonate cells to produce pro- and anti-inflammatory cyto-kines,” Infection and Immunity, vol. 68, no. 9, pp. 5430–5434,2000.

[24] Y. Carlier and C. Truyens, “Maternal-Fetal Transmission ofTrypanosoma cruzi,” in American Trypanosomiasis ChagasDisease One Hundred years of Research, J. Telleria and M.Tibayrenc, Eds., pp. 539–581, Elsevier, New York, NY, USA,2010.

[25] E. Moretti, B. Basso, I. Castro et al., “Chagas’ disease: study ofcongenital transmission in cases of acute maternal infection,”Revista da Sociedade Brasileira de Medicina Tropical, vol. 38,no. 1, pp. 53–55, 2005.

[26] C. D. Lujan, M. F. Triquell, A. Sembaj, C. E. Guerrero, andR. E. Fretes, “Trypanosoma cruzi: productive infection is notallowed by chorionic villous explant from normal humanplacenta in vitro,” Experimental Parasitology, vol. 108, no. 3-4,pp. 176–181, 2004.

[27] W. P. Daley, S. B. Peters, and M. Larsen, “Extracellular matrixdynamics in development and regenerative medicine,” Journalof Cell Science, vol. 121, no. 3, pp. 255–264, 2008.

[28] P. N. Nde, K. J. Simmons, Y. Y. Kleshchenko, S. Pratap, M. F.Lima, and F. Villalta, “Silencing of the larninin γ-1 gene blocksTrypanosoma cruzi infection,” Infection and Immunity, vol. 74,no. 3, pp. 1643–1648, 2006.

[29] A. P. C. A. Lima, P. C. Almeida, I. L. S. Tersariol et al., “Heparansulfate modulates kinin release by Trypanosoma cruzi throughthe activity of cruzipain,” The Journal of Biological Chemistry,vol. 277, no. 8, pp. 5875–5881, 2002.

[30] J. Scharfstein and A. Morrot, “A role for extracellular amastig-otes in the immunopathology of chagas disease,” Memorias doInstituto Oswaldo Cruz, vol. 94, supplement 1, pp. 51–63, 1999.

[31] F. R. S. Gutierrez, M. M. Lalu, F. S. Mariano et al., “Increasedactivities of cardiac matrix metalloproteinases matrix metallo-proteinase (MMP)-2 and MMP-9 are associated with mortal-ity during the acute phase of experimental Trypanosoma cruziinfection,” Journal of Infectious Diseases, vol. 197, no. 10, pp.1468–1476, 2008.

[32] C. M. Calvet, F. O. R. Oliveira, T. C. Araujo-Jorge, and M. C.S. Pereira, “Regulation of extracellular matrix expression anddistribution in Trypanosoma cruzi-infected cardiomyocytes,”International Journal of Medical Microbiology, vol. 299, no. 4,pp. 301–312, 2009.


Review Article

Prevention of Congenital Transmission of Malaria inSub-Saharan African Countries: Challenges and Implications forHealth System Strengthening

Kayode O. Osungbade1 and Olubunmi O. Oladunjoye2

1 Department of Health Policy and Management, Faculty of Public Health, College of Medicine and University College Hospital,University of Ibadan, PMB 5017 General Post Office, 200212 Ibadan, Nigeria

2 Department of Community Medicine, University College Hospital, PMB 5116, 200212 Ibadan, Nigeria

Correspondence should be addressed to Kayode O. Osungbade, [email protected]

Received 1 June 2011; Accepted 27 July 2011


Copyright © 2012 K. O. Osungbade and O. O. Oladunjoye. This is an open access article distributed under the Creative CommonsAttribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work isproperly cited.

Objectives. Review of burden of congenital transmission of malaria, challenges of preventive measures, and implications for healthsystem strengthening in sub-Saharan Africa. Methods. Literature from Pubmed (MEDLINE), Biomed central, Google Scholar,and Cochrane Database were reviewed. Results. The prevalence of congenital malaria in sub-Saharan Africa ranges from 0 to23%. Diagnosis and existing preventive measures are constantly hindered by weak health systems and sociocultural issues. WHOstrategic framework for prevention: intermittent preventive therapy (IPT), insecticide-treated nets (ITNs), and case managementof malaria illness and anaemia remain highly promising; though, specific interventions are required to strengthen the healthsystems in order to improve the effectiveness of these measures. Conclusion. Congenital malaria remains a public health burdenin sub-Saharan Africa. Overcoming the challenges of the preventive measures hinges on the ability of national governments anddevelopment partners in responding to the weak health systems.

1. Introduction

Malaria remains a significant burden in sub-Saharan Africaas it continues to be the leading cause of infant morbidity andmortality in Africa. It is believed that malaria contributes upto about 25% of infant mortality in Nigeria [1]. Furthermore,in areas of Africa with stable transmission, Plasmodium fal-ciparum infection during pregnancy is estimated to cause asmany as 10,000 maternal deaths each year, 8% to 14% of alllow birth weight babies, and 3% to 8% of all infant deaths[2]. Thus, malaria burden, if not reduced, poses a threat tothe attainment of the Millennium Development Goals 4 and6.

Congenital malaria is defined as malaria in a newborn orinfant, transmitted from the mother [3]. Congenital malariais generally defined as malaria acquired by the fetus ornewborn directly from the mother, either in utero or duringdelivery [4]. It is also defined as the presence of malarial

parasites in the peripheral smear of the newborn from twentyfour hours to seven days of life [5]. Malaria is consideredto be congenital in the neonate when asexual parasites aredetected in the peripheral blood within the first week of life[6, 7].

Congenital malaria was thought to be rare in developingcountries [8, 9]. This might be due to a number of reasons.Firstly, the protective effect of the foetal haemoglobin (HbF)in a newborn is expected to exert its influence during theimmediate neonatal period [10, 11]. Secondly, local healthfacilities in resource-limited settings often lack the capacityto diagnose malarial infection [12–14]. Thirdly, the clinicalsigns of neonatal malaria may be indistinguishable fromthose of neonatal sepsis [15]. Thus, congenital malaria mightpass unnoticed, and this might be responsible for its under-reporting. Therefore, the aim of this paper was to highlightthe prevalence and burden of congenital malaria in sub-Saharan Africa as well as the challenges of its diagnosis and


prevention; the findings were used to suggest measures aimedat strengthening health systems and reducing the burden ofcongenital malaria in the population.

2. Methods

Literature providing information on the prevalence, burden,diagnosis, prevention, and control of congenital malaria,published between 2000 and 2010 was reviewed. Searchterms included “congenital,” “Malaria,” “burden,” “diag-nosis,” “prevention”, “control,” and “sub-Saharan Africa.”These literatures were accessed from Pubmed (MEDLINE),Biomed central, Google Scholar, and Cochrane Database ofSystematic Reviews. Searches were also supplemented withrecommendations from outside experts, reviews of bibliogra-phies of other relevant articles, and systematic reviews.

3. Results

3.1. Prevalence of Congenital Malaria in Sub-Saharan AfricanCountries. Congenital malaria is rare in developed countriessuch as the United States of America, where only threeinfants out of 4.1 million live births were reported to havecongenital malaria [16]. The situation is however differentand widely varied in sub-Saharan African countries. To thisend, recent reports have suggested that congenital malariais not as rare in developing countries as previously thought[17]. For example, in a survey covering seven sites in sub-Saharan Africa, Fischer showed a mean prevalence rate of 7%for congenital malaria (range 0–23%) [9].

Furthermore, while a study in Kenya reported a preva-lence of malarial parasitaemia in <0.5% of neonatal admis-sions [18], high prevalence of congenital and neonatal par-asitaemia (>20%) was reported in Uganda and Zambia [19].A study done in Ghana revealed an incidence of 13.6% con-genital malaria infections [20]. In Nigeria, congenital malariawas documented in 13.6% of babies at delivery [21]. Inanother multicentre study in Nigeria, the overall incidenceof congenital malaria was 5.1% (range 1.1%–11.5%) [6]. Inaddition, Runsewe-Abiodun et al. have reported a congenitalmalaria prevalence rate of 17.4% among sick neonates in atertiary hospital in southwestern Nigeria [17].

3.2. Burden of Congenital Malaria in Sub-Saharan Africa. Itis estimated that placental malaria is responsible for 35% ofpreventable low birth weight in developing countries [22]. Itis also associated with intrauterine growth restriction, pre-maturity due to preterm labour and intrauterine foetal death[23]. Malaria in pregnancy is reported to cause between75,000 and 200,000 infant deaths each year in sub-SaharanAfrica [22].

3.3. Challenges in Detection of Congenital Malaria. The de-tection of malaria parasites in the infant’s blood is essentialfor diagnosis, although blood smears can be negative if thereare low parasite counts (50 parasites/µL blood) [23]. Pengsaasuggested that three repeated blood smears over 48 hoursshould be reported negative before excluding the diagnosis

[19]. The capacity to conduct blood test for the diagnosis ofcongenital malaria is however limited in sub-Saharan Africancountries. This is because studies have shown a persistentlack of capacity to conduct quality malaria diagnostic testsby local health facilities [12–14].

Cord malaria can also be detected by polymerase chainreaction (PCR). The use of PCR has suggested that congenitalmalaria may be more frequent, although it is unclear if apositive PCR represents an active infection [4]. Though, PCRis a more sensitive and accurate diagnostic technique thanblood smear microscopy, its use is however very limited indeveloping countries as it is expensive and requires a spe-cialized laboratory [4].

Apart from the laboratory constraints, low antenatal careattendance and skilled delivery rates in African countries willalso affect detection and prevention of congenital malaria.Antenatal care utilization in the developing countries isabout 65%; this is low compared to that of the developedcountries, which is 97% [24]. The implication is that inabout two thirds of pregnant women in sub-Saharan Africa,opportunities for early recognition and prompt treatment ofmalaria infection are missed.

3.4. Challenges in the Clinical Diagnosis. The clinical diag-nosis of congenital malaria usually poses a challenge. Thisis because its clinical findings may be indistinguishablefrom those of neonatal sepsis [15]; hence, the condition iseither detected late or not even suspected unless neonatesare specifically screened for it. Furthermore, as with manycongenital infections, the new born child can manifest withfever, irritability, feeding problems, hepatosplenomegaly,anemia, jaundice, and low birth weight [25].

Thus, the clinical distinction from other congenitalinfections rests primarily on maternal history of exposureto malaria and absence of a skin rash [23]. A study donein Calabar, Nigeria, reported that 35% of newborns withclinical signs of neonatal sepsis were found to have congenitalmalaria [26]. Fever was the most common feature reportedby studies in Kenya and Nigeria among others, whichinclude refusal to feed or poor feeding, irritability, anaemia,hepatosplenomegaly, and jaundice [6].

3.5. Challenges in Prevention of Congenital Malaria in Sub-Saharan African Countries. Prevention of any disease con-dition requires the availability of methods for prediction ofthose at high risk of the disorder. If there were tests whichare adequately sensitive for detecting placenta malaria inthe antenatal period, it would have helped in assessing theefficacy of antimalarial drug during pregnancy and iden-tifying the infants at risk of congenital malaria [27]. However,there are no tests which are adequately sensitive for detectingplacental malaria; hence, the difficulty in identifying those athigh risk of this condition.

Therefore, malaria preventive measures in pregnancystill remain as priority interventions required to protect thefoetus and the newborn against the adverse effects of con-genital malaria. The World Health Organization (WHO)has recommended a three-pronged strategic framework in


areas of high or moderate (stable) malaria transmission ofsub-Saharan Africa: intermittent preventive therapy (IPT),insecticide-treated nets (ITNs), and case management ofmalaria illness and anaemia [28].

