The Journal of Immunology
Human Cord Blood CD4+CD25hiRegulatory T Cells Suppress
Prenatally Acquired T Cell Responses to Plasmodium
Maria S. Mackroth,* Indu Malhotra,* Peter Mungai,* Davy Koech,†Eric Muchiri,‡
and Christopher L. King*,x
In malaria endemic regions, a fetus is often exposed in utero to Plasmodium falciparum blood-stage Ags. In some newborns, this can
result in the induction ofimmune suppression. We have previously shown these modulated immune responses to persist postnatally,
with a subsequent increase in a child’s susceptibility to infection. To test the hypothesis that this immune suppression is partially
mediated by malaria-specific regulatory T cells (Tregs) in utero, cord blood mononuclear cells (CBMC) were obtained from 44
Kenyan newborns of women with and without malaria at delivery. CD4+CD25loT cells and CD4+CD25hiFOXP3+cells (Tregs) were
enriched from CBMC. Tregfrequency and HLA-DR expression on Tregswere significantly greater for Kenyan as compared with
North American CBMC (p , 0.01). CBMC/CD4+T cells cultured with P. falciparum blood-stage Ags induced production of IFN-g,
IL-13, IL-10, and/or IL-5 in 50% of samples. Partial depletion of CD25hicells augmented the Ag-driven IFN-g production in 69%
of subjects with malaria-specific responses and revealed additional Ag-reactive lymphocytes in previously unresponsive individuals
(n = 3). Addition of Tregsto CD4+CD25locells suppressed spontaneous and malaria Ag-driven production of IFN-g in a dose-
dependent fashion, until production was completely inhibited in most subjects. In contrast, Tregsonly partially suppressed malaria-
induced Th2 cytokines. IL-10 or TGF-b did not mediate this suppression. Thus, prenatal exposure to malaria blood-stage Ags
induces Tregsthat primarily suppress Th1-type recall responses to P. falciparum blood-stage Ags. Persistence of these Tregspost-
natally could modify a child’s susceptibility to malaria infection and disease.
first pregnancy, are highly susceptible to malaria (1, 2). Malaria
during pregnancy can lead to the sequestration of Plasmodium
falciparum-infected erythrocytes in the placenta through adhesion
to molecules such as chondroitin sulfate A (3–5) and is associated
with increased risk of maternal anemia, low birth weight, growth
retardation, and premature birth (6, 7). The accumulation of
infected erythrocytes in the placenta may result in transplacental
transport of infected erythrocytes or their soluble components,
thereby exposing and sensitizing the fetal immune system to
P. falciparum Ags (8–12). The reported frequency of malaria blood-
stage–specific T and B cell responses in cord blood mononuclear
The Journal of Immunology, 2011