3.5.1. Intermittent Preventive Therapy (IPT). Intermittentpreventive therapy (IPT), also known as chemoprophylaxisfor pregnant women, especially those in their first pregnan-cies, has been widely used in sub-Saharan Africa. In line withWHO recommendation, most national guidelines stipulatethat all pregnant women should receive at least two dosesof IPT given as sulphadoxine-pyrimethamine (SP) combi-nation after quickening as part of preventive treatmentsat antenatal care [28]. Evidence abound that this measurehas recorded successes. For example, researchers in Kenyaand Malawi have shown that intermittent preventive therapywith sulphadoxine-pyrimethamine significantly reduces theprevalence of maternal anaemia and placental parasitaemia,and the incidence of low birth weight [29–32]. Furthermore,sulphadoxine-pyrimethamine has been found to be safein pregnancy and efficacious in reproductive-age women;hence, its strong acceptance and coverage levels of greaterthan 80% for the first dose [29].

Despite the beneficial impact of sulphadoxine-pyrxim-ethamine on maternal and infant health, its utilization isthreatened by weak health systems and sociocultural issuesin sub-Saharan Africa. Studies have shown that a substantialproportion (20% to 80%) of pregnant women in this settingmake their first antenatal visit in their third trimester[33–35]. Furthermore, unbooked pregnancies are common[36, 37], and antenatal care utilization rate (i.e., antenatalcare clinic attendance of at least once during most recentpregnancy) varies from 60% to 90% [1, 38, 39]. Thesefindings could be partly attributed to low quality of serviceand sociocultural reasons. While evidence abound on thepersistent declining quality of antenatal services provided topregnant women [33, 35, 40], other researchers have foundthat perceived quality of service was the most importantfactor which influenced the choice of facility for obstetriccare [41]; similarly, a perceived lack of quality in the antenatalcare was associated with a late first antenatal visit in Kenya[35].

Within the sociocultural context, it is thought that non-initiation of care in the first trimester seems to be a wide-spread cultural practice in sub-Saharan Africa. In ruralGambia, women do not usually “announce” pregnancybut wait for other family members to discover it, therebypresenting for care well into the third trimester [42]. Whilefurther qualitative research is required to explore the extentto which late first antenatal visit is affected by culturalpractices, the implication of the findings is that a substantialproportion of pregnant women always remains unprotectedagainst malaria infection because of missed opportunity oftreatment in the second trimester. This therefore exposesfetuses of such unprotected women to placental malaria andits adverse effects, thereby contributing to the challengesmilitating against prevention of congenital malaria. This

situation is further worsened by the practice of partial pre-ventive treatments, including sulphadoxine-pyrimethamine(SP) therapy provided by health workers at antenatal careservice [33, 35, 43].

3.5.2. Insecticide-Treated Nets (ITNs). Randomized controltrials in sub-Saharan Africa have consistently shown theeffectiveness of insecticide-treated nets (ITNs) in the pre-vention of malaria in pregnancy. Studies in Ghana andKenya have documented reduced placental malaria, low birthweight, and fetal loss resulting from use of ITNs [44].Though the use of insecticide-treated nets (ITNs) is re-portedly considered as the most cost-effective method ofmalaria prevention in highly endemic areas, there havealways been some concerns about the preventive strategyin sub-Saharan Africa; these concerns include access, cost,retreatment, and gaps between ownership and use of ITN.

Access to ITNs by vulnerable populations includingpregnant women continues to increase due to the efforts ofnational governments and supporting development agenciesthrough multiple approaches such as stand-alone campaigns,health facilities, and antenatal clinic [1, 38]. Access is alsoenhanced by free distribution and sales of ITNs at subsidizedcost or through discounted voucher systems [1, 38, 45], whilethe introduction of long-lasting insecticidal nets (LLINs) hasreduced the burden of frequent retreatments.

Despite these efforts, recent findings revealed low utiliza-tion of ITNs among pregnant women. Though increasingtrends in ownership and use of ITNs by households werereported in national demographic surveys, the reportsindicated that 5% to 57% of pregnant women aged 15–49years slept under an ITN the past night. Furthermore, gapsbetween ownership and use of an ITN continue to exist asabout 50% or less of pregnant women aged 15–49 years inhouseholds with an ITN slept under the net the past night[1, 38, 46, 47].

3.5.3. Case Management of Malaria Illness. Supporting andpromoting access to correct, affordable, and appropriatetreatment within 24 hours of the onset of symptomsof malaria is the third essential component of malariaprevention and control during pregnancy in malariousendemic countries of sub-Saharan Africa [28]. The WorldHealth Organization (WHO) had recommended chloro-quine (CQ) in chloroquine-sensitive areas and sulphadoxine-pyrimethamine (SP) in areas with CQ resistance for treat-ment of uncomplicated malaria in pregnancy. Quinine isanother alternative in areas where both CQ and SP are noteffective [28].

However, prompt and effective case management ofmalaria illness is hinged on early and correct diagnosis ofthe condition. As noted above, this is seriously hindered bylack of capacity to conduct quality malaria diagnostic tests bylocal health facilities in sub-Saharan African countries [12–14]. While this situation might have justified the presumptivetreatment of fever with antimalarials (i.e., treating all febrileepisodes suspected of malaria with a full therapeutic doseof antimalarials), such practice might have led to a reduced


susceptibility of parasites to the commonly used antimalarialdrugs in these settings.

3.6. Recommendations

3.6.1. Health System Strengthening. The World Health Orga-nization (WHO) recently recommended prompt parasito-logic confirmation by microscopy or alternatively by rapiddiagnostic tests (RDTs) in all patients suspected of malariabefore treatment is commenced unless parasitological diag-nosis is not accessible [48]. Findings from the previousreview revealed that many peripheral health facilities inresource-poor settings of sub-Saharan Africa lacked thecapacity to conduct quality parasitological diagnosis ofmalaria by microscopy; this is because the procedure islabour-intensive requiring trained staff and quality equip-ment [49, 50].

On the other hand, malaria rapid diagnostic testing(RDT) has been found useful as an attractive alternative toroutine microscopy with good sensitivity and specificity pro-files [48, 51]. In addition, RDTs have the potential to be cost-effective, require no capital investment or electricity, are sim-ple to perform, and are easy to interpret [51, 52]. If appropri-ately introduced and implemented, it also has been shown tohave the potential of improving drug-prescribing behaviourof health workers [53], thereby reducing overtreatment ofmalaria and the problem of drug resistance.

From the foregoing, utilization of RDTs in resource-poorsettings would seem to be an appropriate technology as it hasthe potential of improving the quality of malaria diagnosisand treatment services provided to all cases of malaria,including pregnant women and newborns. Therefore, itis suggested that national governments and developmentpartners in sub-Saharan Africa should support widespreaduse of rapid diagnostic tests (RDTs) for malaria diagnosis.The capacity of local health facilities providing maternaland child health services should be strengthened with theprovision of adequate supplies of equipment and con-sumables required to provide the diagnostic services. Inaddition, targeted trainings and supportive supervision oflocal health staff are highly desirable in order to meet up withthe challenges of newly-assigned task.

In order to increase uptake of IPTs and facilitate promptdiagnosis and treatment of malaria in pregnancy, nationalgovernments and development partners in sub-SaharanAfrican countries should also consider improving the poormaternal health service indicators—nonbooking, late ante-natal visits and low antenatal service rate—as an essentialtask to be concertedly pursued. To achieve this task, strategiesaimed at improving maternal health service should beimplemented. For example, proven-specific interventionssuch as the World Health Organization (WHO) trainingon Life Saving Skills [54] are suggested for use of maternalhealth service managers in addressing the weak healthsystems. Periodic trainings conducted for local health staffcan help address and strengthen health systems, particularlyif targeted at previously reported areas of deficiency suchas declining quality of maternal health service, poor record

keeping practices of health workers, and poor utilization ofintrapartum monitoring tools such as a partograph [33, 34,55].

In addition, health staffs’ familiarization and adherenceto WHO guidelines on provision of effective, efficient, safe,and culturally appropriate services to pregnant women andnewborn under the Integrated Management of Pregnancyand Childbirth (IMPAC) would assist to ensure best prac-tices [56]. It is expected that these efforts would improveutilization of maternal health services, increase IPT uptake,enhance prompt diagnosis and treatment of malaria inpregnancy, and consequently prevent congenital malaria.

On a long term basis, strategies which would bringabout over 80% antenatal service utilization rate should bepursued vigorously. Therefore, it is suggested that redirectingand repackaging health education and service content ofantenatal care service using a social marketing approachacceptable within the sociocultural context would be founduseful. Therefore, placing the antenatal care service contentand its benefits on public agenda through different mediaand matrices may be helpful in the study setting as opposedto the current practice of restricting information to womenwho make antenatal visits.

Despite male economic dominance and decision mak-ing power in developing countries, their involvement inreproductive health issues is reportedly low [57]. However,studies had shown that male involvement would significantlyimprove indicators of maternal health service utilization,[58] and, therefore, it could also improve uptake of IPTsand case management of malaria in pregnancy. Hence, itis recommended that father’s roles in maternal health bedefined, packaged, and incorporated in the antenatal careservice messages meant for dissemination to the public.

With regard to closing the gap between ownership anduse of an ITN, a carefully designed qualitative research maybe useful in eliciting the factors responsible or reasons fornot sleeping under an ITN despite its availability in thehousehold. While it is suggested that national governmentsand development partners should not relent on their effortsin making ITNs universally accessible to pregnant womeneither free of charge or at a subsidized price, experiences ofother researchers on factors which promote or inhibit ITNusage may be found useful in packaging educational mes-sages aimed at promoting its usage. ITN usage promotingfactors such as high perception on the seriousness of malariaand its effect on pregnant women and children, high per-ceived benefit of ITNs in protecting children and pregnantwomen against malaria, and high awareness of the preven-tion of malaria as a better and cheaper option compared withtreatment should be intensified. Whereas inhibitory factorssuch as fear of the chemical that is used to treat nets andunsupportive spouses should be demystified [59].

4. Conclusion

Evidence abound that congenital malaria constitutes a publichealth burden in sub-Saharan Africa. However, efforts ofthe national governments and development partners at


instituting the recommended cost-effective interventionsare continuously thwarted by challenges brought about byweak health systems and sociocultural factors among others,thereby, militating against the progress towards attainmentof Millennium Development Goal (MDG) 6 (Indicator 22,Target 8). Health system strengthening and appropriatepublic health promoting and educating messages deliveredthrough a social marketing approach may be found useful inputting back on track the race towards 2015 with respect toattainment of MDG 6.

References

[1] National Population Commission [Nigeria] and ICF Macro,Nigeria Demographic and Health Survey 2008, National Popu-lation Commission and ICF Macro, Abuja, Nigeria, 2009.

[2] Roll Back Malaria, Malaria in Pregnancy, http://www.rbm.who.int/cmc upload/0/000/015/369/RBMInfosheet 4.htm.

[3] Centers for Disease Control and Prevention, http://www.cdc.gov/malaria/glossary.html.

[4] O. Coll, C. Menendez, F. Botet et al., “Treatment and pre-vention of malaria in pregnancy and Newborn,” Journal ofPerinatal Medicine, vol. 36, pp. 15–29, 2008.

[5] O. Basipinar, Z. Bayraktaroglu, T. Karsligil, A. Bavram, andY. Coskun, “A rare cause of anemia and thrombocytopeniain a newborn: congenital malaria,” The Turkish Journal of Pe-diatrics, vol. 48, pp. 63–65, 2006.

[6] C. Falade, O. Mokuolu, H. Okafor et al., “Epidemiology ofcongenital malaria in Nigeria: a multi-centre study,” TropicalMedicine and International Health, vol. 12, no. 11, pp. 1279–1287, 2007.

[7] Ghana Health Service, “Guidelines for malaria in pregnancy,”http://www.ghanahealthservice.org/.

[8] A. A. Orogadeav, C. Falade, H. U. Okafor et al., “Clinical andlaboratory features of congenital malaria in Nigeria,” Journalof Pediatric Infectious Diseases, vol. 3, pp. 181–187, 2008.

[9] P. R. Fischer, “Congenital malaria: an African survey,” ClinicalPediatrics, vol. 36, no. 7, pp. 411–413, 1997.

[10] H. L. Shear, L. Grinberg, J. Gilman et al., “Transgenic mice ex-pressing human fetal globin are protected from malaria by anovel mechanism,” Blood, vol. 92, no. 7, pp. 2520–2526, 1998.

[11] C. Shulman and E. Dorman, “Clinical features of malaria inpregnancy,” in Essential Malariology, D. A. Warrell and H. M.Gilles, Eds., p. 223, BookPower Formerly ELST, London, UK,4th edition, 2002.

[12] P. A. Phillips-Howard, K. A. Wannemuehler, F. O. Ter Kuile etal., “Diagnostic and prescribing practices in peripheral healthfacilities in rural western Kenya,” American Journal of TropicalMedicine and Hygiene, vol. 68, supplement 4, pp. 44–49, 2003.

[13] B. Hailegiorgis, S. Girma, Z. Melaku et al., “Laboratory ma-laria diagnostic capacity in health facilities in five adminis-trative zones of Oromia Regional State, Ethiopia,” TropicalMedicine and International Health, vol. 15, no. 12, pp. 1449–1457, 2010.

[14] D. Ishengoma, Y. Derua, R. Rwegoshora et al., “The perform-ance of health laboratories and the quality of malaria diagnosisin six districts of Tanzania,” Annals of Tropical Medicine andParasitology, vol. 104, no. 2, pp. 123–135, 2010.

[15] W. I. Olanrewaju, “Malaria in the neonate: report of 2 cases,”West African Journal of Medicine, vol. 18, no. 2, pp. 139–140,1999.

[16] C. R. Lesko, P. M. Arguin, and R. D. Newman, “Congenital ma-laria in the United States: a review of cases from 1966 to 2005,”Archives of Pediatrics and Adolescent Medicine, vol. 161, no. 11,pp. 1062–1067, 2007.

[17] I. T. Runsewe-Abiodun, O. B. Ogunfowora, and B. M. Fetuga,“Neonatal malaria in Nigeria— a 2 year review,” BMCPediatrics, vol. 6, article 19, 2006.

[18] M. K. Mwaniki, A. W. Talbert, F. N. Mturi et al., “Congenitaland neonatal malaria in a rural Kenyan district hospital: aneight-year analysis,” Malaria Journal, vol. 9, no. 1, article 313,2010.

[19] K. Pengsaa, “Congenital malaria in Thailand,” Annals of Trop-ical Paediatrics, vol. 27, no. 2, pp. 133–139, 2007.

[20] G. Wagner, K. Koram, D. McGuinness, S. Bennett, F. Nkru-mah, and E. Riley, “High incidence of asymptomatic malarainfections in a birth cohort of children less than one yearof age in Ghana, detected by multicopy gene polymerasechain reaction,” The American Journal of Tropical Medicine andHygiene, vol. 59, no. 1, pp. 115–123, 1998.

[21] F. Lesi, M. Mukhtar, E. Iroha, and M. Egri-Okwaji, “Clinicalpresentation of congenital malaria at the Lagos UniversityTeaching Hospital,” The Nigerian Journal of Clinical Practice,vol. 13, no. 2, pp. 134–138, 2010.

[22] R. W. Steketee, B. L. Nahlen, M. E. Parise, and C. Menendez,“The burden of malaria in pregnancy in malaria-endemicareas,” The American Journal of Tropical Medicine and Hygiene,vol. 64, supplement 1-2, pp. 28–35, 2001.

[23] B. J. Brabin, “Congenital malaria— a recurrent problem,” An-nals of Tropical Paediatrics, vol. 27, no. 2, pp. 95–98, 2007.

[24] Maternal Mortality in 2005, World Health Organization,Geneva, Switzerland, 2007.

[25] C. Menendez, “Malaria during pregnancy: a priority area ofmalaria research and control,” Parasitology Today, vol. 11, no.5, pp. 178–183, 1995.

[26] A. D. Ekanem, M. U. Anah, and J. J. Udo, “The prevalence ofcongenital malaria among neonates with suspected sepsis inCalabar, Nigeria,” Tropical Doctor, vol. 38, no. 2, pp. 73–76,2008.

[27] H. Guyatt and R. Snow, “Malaria in pregnancy as an indirectcause of infant mortality in sub-SaharanAfrica,” Transactionsof the Royal Society of Tropical Medicine and Hygiene, vol. 95,no. 6, pp. 569–576, 2001.

[28] World Health Organization, “A strategic framework for ma-laria prevention and control during pregnancy in the Africanregion,” WHO Regional Office for Africa, Brazzaville, The Re-public of the Congo, AFR/MAL/04/01, 2004.

[29] F. Verhoeff, B. Brabin, L. Chimsuku, P. Kazembe, W. Russell,and R. Broadhead, “An evaluation of the effects of intermittentsulfadoxine-pyrimethamine treatment in pregnancy on par-asite clearance and risk of low birthweight in rural Malawi,”Annals of Tropical Medicine and Parasitology, vol. 92, no. 2, pp.141–150, 1998.

[30] L. J. Schultz, R. W. Steketee, A. Macheso, P. Kazembe, L. Chit-sulo, and J. J. Wirima, “The efficacy of antimalarial regimenscontaining sulfadoxine-pyrimethamine and/or chloroquine inpreventing peripheral and placental Plasmodium falciparuminfection among pregnant women in Malawi,” The AmericanJournal of Tropical Medicine and Hygiene, vol. 51, no. 5, pp.515–522, 1994.

[31] M. E. Parise, J. G. Ayisi, B. L. Nahlen et al., “Efficacy of sul-fadoxine-pyrimethamine for prevention of placental malariain an area of Kenya with a high prevalence of malaria andhuman immunodeficiency virus infection,” The American


Journal of Tropical Medicine and Hygiene, vol. 59, no. 5, pp.813–822, 1998.

[32] C. Shulman, E. Dorman, F. Cutts et al., “Intermittent sul-phadoxine-pyrimethamine to prevent severe anaemia sec-ondary to malaria in pregnancy: a randomised placebo-controlled trial,” The Lancet, vol. 353, no. 9153, pp. 632–636,1999.

[33] K. Osungbade, S. Oginni, and A. Olumide, “Content of an-tenatal care services in secondary health care facilities inNigeria: implication for quality of maternal health care,” In-ternational Journal for Quality in Health Care, vol. 20, no. 5,pp. 346–351, 2008.

[34] K. O. Osungbade, V. N. Shaahu, O. C. Uchendu et al., “Clinicalaudit of antenatal service provision in Nigeria,” Health Care forWomen International, vol. 32, no. 5, pp. 441–452, 2011.

[35] A. van Eijk, H. Bles, F. Odhiambo et al., “Use of antenatal ser-vices and delivery care among women in rural Western Kenya:a community based survey,” Reproductive Health, vol. 3, article2, 2006.

[36] B. Chigbu, S. Onwere, C. I. Kamanu, C. Aluka, O. Okoro,and E. Adibe, “Pregnancy outcome in booked and unbookedmothers in South Eastern Nigeria,” East African MedicalJournal, vol. 86, no. 6, pp. 267–271, 2009.

[37] A. T. Owolabi, A. O. Fatusi, O. Kuti, A. Adeyemi, S. O. Faturoti,and P. O. Obiajuwa, “Maternal complications and perinataloutcomes in booked and unbooked Nigerian mothers,” Sin-gapore Medical Journal, vol. 49, no. 7, pp. 526–531, 2008.

[38] Ghana Statistical Service, Ghana Health Service, and ICFMacro, Ghana Demographic and Health Survey 2008, GSS,GHS, and ICF Macro, Accra, Ghana, 2009.

[39] S. Babalola and A. Fatusi, “Determinants of use of maternalhealth services in Nigeria—looking beyond individual andhousehold factors,” BMC Pregnancy and Childbirth, vol. 9,article 1471, p. 43, 2009.

[40] A. Sagay, C. Ekwempu, M. Kabiru, P. Daru, and A. Aisien,“Audit of antenatal services in primary healthcare centres inJos, Nigeria,” Tropical Journal of Obstetrics & Gynaecology, vol.22, no. 2, pp. 147–151.

[41] C. Iyaniwura and Q. Yussuf, “Utilization of antenatal care anddelivery services in Sagamu, South Western Nigeria,” AfricanJournal of Reproductive Health, vol. 13, no. 3, pp. 111–122,2009.

[42] M. L. Telfer, J. T. Rowley, and G. E. Walraven, “Experiences ofmothers with antenatal, delivery and postpartum care in ruralGambia,” AfricanJournal of Reproductive Health, vol. 6, no. 1,pp. 74–83, 2002.

[43] National Population Commission (Nigeria) and ORC Macro,Nigeria Demographic and Health Survey 2003, National Popu-lation Commission (Nigeria) and ORC Macro, Calverton, Md,USA, 2004.

[44] C. Gamble, J. P. Ekwaru, and F. O. ter Kuile, “Insecticide-treated nets for preventing malaria in pregnancy,” CochraneDatabase of Systematic Reviews, no. 2, p. CD003755, 2006.

[45] A. Tami, J. Mbati, R. Nathan, H. Mponda, C. Lengeler, and J.Schellenberg, “Use and misuse of a discount voucher schemeas a subsidy for insecticide-treated nets for malaria control insouthern Tanzania,” Health Policy and Planning, vol. 21, no. 1,pp. 1–9, 2005.

[46] Kenya National Bureau of Statistics and ICF Macro, KenyaDemographic and Health Survey 2008–2009, KNBS and ICFMacro, Calverton, Md, USA, 2010.

[47] Tanzania National Bureau of Statistics and ICF Macro, Tan-zania Demographic and Health Survey 2009–2010, KNBS andICF Macro, Calverton, Md, USA, 2010.

[48] World Health Organization, World Malaria Report 2009, 2009.

[49] P. Bloland, S. Kachur, and H. Williams, “Trends in antimalarialdrug deployment in sub-Saharan Africa,” Journal of Experi-mental Biology, vol. 206, no. 21, pp. 3761–3769, 2003.

[50] F. Moerman, C. Lengeler, J. Chimumbwa et al., “The contri-bution of the health-care service to a sound and sustainablemalaria-control policy,” The Lancet Infectious Diseases, vol. 3,no. 2, pp. 99–102, 2003.

[51] C. Wongsrichanalai, M. J. Barcus, S. Muth, A. Sutamihardja,and W. H. Wernsdorfer, “A review of malaria diagnostic tools:microscopy and rapid diagnostic test (RDT),” The AmericanJournal of Tropical Medicine and Hygiene, vol. 77, no. 6, pp.119–127, 2007.

[52] S. Shillcutt, C. Morel, C. Goodman et al., “Cost-effectivenessof malaria diagnostic methods in sub-Saharan Africa in anera of combination therapy,” Bulletin of the World HealthOrganization, vol. 86, no. 2, pp. 101–110, 2008.

[53] G. Bastiaens, E. Schaftenaar, A. Ndaro et al., “Malaria di-agnostic testing and treatment practices in three different Plas-modium falciparum transmission settings in Tanzania: beforeand after a government policy change,” Malaria Journal, vol.10, p. 76, 2011.

[54] World Health Organization, “Making pregnancy safer: thecritical role of the skilled attendant. A joint statement byWHO, ICM and FIGO,” Department of Reproductive Healthand Research, WHO, 2004 http://www.who.int/reproductive-health/publications/2004/skilled attendant.pdf.

[55] O. Oladapo, O. Daniel, and A. Olatunji, “Knowledge anduse of the partograph among health care personnel at theperipheral maternity centres in Nigeria,” Journal of Obstetricsand Gynaecology, vol. 26, no. 6, pp. 538–541, 2006.

[56] World Health Organization, Pregnancy, Childbirth, Postpartumand Newborn Care: A Guide for Essential Practice (IntegratedManagement of Pregnancy and Childbirth), World HealthOrganization, 2nd edition, 2006, http://www.who.int/makingpregnancy safer/publications/PCPNC 2006 03b.pdf.

[57] Z. Iliyasu, I. S. Abubakar, H. S. Galadanci, and M. H. Aliyu,“Birth preparedness, complication readiness and fathers’ par-ticipation in maternity care in a northern Nigerian commu-nity,” African Journal of Reproductive Health, vol. 14, no. 1, pp.21–32, 2010.

[58] L. Martin, M. McNamara, A. Milot, T. Halle, and E. C. Hair,“The effects of father involvement during pregnancy on re-ceipt of prenatal care and maternal smoking,” Maternal andChild Health Journal, vol. 11, no. 6, pp. 595–602, 2007.

[59] U. M. Chukwuocha, I. N. Dozie, C. O. Onwuliri et al.,“Perceptions on the use of insecticide treated nets in partsof the Imo River Basin, Nigeria: implications for preventingmalaria in pregnancy,” African Journal of Reproductive Health,vol. 14, no. 1, pp. 117–128, 2010.


Research Article

In Vitro Infection of Trypanosoma cruzi CausesDecrease in Glucose Transporter Protein-1 (GLUT1) Expressionin Explants of Human Placental Villi Cultured under Normaland High Glucose Concentrations

Luciana Mezzano,1 Gaston Repossi,1, 2 Ricardo E. Fretes,1, 2 Susana Lin,1, 3

Marıa Jose Sartori,1 and Sofıa G. Parisi de Fabro1

1 Catedra de Biologıa Celular, Histologıa y Embriologıa, Instituto de Biologıa Celular, Facultad de Ciencias Medicas,Universidad Nacional de Cordoba, Ciudad Universitaria, 5016 Cordoba, Argentina

2 Catedra de Histologıa, Embriologıa y Genetica, IICSHUM, Universidad Nacional de La Rioja, 5300 La Rioja, Argentina3 Division of Environmental Health and Occupational Medicine, National Health Research Institutes,35053 Zhunan Town, Miaoli County, Taiwan

Correspondence should be addressed to Luciana Mezzano, [email protected]

Received 2 June 2011; Accepted 15 July 2011

Academic Editor: Ulrike Kemmerling

Copyright © 2012 Luciana Mezzano et al. This is an open access article distributed under the Creative Commons AttributionLicense, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properlycited.

Trypanosoma cruzi, the etiologic Chagas’ disease agent, induces changes in protein pattern of the human placenta syncytiotro-phoblast. The glucose transporter protein-1 (GLUT1) is the primary isoform involved in transplacental glucose transport. Wecarried out in vitro assays to determine if T. cruzi infection would induce changes in placental GLUT1 protein expression undernormal and high concentration of glucose. Using Western blot and immunohistological techniques, GLUT1 expression wasdetermined in normal placental villi cultured under normal or high concentrations of glucose, with or without in vitro T. cruziinfection, for 24 and 48 hours. High glucose media or T. cruzi infection alone reduced GLUT1 expression. A yet more accentuatedreduction was observed when infection and high glucose condition took place together. We inform, for the first time, that T. cruziinfection may induce reduction of GLUT1 expression under normal and high glucose concentrations, and this effect is synergic tohigh glucose concentrations.

1. Introduction

Chagas’ disease, endemic in Latin America, is caused byTrypanosoma cruzi a flagellated protozoan with a life cycleinvolving an insect vector and a mammalian host. Thecongenital Chagas’ disease is associated with prematurelabor, miscarriage, and placentitis [1]. In endemic countries,maternal-fetal transmission of T. cruzi is between 1 and 17%of pregnancies in chronically infected mothers, depending ongeographic area [2]. The infectious form of the parasite (try-pomastigote) adheres to specific receptors on the outer mem-brane of host cells previous to intracellular invasion [3], andthe process of invasion requires activation of signal transduc-tion pathways in the parasite and the host cell [4–10].

Pancreas is one of the organs affected in Chagas’ Disease.Patients with this disease have plasma pancreatic glucagonand pancreatic polypeptide levels reduced [11], lower insulinactivity [12], and morphometric and morphologic alter-ations of pancreatic ganglia and islets [13]. Experimentalinfections in hamsters caused pancreatitis, erratic bloodglucose levels, and a tendency to hypoinsulinemia [14].T. cruzi infection-resistant C57BL/6 mice developed highparasitemia and mortality when diabetes was induced withstreptozotocin in them [15]. Above clinical and animalstudies suggest that patients with Chagas’ Disease are moresusceptible to develop hyperglycemia, and this would worsenthe infection. In fact, patients with both conditions havebeen reported, and diabetes or hyperglycemia prevalence was


higher in patients with the cardiac form of Chagas’ disease[16].

Hyperglycemia during pregnancy is a well-recognizedpathogenic factor. Adequate glucose transfer from the mater-nal to the fetal compartment is crucial for the normalsurvival and development of the fetus during pregnancy [17].GLUT1 is the primary isoform involved in the transplacentalmovement of glucose and distributed asymmetrically on themicrovillus and basal membranes syncytiotrophoblast. Themicrovillous membrane contains more transporters than thebasal. GLUT is inversely regulated by glucose concentration,and basal membrane GLUT1 is positively regulated byinsulin-like growth factor I, placental growth hormone, andhypoxia [18]. In vivo, basal membrane GLUT1 is upregu-lated over gestation, increased in diabetic pregnancy, anddecreased in chronic hypoxia, while microvillous membraneGLUT1 is unaffected [18, 19]. As the rate-limiting step intransplacental glucose transport, changes in the density ofbasal membrane GLUT1 will have a significant impact ontransplacental glucose flux [18, 20].

Although increased expression of GLUT1 in placentafrom diabetic pregnant women has been reported [18, 20],decrease in GLUT1 mRNA and protein levels in diabetic micecompared with the control and placental cell cultures underhigh glucose concentration conditions was also informed[20, 21]. Glucose would also alter GLUT1 partitioningbetween the plasma membrane and intracellular sites in favorof the latter [22].

In the invasion process, T. cruzi has been found to affectnumerous surface molecules of the placental villi, probablycausing placental dysfunction as consequence. We wonderedif this parasite would somehow also affect GLUT1 expressionpattern, especially under high glucose (HG) concentration,since T. cruzi has been reported to affect pancreatic functionand alters the insulin-glycemia axis, and there is not yeta marker for placental or fetus infection. In this work,we compared the protein expression of GLUT1 of humanplacental explants infected in vitro with trypomastigotes,cultured under normal and high glucose (HG) con-centration.


2.1. Placentas. Placentas (n = 17) from clinically and sero-logically healthy women at 38 to 40 weeks of gestation wereobtained by caesarian delivery, in order to assure asepsis.Women signed an informed consent. Placentas were kept inglucose solution 0.29 mM at 4◦C for transportation. Oncein the laboratory, central villi of placental cotyledons wereisolated, rinsed with PBS several times, and cut into pieces of3 mm in diameter.

2.2. Parasites. Trypomastigotes of T. cruzi (Tulahuen strain)were isolated according to Andrews and Colli [23] andFretes and Fabro [24], from infected Albino/Swiss micebloodstream at the peak of parasitism. Briefly, mice bloodwas centrifuged for 10 minutes at 100 g and kept still for 1 hat 37◦C. Plasma was then separated and centrifuged for 10

minutes at 590 g. The pellets containing the parasites werewashed twice and suspended in MEM-199 (Gibco Lab., NY,USA).

2.3. Treatments of Placental Explants. Explants of normalhuman placental villi were cultured at 37◦C, in normalatmosphere supplemented with 5% CO2, in MEM-199 (pH7) with 0.1% penicillin and 0.01% streptomycin and eitherwith 5 mM D-glucose (normal D-glucose: NG) (J.T. Baker,NJ, USA) or 25 mM D-glucose (high glucose: HG) con-centrations [22].

After 24 hours, 7 × 105 trypomastigotes Tulahuen strainof T. cruzi were added with the refresh of culture media,prepared as mentioned above. Controls without parasiteswere carried out for both normal and HG concentrations.Cocultures with trypomastigotes and controls were incu-bated for another 24 h or 48 h.

After treatment, the placental explants were rinsed withPBS. Part of the explants were fixed with 10% formaldehydeand included in paraffin. Approximately 30 mg of placentalexplants from each treatment were homogenized in 0.25 mLPBS with an OMNI 1000 homogenizer for five cycles of highspeed application, each lasting ten seconds.

2.4. Analysis of Placental Explants Infection. At the end ofcultures, placental explants were collected, fixed, and stainedwith hematoxylin/eosin and PAS/hematoxylin and observedunder low and high magnifications in a Zeiss Axioskop20 microscope. Infection of placental explants was assessedobserving amastigote groups of the T. cruzi in trophoblast orstromal cells of the chorionic villi.

2.5. Immunostaining of GLUT1 Protein Expression in Pla-cental Explants. Deparaffinized histological sections wereembedded in TBS (pH 7.2), pretreated with 0.05% saponin(15 min) for the unmasking of antigens, then with 3% H2O2

(15 min) to block internal peroxidases, and treated with3% nonfat dry milk in TBS (15 min) to block nonspecificepitopes. GLUT1 protein was detected by incubating thetreated sections with an anti-GLUT1 polyclonal antiserum(rabbit, CHEMICOM International Inc, Temecula, Calif,USA) diluted in TBS/Tween (1 : 500) for overnight, at 4◦C,and revealed with a secondary antirabbit immunoglobu-lin conjugated with peroxidase (Sigma-Aldrich Co, Mo,USA). Peroxidase activity was developed using H2O2/4-Cl-1-naphthol. Background control without addition of anti-GLUT1 antiserum was carried out. We also used preculturedplacenta controls. Images stored as jpg format were analyzedwith “Image Tool” UTHSCSA version 3.00 (download-able from http://ddsdx.uthscsa.edu/dig/download.html) asdescribed previously [25]. Five optical fields were measuredper slide; each treatment for each placenta provided 3 slides.Immunostained area was expressed as ratio of total area(manually selected surface area).

2.6. Western Blot for GLUT1. Homogenized placental villiwere mixed with lysis buffer containing protease inhibitors(1% w/v de SDS, 1 mM EDTA, 1 µg/mL of leupeptin,


(a) (b)

(c) (d)

Figure 1: Paraffin-embedded placental villi cultured for 48 hs, Pas/H stained. (a) Control placental villi without T. cruzi infection culturedwith normal glucose concentration media; (b) normal placental villi in vitro infected with T. cruzi, cultured with normal glucose media; (c)normal placental villi cultured under high glucose concentration media; (d) normal placental villi cultured under high glucose concentrationmedia, in vitro infected with T. cruzi. Amastigotes nests were observed inside the villous from infected placentas (arrows). Arrow pointsindicate increased glycogen deposits in placental villi explants cultured under HG concentration (c and d). Original magnification: 1000x.The scale bar represents 1 µm.

100 mM of Hepes pH 7.4) and centrifuged at 12000 g for 10minutes. Protein content was measured with Lowry proteinassay [26]. The samples were not heated prior electrophore-sis; 40 µg of sample were loaded per lane in a SDS-PAGE gelthat was carried out with 3% stacking gel and 10% resolvinggel at 200V for 1 hr [27] and blotted onto nitrocelluloseat 300 mA for 1.5 hr using a Trans Blot Mini-Protean IIapparatus (BioRad, Richmond, Calif, USA). GLUT1 onnitrocellulose was detected with rabbit anti-GLUT1 poly-clonal antiserum (CHEMICOM International Inc, Temecula,Calif, USA), diluted 1 : 5000 in TBS-Tween and revealedwith a secondary antirabbit immunoglobulin conjugatedwith peroxidase (Sigma-Aldrich Co, Mo, USA). Peroxidaseactivity was developed using H2O2/4-Cl-1-naphthol. Back-ground control without addition of anti-GLUT1 antiserumand precultured placenta controls were carried out. Actin wasdetected as a positive control for protein content present insamples, due the stable expression of this protein.

GLUT1 expression was evaluated with the “Scion imagefor windows” program (version: Beta 4.0.2) to measure thearea units marked by Western blot.

2.7. Statistical Analysis. Data were expressed as the mean ±SE and were analyzed statistically by ANOVA test followed by“post hoc” LSD Fisher test. Paired t-test was performed, toestablish significance between groups (different placentas).A level of less than 5% (P < 0.05) was chosen to detectsignificant differences.

3. Results

Placental explants showed groups of the reproductive formsof the parasite (amastigotes) mainly in stromal cells ofthe chorionic villi under normal (NG) and high D-glucose(HG) concentrations, as they were seen in placental villislides stained with PAS/H. Furthermore, the presence ofthe parasite was more evident when placental explants werecultured with high D-glucose concentration (Figure 1).

Placental villi explants cultured under HG concentration(HGC) underwent structural modifications mainly thick-ening of the syncytiotrophoblast and basal membranes,increased glycogen deposits, and a fetal capillary thickening,similar to those described in the bibliography [28–30]. Thesealterations were maintained and also were more notoriouswhen these placental explants were infected with T. cruzi,compared to healthy controls incubated with normal glucoseconcentration (NGC), as shown in placental sections withPAS/H staining (Figure 1).

GLUT1 protein was intensively stained on the apical andbasal membrane of the syncytiotrophoblast in controls withNGC and no infection. The apical expression of GLUT1 wasscarce under HGC, infection with T. cruzi, or both conditionstogether, even thought the label increased in basal area underthese conditions (Figures 2 and 3).

Image quantification of immunostained areas showedthat placental explants cultured under HGC per se reducedthe expression of GLUT1 to 34.7% at 24 h (Figure 2) and the


(a) (b)

(c) (d)

(e)

∗

∗∗

EDCBA

Are

a(%

)

(f)

Figure 2: GLUT1 immunodetection in paraffin-embedded placental villi sections cultured for 24 hours. (a) Uninfected control placental villi.GLUT1 expression was intensively marked on the apical and basal syncytiotrophoblast cell membrane (arrows); (b) normal placental villi invitro infected with T. cruzi; (c) normal placental villi cultured under high glucose concentration media; (d) normal placental villi culturedunder high glucose concentration media, and in vitro infected with T. cruzi; (e) negative control placental villi (without GLUT1 antiserumincubation); (f) image quantification made by “Image Tool” UTHSCSA version 3.00, which indicates the percentage of the marked area;values shown are the mean ± SE of four samples from representative experiments (A: control placental villi, without infection; B: infectednormal placental villi; C: normal placental villi cultured under high glucose media; D: infected normal placental villi cultured under highglucose media; E: negative control placental villi). Original magnification: 400x. The scale bar represents 1 µm. ∗Significantly different fromthe control placental villi (P < 0.05; n = 4; paired t-test).

level of GLUT1 protein expression recovered to 76.3% at 48 h(Figure 3). In explants invaded by T. cruzi, the expressionof GLUT1 protein was reduced under both NG and HGconcentrations. At 24 h, the protein expression was reducedto 32% under NGC and to 14.8% under HGC. At 48 h, theGLUT1 protein expression recovered to 37.4% of the initiallevels under NGC and to 48.6% under HGC (paired t-test,P < 0.05).

Western-blotting image of one of the placental samplesas representative of the experiment is shown in Figure 4.

Placental explants cultured under HGC per se reduced theexpression of GLUT1 to 40.5% at 24 h and the level of GLUT1protein expression recovered to 56.5% at 48 h. In explantscultured with T. cruzi, the expression of GLUT1 proteinwas reduced under both NG and HG concentrations. At24 h, the protein expression was reduced to 59.5% underNGC and to 20% under HGC. At 48 h, the GLUT1 proteinexpression was at 52.1% and 14.85% of the initial levels,in cultures under NG and HG concentrations, respectively(P < 0.05).


(a) (b)

(c) (d)

(e)

∗∗

EDCBA

Are

a(%

)

(f)

Figure 3: GLUT1 immunodetection in paraffin-embedded placental villi sections cultured for 48 hours. (a) Uninfected control placentalvilli (GLUT1 expression was intensively marked on the apical and basal syncytiotrophoblast cell membrane); (b) normal placental villi invitro infected with T. cruzi; (c) normal placental villi cultured under high glucose concentration media; (d) normal placental villi culturedunder high glucose concentration media and in vitro infected with T. cruzi; (e) negative control placental villi (without GLUT1 antiserumincubation); (f) image quantification made by “Image Tool” UTHSCSA version 3.00, which indicates the percentage of the marked area;values shown are the mean ± SE of seven samples from representative experiments (A: control placental villi, without infection; B: infectednormal placental villi; C: normal placental villi cultured under high glucose media; D: infected normal placental villi cultured under highglucose concentration media; E: negative control placental villi). Original magnification: 400x. The scale bar represents 1 µm.∗Significantlydifferent from the control placental villi (P < 0.05; n = 7; paired t-test).

4. Discussion

Different authors have reported that diabetics have anincreased susceptibility to a variety of infectious agents [31,32]. Hyperglycemia has been previously observed to increasethe morbidity and mortality of murine T. cruzi infection[15]. Diabetes and hyperglycemia were also reported tobe more prevalent in chagasic human patients with thecardiac form of the disease, than in control ones [16].But, according to the analysis of the bibliography, there isnot any study analyzing an association between pregnantwomen affected with both Chagas’ disease and diabetes

with congenital transmission or with the effect on the newborn. Furthermore, there is no marker for placental or fetusinfection in Chagas’ disease. Due the effect of the Chagasparasite on some proteins located at the lipid raft of chorionicvilli trophoblast [24, 33], as well as in trophoblast lipids [34],we aimed to analyze the possible modification of the mainglucose transporter located at the syncytiotrophoblast, theGLUT1 protein, produced by placental T. cruzi infection.

T. cruzi crosses the placental barrier and infects the fetus,causing the congenital form of the disease [1]. In order tounderstand the mechanism used by the parasite to cross thisbarrier, the interaction between syncytiotrophoblast plasma


∗∗

∗

G + TGN + TNWCN

Are

a(%

)

G + TGN + TNWCN

55 kDa

Actin42 kDa

(a)

∗

∗∗

G + TGN + TN

Are

a(%

)

G + TGN + TN

55 kDa

Actin42 kDa

(b)

Figure 4: Western blot to detect GLUT1 transporter protein expression in homogenates from normal placental villi, cultured for 24 hours(a) and 48 hours (b). Actin presence was also detected as a positive control for protein content in samples. GLUT1 expression was evaluatedwith the “Scion Image for Windows” program that measured the area units marked. WCN: without culture normal placental villi (positivecontrol); N: normal placental villi; N + T: normal placental villi cocultured with T. cruzi; G: normal placental villi cultured under high glucosemedia; G + T: normal placental villi cocultured with T. cruzi under high glucose media. Values shown are the mean ± SE. ∗Significantlydifferent from the control (P < 0.05; n = 8; ANOVA test).

membranes from the human placenta and the parasite waspreviously studied, with modifications in lipid and proteinpatterns from trophoblast membranes being found [35].However, the mechanism by which the parasite infects theplacenta as well as the effects upon the protein contents ofthe placental barrier is still not well understood.

Our experiments showed that GLUT1 protein expressionwas significantly diminished in normal placental villi cul-tured under HGC in vitro infected with T. cruzi, compared tocontrols, as was observed by GLUT1 immunodetection andwestern blot. Previous studies have shown that the down-regulation of GLUT1 mRNA expression is correlated withdecreased glucose uptake in placental trophoblasts culturedunder high D-glucose concentration [22]. In the presentwork, a significant reduction in GLUT1 protein expressionwas observed in normal placental syncytiotrophoblast, cul-tured under HGC for either 24 or 48 hours. This corroboratesthe inverse relation existing between GLUT1 expression andextracellular glucose concentration, as described by variousauthors [18, 20, 36]. Our results are consistent with previousobservations that a reduction in mRNA expression of GLUT1is induced by a high glucose concentration in culturedplacental trophoblast cells [22, 36–38].

T. cruzi produces plasmatic membrane modifications, byaltering their lipid [34, 35] and protein components, suchas placental alkaline phosphatase [7–9, 39–41] and GLUT1protein, as observed in the present study. On the otherhand, we observed in our laboratory, that Gamma-glutamyltranspeptidase, another enzyme present in the placenta’sbrush border, is not affected in cells cocultured with T. cruzi[33], suggesting that the parasite affects molecules insertedin lipid microdomains of the membrane, as PLAP [42] andGLUT1 [43]. As the hyperglycemia characteristic of diabetes

mellitus also affects these membrane components, we suggestthat both the parasite and the high glucose conditions couldhave been provoking, by different ways, a lower placentaefficacy transporting glucose to fetus. The results obtainedin this work demonstrate that placental explants infectedwith T. cruzi induce changes in GLUT1 expression fromhuman term placenta under high D-glucose concentration.As a consequence, it is of importance to perform a systematicstudy analyzing new born from pregnant women who haveboth conditions diabetes and Chagas.

Reduced GLUT1 expression observed in placental culturewith parasites could imply a downregulated GLUT1 activityin pregnant women with Chagas’ disease. If this diseaseis causing damage to pancreas islets [11–13], or happenstogether with a previous diabetic condition, the adverseeffects of reduced GLUT1 activity may be exacerbated.The level of GLUT1 is regulated by glucose concentration,insulin-like growth factor I, placental growth hormone, andhypoxia [18]. It would be interesting to further study themechanisms by which GLUT1 is affected in Chagas’ disease.It is likely that the parasite would perturb other componentsof the maternal and fetal glucose-insulin-GLUT1 axis. Asglucose and GLUT are inversely regulated, changes in GLUTexpression might alter the insulin level in plasma, pointingout to determine a marker for placental infection by T. cruziin pregnant chagasic women. This matter could be of theupmost importance in congenital Chagas’ disease.

Histological studies have demonstrated that placentasfrom poorly controlled diabetic pregnancies show a thick-ening of the basal membranes and reduced vascularizationof the villi [30]. Similar to observations reported by otherauthors [28, 29], in the present work we detected thatplacental villi cultured under high glucose conditions were


morphologically altered, with thickening of the basementmembranes. The presence of T. cruzi maintains this alter-ation. The parasite causes disorganization of the basalmembrane of the trophoblast and the stroma of the chorionicvilli [44], as was observed in in vitro experiments similarto those performed in this work. So, both conditionshyperglycemia and T. cruzi can modify the stroma of thechorionic villi. According to our results, the alteration ofthe trophoblast basal membrane in the presence of T. cruziunder high D-glucose concentration might be caused by twodifferent mechanisms or not. This important matter shouldbe elucidated.

The protein membrane pattern was also altered, asnoted in others works on Placental Alkaline Phosphatase(PLAP) [7–9, 24, 25, 39–41]; and in the present studywith GLUT1 protein. These characteristics could modifythe human placenta efficacy as a barrier against infections.It has been described that chorionic villi in vitro has lowsusceptibility to infection by T. cruzi in normal D-glucoseconcentration [45, 46]. In order to clarify if high D-glucoseconcentration can increase the infection of chorionic villi byT. cruzi, it is necessary to quantify the infection by T. cruziof the placental explants under high glucose concentration,in order to analyze the susceptibility of the invasion and thereproduction of the parasite in the placenta, aspect that weare planning to do in further experiments.

With the present work, we report for the first time theeffect of T. cruzi on GLUT1 protein expression, adding it tothe increasing list of affected proteins altered by this parasite[7–9, 24, 25, 40, 41].

Acknowledgments

The authors are thankful to Dr. Patricia Paglini and Dr.Walter Rivarola for providing trypomastigotes of T. cruzi,to Miriam Rabino for technical support, and to SanatorioAllende and Hospital Nacional de Clınicas de Cordoba forproviding human term placentas. This work was supportedby grants from Secretarıa de Ciencia y Tecnologıa de laUniversidad Nacional de Cordoba, Secretarıa de Ciencia yTecnologıa de la Universidad Nacional de La Rioja, andMINCyT-Cordoba.

References

[1] A. L. Bittencourt, “Congenital Chagas disease,” American Jour-nal of Diseases of Children, vol. 130, no. 1, pp. 97–103, 1976.

[2] I. Oliveira, F. Torrico, J. Munoz, and J. Gascon, “Congenitaltransmission of Chagas disease: a clinical approach,” ExpertReview of Anti-Infective Therapy, vol. 8, no. 8, pp. 945–956,2010.

[3] M. Ming, M. Chuenkova, E. Ortega-Barria, and M. E. A.Pereira, “Mediation of Trypanosoma cruzi invasion by sialicacid on the host cell and trans-sialidase on the trypanosome,”Molecular and Biochemical Parasitology, vol. 59, no. 2, pp. 243–252, 1993.

[4] S. Schenkman, N. W. Andrews, V. Nussenzweig, and E. S.Robbins, “Trypanosoma cruzi invade a mammalian epithelialcell in a polarized manner,” Cell, vol. 55, no. 1, pp. 157–165,1988.

[5] I. Tardieux, M. H. Nathanson, and N. W. Andrews, “Role inhost cell invasion of Trypanosoma cruzi-induced cytosolic-freeCa2+ transients,” Journal of Experimental Medicine, vol. 179,no. 3, pp. 1017–1022, 1994.

[6] A. G. Todorov, M. Einicker-Lamas, S. L. de Castro, M. M.Oliveira, and A. Guilherme, “Activation of host cell phos-phatidylinositol 3-kinases by Trypanosoma cruzi infection,”Journal of Biological Chemistry, vol. 275, no. 41, pp. 32182–32186, 2000.

[7] M. J. Sartori, L. Mezzano, S. Lin, S. Munoz, and S. P. de Fabro,“Role of placental alkaline phosphatase in the internalizationof trypomatigotes of Trypanosoma cruzi into HEp2 cells,”Tropical Medicine & International Health, vol. 8, no. 9, pp. 832–839, 2003.

[8] M. J. Sartori, L. Mezzano, S. Lin, G. Repossi, and S. P. Fabro,“Cellular components and placental alkaline phosphatase inTrypanosoma cruzi infection,” Revista da Sociedade Brasileirade Medicina Tropical, vol. 38, supplement 2, pp. 87–91, 2005.

[9] L. Mezzano, M. J. Sartori, S. Lin, G. Repossi, and S. P.de Fabro, “Placental alkaline phosphatase (PLAP) study indiabetic human placental villi infected with Trypanosomacruzi,” Placenta, vol. 26, no. 1, pp. 85–92, 2005.

[10] F. Nagajyothi, L. M. Weiss, D. L. Silver et al., “Trypanosomacruzi utilizes the host low density lipoprotein receptor ininvasion,” PLoS Neglected Tropical Diseases, vol. 5, no. 2, p.e953, 2011.

[11] R. G. Long, R. H. Albuquerque, A. Prata et al., “Responseof plasma pancreatic and gastrointestinal hormones andgrowth hormone to oral and intravenous glucose and insulinhypoglycaemia in Chagas’s disease,” Gut, vol. 21, no. 9, pp.772–777, 1980.

[12] M. E. Guariento, E. Olga, A. Muscelli, and J. A. Gontijo,“Chronotropic and blood pressure response to oral glucoseload in Chagas’ disease,” Sao Paulo Medical Journal, vol. 112,no. 3, pp. 602–606, 1994.

[13] J. C. Saldanha, V. M. Dos Santos, M. A. Dos Reis, D. F. DaCunha, and V. P. A. Teixeira, “Morphologic and morphometricevaluation of pancreatic islets in chronic Chagas’ disease,”Revista do Hospital das Clinicas de Faculdade de Medicinada Universidade de Sao Paulo, vol. 56, no. 5, pp. 131–138,2001.

[14] V. M. Dos Santos, M. A. de Lima, M. Cabrine-Santos et al.,“Functional and histopathological study of the pancreas inhamsters (Mesocricetus auratus) infected and reinfected withTrypanosoma cruzi,” Parasitology Research, vol. 94, no. 2, pp.125–133, 2004.

[15] H. B. Tanowitz, B. Amole, D. Hewlett, and M. Wittner,“Trypanosoma cruzi infection in diabetic mice,” Transactionsof the Royal Society of Tropical Medicine and Hygiene, vol. 82,no. 1, pp. 90–93, 1988.

[16] V. M. Dos Santos, S. F. Da Cunha, V. P. Teixeira et al.,“Frequency of diabetes mellitus and hyperglycemia in chagasicand non-chagasic women,” Revista da Sociedade Brasileira deMedicina Tropical, vol. 32, no. 5, pp. 489–496, 1999.

[17] H. Li, Y. Gu, Y. Zhang, M. J. Lucas, and Y. Wang, “Highglucose levels down-regulate glucose transporter expressionthat correlates with increased oxidative stress in placentaltrophoblast cells in vitro,” Journal of the Society for GynecologicInvestigation, vol. 11, no. 2, pp. 75–81, 2004.

[18] M. U. Baumann, S. Deborde, and N. P. Illsley, “Placentalglucose transfer and fetal growth,” Endocrine, vol. 19, no. 1,pp. 13–22, 2002.


[19] T. Jansson, M. Wennergren, and T. L. Powell, “Placental glu-cose transport and GLUT 1 expression in insulin-dependentdiabetes,” American Journal of Obstetrics and Gynecology, vol.180, no. 1, part 1, pp. 163–168, 1999.

[20] N. P. Illsley, “Glucose transporters in the human placenta,”Placenta, vol. 21, no. 1, pp. 14–22, 2000.

[21] K. Ogura, M. Sakata, M. Yamaguchi, H. Kurachi, and Y.Murata, “High concentration of glucose decreases glucosetransporter-1 expression in mouse placenta in vitro and invivo,” Journal of Endocrinology, vol. 160, no. 3, pp. 443–452,1999.

[22] T. Hahn, S. Barth, U. Weiss, W. Mosgoeller, and G. Desoye,“Sustained hyperglycemia in vitro down-regulates the GLUT1glucose transport system of cultured human term placentaltrophoblast: a mechanism to protect fetal development?”FASEB Journal, vol. 12, no. 12, pp. 1221–1231, 1998.

[23] N. W. Andrews and W. Colli, “Adhesion and interiorization ofTrypanosoma cruzi in mammalian cells,” Journal of Protozool-ogy, vol. 29, no. 2, pp. 264–269, 1982.

[24] R. E. Fretes and S. P. de Fabro, “Trypanosoma cruzi: modifica-tion of alkaline phosphatase activity induced by trypomastig-otes in cultured human placental villi,” Revista do Instituto deMedicina Tropical de Sao Paulo, vol. 32, no. 6, pp. 403–408,1990.

[25] S. Lin, M. J. Sartori, L. Mezzano, and S. P. de Fabro, “Placentalalkalinephosphatase (PLAP) enzyme activity and binding toIgG in Chagas’ disease,” Placenta, vol. 26, no. 10, pp. 789–795,2005.

[26] O. H. Lowry, N. J. Rosebrough, A. L. Farr, and R. J. Randall,“Protein measurement with the Folin phenol reagent,” TheJournal of Biological Chemistry, vol. 193, no. 1, pp. 265–275,1951.

[27] U. K. Laemmli, “Cleavage of structural proteins during theassembly of the head of bacteriophage T4,” Nature, vol. 227,no. 5259, pp. 680–685, 1970.

[28] P. Vannini, “Pregnancy and diabetes: physiopathologicalaspects,” Minerva Endocrinologica, vol. 19, no. 2, pp. 45–50,1994.

[29] P. Garner, “Type I diabetes mellitus and pregnancy,” TheLancet, vol. 346, no. 8968, pp. 157–161, 1995.

[30] G. Desoye and E. Shafrir, “The human placenta in diabeticpregnancy,” Diabetes Reviews, vol. 4, no. 1, pp. 70–89, 1996.

[31] J. F. Plouffe, J. Silva Jr., R. Fekety, and J. L. Allen, “Cell-mediated immunity in diabetes mellitus,” Infection & Immu-nity, vol. 21, no. 2, pp. 425–429, 1978.

[32] J. Casey and C. Sturm Jr., “Impaired response of lymphocytesfrom non-insulin-dependent diabetics to staphage lysate andtetanus antigen,” Journal of Clinical Microbiology, vol. 15, no.1, pp. 109–114, 1982.

[33] S. Priotto, M. J. Sartori, G. Repossi, and M. A. Valentich,“Trypanosoma cruzi: participation of cholesterol and placentalalkaline phosphatase in the host cell invasion,” ExperimentalParasitology, vol. 122, no. 1, pp. 70–73, 2009.

[34] A. E. Fabro and G. Calzolari, “Increase of lipids in humanchagasic placentas: cytochemical and biochemical study,”Revista de la Facultad de Ciencias Medicas de la UniversidadNacional de Cordoba, vol. 48, no. 1-2, pp. 25–32, 1990.

[35] R. O. Calderon and S. P. de Fabro, “Trypanosoma cruzi:fusogenic ability of membranes from cultured epimastigotesin interaction with human syncytiotrophoblast,” ExperimentalParasitology, vol. 56, no. 2, pp. 169–179, 1983.

[36] N. P. Illsley, M. C. Sellers, and R. L. Wright, “Glycaemicregulation of glucose transporter expression and activity in thehuman placenta,” Placenta, vol. 19, no. 7, pp. 517–524, 1998.

[37] S. Hauguel-De Mouzon, M. Loizeau, and J. Girard, “Develop-mental expression of Glut1 glucose transporter and c-fos genesin human placental cells,” Placenta, vol. 15, no. 1, pp. 35–46,1994.

[38] S. Barth, T. Hahn, R. Zechner, and G. Desoye, “Prolongedhyperglycemia in vitro affects glucose transporter protein Glut1 and glucose uptake in cultured term trophoblast cells,”Placenta, vol. 15, p. 4A, 1994.

[39] R. E. Fretes and S. P. de Fabro, “Human chagasic placentallocalization of enzymes associated to syncytiotrophoblastmembrane,” Comunicaciones Biolo6gicas, vol. 9, pp. 51–59,1990.

[40] M. J. Sartori, P. Pons, L. Mezzano, S. Lin, and S. P. deFabro, “Trypanosoma cruzi infection induces microfilamentdepletion in human placenta syncytiotrophoblast,” Placenta,vol. 24, no. 7, pp. 767–771, 2003.

[41] S. Lin, M. J. Sartori, L. Mezzano, and S. P. de Fabro, “Epidermalgrowth factor (EGF) in the human placental infection withTrypanosoma cruzi,” Placenta, vol. 25, no. 4, pp. 283–286, 2004.

[42] Z. Salamon, S. Devanathan, I. D. Alves, and G. Tollin,“Plasmon-waveguide resonance studies of lateral segregationof lipids and proteins into microdomains (rafts) in solid-supported bilayers,” Journal of Biological Chemistry, vol. 280,no. 12, pp. 11175–11184, 2005.

[43] T. Sakyo, H. Naraba, H. Teraoka, and T. Kitagawa, “Theintrinsic structure of glucose transporter isoforms Glut1 andGlut3 regulates their differential distribution to detergent-resistant membrane domains in nonpolarized mammaliancells,” FEBS Journal, vol. 274, no. 11, pp. 2843–2853, 2007.

[44] J. Duaso, G. Rojo, G. Cabrera et al., “Trypanosoma cruziinduces tissue disorganization and destruction of chorionicvilli in an ex vivo infection model of human placenta,”Placenta, vol. 31, no. 8, pp. 705–711, 2010.

[45] C. D. Lujan, M. F. Triquell, A. Sembaj, C. E. Guerrero, andR. E. Fretes, “Trypanosoma cruzi: productive infection is notallowed by chorionic villous explant from normal humanplacenta in vitro,” Experimental Parasitology, vol. 108, no. 3-4,pp. 176–181, 2004.

[46] M. F. Triquell, C. Dıaz-Lujan, H. Freilij, P. Paglini, andR. E. Fretes, “Placental infection by two subpopulations ofTrypanosoma cruzi is conditioned by differential survival of theparasite in a deleterious placental medium and not by tissuereproduction,” Transactions of the Royal Society of TropicalMedicine and Hygiene, vol. 103, no. 10, pp. 1011–1018, 2009.


Research Article

Prevalence of Congenital Malaria in Minna, North Central Nigeria

Innocent Chukwuemeka James Omalu,1 Charles Mgbemena,2

Amaka Mgbemena,1 Victoria Ayanwale,1 Israel Kayode Olayemi,1

Adeniran Lateef,3 and Victoria I. Chukwuemeka1

1 Department of Biological Sciences, Federal University of Technology, Minna, Nigeria2 Dentistry Department, Niger State General Hospital, Minna 900002, Nigeria3 Department of Biochemistry/Physiology, University of Abuja, FCT, Nigeria

Correspondence should be addressed to Innocent Chukwuemeka James Omalu, omalu [email protected]

Received 14 April 2011; Revised 24 June 2011; Accepted 28 June 2011


Copyright © 2012 Innocent Chukwuemeka James Omalu et al. This is an open access article distributed under the CreativeCommons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided theoriginal work is properly cited.

The study was designed to determine the true prevalence of congenital, cord, and placental malaria in General Hospital Minna,North Central Nigeria. Peripheral blood smears of near-term pregnant women, as well as the placental, cord, and peripheral bloodsmears of their newborn babies, were examined for malaria parasites, using the Giemsa staining technique. Out of 152 pregnantwomen screened, 21 (13.82%) of them were infected with malaria parasites. Of the 152 new born babies, 4 (2.63%) showed positiveperipheral parasitaemia. Placental parasitaemia was 7/152 (4.61%), while cord blood parasitaemia was 9/152 (5.92%). There werestrong associations between peripheral and cord malaria parasitaemia and congenital malaria (P < 0.05). Plasmodium falciparumoccurred in all, and none had mixed infection. The average birth weights of the babies delivered of nonmalarious pregnant womenwere higher than those delivered by malarious pregnant women, though not significant (P > 0.05). Malaria parasitaemia occurredmore frequently in primigravidae than multigravidae.

1. Introduction

Congenital malaria was first described in 1876 [1]. It can beacquired by transmission of parasites from mother to childduring pregnancy or perinatally during labour [2].

Congenital malaria has been documented for many years,but it was previously thought to be uncommon especiallyin indigenous populations; more recent studies, however,suggest that incidence has increased, and values between0.30 to 33.00% have been observed from both endemic andnonendemic areas [3].

Malaria and pregnancy are generally believed to be mutu-ally aggravating conditions. The pathological changes due tomalaria and the physiological changes associated with preg-nancy have a synergistic effect on the course of each other[4].

Pregnancy exacerbates malaria through a nonspecifichormone-dependant depression of the immune system; pro-tective antiplasmodial activity is suppressed at pregnancy [5].

The following hypotheses to explain the altered immu-nity associated with pregnancy were offered.

Reduced lymphoproliferative response sustained by ele-vated levels of serum cortisol, loss of cell-mediated immunityin the mother, the presence of placenta, a new organ inthe primigravidae, allows the parasite to bypass the existinghost immunity, or allows placenta-specific phenotypes of P.falciparum to multiply; pregnant women display a bias to-wards type-2 cytokines and are therefore susceptible todiseases requiring type-1 responses for protection like TB,malaria, and P. falciparum has the unique ability of cytoad-hesion; chondroitin sulfate A and hyaluronic acid are theadhesion molecules for parasite attachment to placental cells[4].

In pregnancies complicated by malaria, both fetal growthretardation and preterm birth contribute to low birth weight[6]. Malaria control still remains the foremost public healthchallenge in Africa, and world malaria report, which indi-cated that Nigeria accounts for a quarter of all malaria casesin the 45 malaria endemic countries in Africa, clearly showedthe enormity of the socioeconomic and health burdens ofthe disease in the country, even in the face of dearth of


Table 1: Prevalence of Plasmodium falciparum in blood samples of different groups of women and tissues at General Hospital Minna, NorthCentral Nigeria.

Source of bloodOverall Primigravidae Multigravidae

No examdNo

positiveInfect rate

(%)No examd

Nopositive

Infect rate(%)

No examdNo

positiveInfect rate

(%)

Near-termPregnant women

152 21 13.82 76 12 15.79 76 9 11.84

Nonpregnwomen

100 13 13.00 — — — — — —

Neonate 152 4 2.63 — 2 — — 2 —

Placenta 152 7 4.61 76 3 3.95 76 4 5.26

Cord 152 9 5.92 76 4 5.26 76 5 6.58

Total 708 54 7.63 226 21 9.29 228 20 8.77

information on the epidemiology of the disease. This, thus,informed this study on the prevalence of congenital malariain Minna, an often neglected but germane aspect of theepidemiology of malaria in endemic communities [7].


2.1. Study Site and Population. The study was carried out atGeneral Hospital Minna, North Central Nigeria. Minna, thecapital of Niger State, Nigeria, is located within longitude6◦33′E and latitude 9◦37′N, covering a land area of 88 km2

with a population of 1.2 million. Minna has a tropicalclimate with mean annual temperature, relative humidity,and rainfall of 30.20◦, 61.00%, and 1334.00 cm, respectively.The climate presents two distinct seasons: a rainy season(April–October) and dry season (November–March). Minnais an endemic area for malaria.

The study was conducted from October 2010 to Febru-ary 2011. The subjects were near-term (close to delivery)pregnant women who were delivered of their babies at thehospital. Sample sizes were determined from the numberof pregnant women that attended antenatal care during theperiod of study. The nonpregnant women served as controlto help compare malaria prevalence in pregnancy only. Thepregnant women did not take any chemoprophylaxis and didnot use mosquito treated nets for protection. The HIV statusof the subjects were not determined.

2.2. Ethical Considerations. All work was performed accord-ing to the guidelines for human experimentations in clinicalresearch stated by the Federal Ministry of Health of Nigeria.This study was approved by the ethical committee of GeneralHospital Minna, Nigeria. All women gave oral informedconsent.

2.3. Sample Collections. Blood samples were obtained fromthe peripheral blood of 152 near-term pregnant womenusing sterile syringes. After delivery, blood was collectedfrom the placentae by placental biopsies. The placentae wereincised between the maternal and foetal surface and a smallquantity of blood pipette out from the intervillous with asterile syringe. Cord blood was collected immediately after

separation of placenta, and peripheral blood of neonates wascollected by heel pricking using a sterile blood lancet onclean glass slides. Labelled ethylene-diamine tetraacetic acid(EDTA) bottles were used in collecting blood samples forparasitological examinations. Blood collections were madepossible with the assistance of midwives on duty duringchild delivery. Blood samples were also collected from 100nonpregnant women.

The ages, types of birth, and weights of newborn babieswere recorded.

2.4. Parasitological Examination. Thick and thin blood filmswere stained with 10% Giemsa and read for malaria parasitesby two trained microscopists following standard quality con-trol procedure. Parasitaemia was expressed as the number ofasexual forms of P. falciparum per microlitre; a result wasconsidered negative after a reading of 1000 leucocytes in themicroscope (×1000). Parasitaemia was graded as low (1–999/µL), moderate (1000–9999/µL), and high (>10000/µL).Transplacental passage of Plasmodium falciparum was con-firmed by detection of malaria parasite within 7 days of birth.

2.5. Statistical Analysis. All data were analysed using SPSSversion 10.1 for windows. Descriptive statistics were com-puted for all relevant data. Chi square analysis was used tocompare proportions within and among groups, for statisti-cal significance.

3. Results

The prevalence of malaria infection in blood smears of near-term pregnant mothers, their newborns, cords, placentae,and nonpregnant women was shown in Table 1. Out ofa total of 152 near-term pregnant women screened, 21(13.82%) had parasite in their peripheral blood. Among 100nonpregnant women screened, 13 (13.00%) had peripheralmalaria parasite. Infection rate was higher in pregnantwomen but not significant (P > 0.05) using the chi-square.The inclusion of nonpregnant women helped to comparethe prevalence of malaria. Also, there was no significantdifference in infection rates between the primigravidae(15.79%) and the multigravidae (11.84%), (P > 0.05).


Table 2: Average parasite densities (Parasites/uL) of blood of different groups of women and tissues at General Hospital Minna, Northcentral Nigeria.

Parasite densities Pregnant women Nonpregnant women Newborn Placenta Cord PG MG

Parasites/uL 1415 411 128 689 196 1281 134

PG: Neonate with primigravid mothers,MG: Neonate with multigravid mothers.

Table 3: Erythrocytic stages of Plasmodium falciparum in blood samples of different groups of women and tissues at General HospitalMinna, North central Nigeria.

Source of blood No positiveTR GM SCH TR & SCH TR & GM TR & GM & SCH Total

No % No % No % No % No % No %

Pregnant women 21 60 64.50 25 26.80 — — — — 8 8.60 — — 93

Nonpreg women 13 15 46.80 12 37.50 — — — — 5 15.60 — — 32

Newborn 4 5 45.50 5 45.50 — — — — 1 9.00 — — 11

Placenta 7 20 62.50 10 31.20 — — — — 2 6.30 — — 32

Cord 9 12 36.30 10 30.30 — — — — 3 9.00 — — 33

Total 54 112 62 19 201

TR: trophozoites, GM: gametocytes, SCH: schizonts.

Of the 152 newborn babies screened, 4 (2.63%) had positiveperipheral parasitaemia. Examination of the 152 placentaegave a prevalence of 4.63% (7) for malaria parasite, outof which 4 placentae were for the 4 babies with positiveperipheral parasitaemia and known showed any clinicalsymptom. The only malaria parasite encountered in thisstudy was the Plasmodium falciparum.

The prevalence of cord blood parasitaemia was 5.92%(9/152), 5 of which came from multigravidae (6.58%)and 4 from primigravidae (5.26%). The average parasitedensity was 1,415 p/uL for pregnant women which wassignificantly (P < 0.05) higher than the 411 p/uL recorded fornonpregnant women. Average parasite density for newbornswas 128 p/uL, slightly lower than that for the cords whichwas 196 p/uL; however, both were significantly lower thanparasite density recorded in the placentae (mean = 689 p/µL).Parasite density was significantly (P < 0.05) higher in ne-onates from primigravid (1281/uL) mothers than multi-gravids (134/uL) (Table 2). Parasite stages encountered weredominated by trophozoites (64.50%), distantly followed bygametocytes (26.80%) in pregnant women; the same trendwas observed for nonpregnant women, placenta, cord, andneonates (Table 3).

The 4 parasitized newborns all had birth weights above3 kg. On the average, however, the birth weights of babiesfrom parasitized mothers were lower than those from non-parasitized mothers, albeit, the difference was not significant(P > 0.05). Also, newborns from primigravid mothersweighed averagely less than those from multigravid mothers;again the difference was not significant (Table 4).

4. Discussion

Many researchers have reported high prevalence of malaria inpregnancy in different parts of Nigeria, ranging from 19.70%to 72.00% [8–11]. The prevalence of malaria in pregnant

women in this study was 14%; though consistent with thereported Nigerian situation, the relatively low percentagecould be due to the fact that the study was carried out duringthe dry season, a period of low mosquito density and, per-haps, low level malaria transmission rates. This observationsupports the position that in areas of malaria endemicity,pregnancy is associated with increased susceptibility tomalaria, arising from pregnancy-induced altered immunity[6], Immunosuppression from raised serum cortisol, loss ofcell-mediated immunity, effects of a new organ, the placenta,and loss of type-1 cytokine responses [4].

The prevalence of malaria in nonpregnant controls in thisstudy was 13%, although this was lower than the pregnantsubjects; the difference was not significant. The inclusionof nonpregnant women in the study allows comparison ofmalaria prevalence in pregnancy with nonpregnant femalesubjects. Parasite densities for pregnant women were com-paratively higher than that of nonpregnant women. Thisis consistent with the findings of Agomo et al. [8], in astudy done in Lagos, Nigeria. Although none of the pregnantwomen screened took malaria chemoprophylaxis duringpregnancy, nor used any form of mosquito nets, the parasitedensity figures were low. However, one may conclude thatthis may have to do with the period of the study, Octoberto February, that is, early to mid-dry season. A seasonthat coincides with decreasing densities of female Anophelesspecies and therefore reduced inoculation with parasites.

Low parasites densities were also recorded for the placen-tae, cords, and peripheral blood of newborns.

Each placentae of the 4 parasitaemia neonates in thisstudy also had positive parasitaemia. This agrees with theposition of Chedraui et al. [6], that placental infection isa prerequisite for, but does not predict, congenital malaria.A strong correlation between placental and congenital para-sitaemia has been shown [12].


Table 4: Mean (±SD) birthweights (kg) of neonate of pregnant women at General Hospital Minna, North Central Nigeria.

Pregnant women Primigravidae Multigravidae

Male Female Male Female

Positive pregnant women 3.03± 0.5 2.83± 0.44 3.18± 0.31 2.83± 0.45

Negative pregnant women 3.01± 0.52 2.89± 0.25 3.31± 0.14 2.94± 0.14

However, an increasing trend in prevalence of congenitalmalaria has been reported recently. In a multicentre studydone at Ibadan a prevalence of 5.10% was reported inUniversity College Hospital [13]. A prevalence of 46.70%was reported in a study of 120 newborn babies at Ile-Ife,Southwestern Nigeria [14]. Also a prevalence of 13.00%was reported among 546 in-born neonates at CalabarTeaching Hospital [15]. These findings represent a new trendsince parasitaemia in peripheral blood of newborns wasconsidered rare in highly endemic areas.

Infection with Plasmodium falciparum was the only oneencountered in this study. This agrees with a similar studydone in Libreville, Gabon by Bouyou-Akotet et al. [16],wherein only Plasmodium falciparum was encountered. Italso agrees with widely accepted view that Plasmodiumfalciparum is the predominant species in Nigeria [16]. Thepredominant forms of this parasite seen were the tropho-zoites (ring stages).

The only stillbirth encountered in this study resultedfrom foetal distress. The mother of the baby was negative formalaria parasites (nonmalarious).

On the average, the birth weights of babies fromparasitaemia mothers were lower than those of babies fromnonparasitaemia mothers, but not significantly so. This dis-agrees with the widely held view that babies with parasitizedplacentae often have low birth weight. It also contradicts thefindings of Adebami et al. [17], who stated that babies ofmothers with parasitized placentae have mean birth weightsignificantly lower than babies of mothers with nonpara-sitized placentae. The generally low parasite densities foundin this study may explain the minimal effect on birthweight, as observed by Opara [2] who recorded parasitizedneonate twins with normal birth weights. Primigravidae aremore susceptible to malaria infection than multigravidaein endemic areas [18]. The findings of this study supportsthis position, albeit, the difference in prevalence was notsignificant.

Congenital malaria was previously thought to be rare,especially, in areas of malaria endemicity. Though this studyappears not to support this view, an increasing prevalence ofcongenital malaria among newborns in other areas of malariaendemicity has been observed.

Conflict of Interests

The authors declare that there is no conflict of interests.

Acknowledgments

The authors gratefully acknowledge the assistance of thetechnologists and midwives and for the use of laboratory

facilities of General Hospital Minna and the Department ofBiological Sciences, Federal University of Technology, Minna.

References

[1] S. Romand, P. Bouree, J. Gelez, B. Bader-Meunier, F. Bisaro,and J. P. Dommergues, “Congenital malaria. Infected twinsborn to an asymptomatic mother,” Presse Medicale, vol. 23, no.17, pp. 797–800, 1994.

[2] D. A. Opara, “Congenital malaria in newborn twins,” GhanaMedical Journal, vol. 44, no. 2, pp. 76–78, 2010.

[3] S. A. Sotimehin, T. I. Runsewe-Abiodun, O. T. Oladapo, O. F.Njokanma, and D. M. Olanrewaju, “Possible risk factors forcongenital malaria at a tertiary care hospital in Sagamu, OgunState, South-West Nigeria,” Journal of Tropical Pediatrics, vol.54, no. 5, pp. 313–320, 2008.

[4] B. S. Kakkilaya, “Malaria and pregnancy,” 2009, http://www.malariasite.com

[5] O. O. Okwa, “The status of malaria among pregnant women:a study in Lagos, Nigeria,” African Journal of ReproductiveHealth, vol. 7, no. 3, pp. 77–83, 2003.

[6] P. A. Chedraui, J. Daily, B. Wylie, P. F. Weller, S. M. Ramin,and V. Barss, “Overview of malaria in pregnancy,” 2009,http://www.uptodate.com.

[7] World Health Organisation, “World Malaria Report,” WHO,Geneva, Switzerland, pp. 99-101, 2008.

[8] C. O. Agomo, W. A. Oyibo, R. I. Anorlu, and P. U. Agomo,“Prevalence of malaria in pregnant women in Lagos, South-West Nigeria,” Korean Journal of Parasitology, vol. 47, no. 2,pp. 179–183, 2009.

[9] O. O. Okwa, “The status of malaria among pregnant women:a study in Lagos, Nigeria,” African Journal of ReproductiveHealth, vol. 7, no. 3, pp. 77–83, 2003.

[10] M. B. Kagu, M. B. Kawuwa, and G. B. Gadzama, “Anaemiain pregnancy: a cross-sectional study of pregnant women in aSahelian tertiary hospital in Northeastern Nigeria,” Journal ofObstetrics and Gynaecology, vol. 27, no. 7, pp. 676–679, 2007.

[11] C. J. Uneke, F. E. Iyare, P. Oke, and D. D. Duhlinska,“Assessment of malaria in pregnancy using rapid diagnostictests and its association with HIV infection and hematologicparameters in South-Eastern Nigeria,” Haematologica, vol. 93,no. 1, pp. 143–144, 2008.

[12] O. A. Oduwole, G. C. Ejezie, and M. Meremekwu, “CongenitalMalaria,” American Journal of Tropical Medicine and Hygiene,vol. 84, pp. 386–389, 2011.

[13] C. Falade, O. Mokuolu, H. Okafor et al., “Epidemiology ofcongenital malaria in Nigeria: a multi-centre study,” TropicalMedicine and International Health, vol. 12, no. 11, pp. 1279–1287, 2007.

[14] P. O. Obiajunwa, J. A. Owa, and O. O. Adeodu, “Prevalenceof congenital malaria in Ile-Ife, Nigeria,” Journal of TropicalPediatrics, vol. 51, no. 4, pp. 219–222, 2005.

[15] A. D. Ekanem, M. U. Anah, and J. J. Udo, “The prevalence ofcongenital malaria among neonates with suspected sepsis in


Calabar, Nigeria,” Tropical Doctor, vol. 38, no. 2, pp. 73–76,2008.

[16] M. K. Bouyou-Akotet, D. E. Ionete-Collard, M. Mabika-Manfoumbi et al., “Prevalence of Plasmodium falciparuminfection in pregnant women in Gabon,” Malaria Journal, vol.2, no. 1, article 1, p. 18, 2003.

[17] O. J. Adebami, J. A. Owa, G. A. Oyedeji, O. A. Oyelami,and G. O. Omoniyi-Esan, “Associations between placental andcord blood malaria infection and fetal malnutrition in anarea of malaria holoendemicity,” American Journal of TropicalMedicine and Hygiene, vol. 77, no. 2, pp. 209–213, 2007.

[18] O. A. Idowu, C. F. Mafiana, and S. Dapo, “Malaria amongpregnant women in Abeokuta, Nigeria,” Tanzania HealthResearch Bulletin, vol. 8, no. 1, pp. 28–31, 2006.

congenital transmission by protozoandownloads.hindawi.com/journals/specialissues/829362.pdf ·...

Documents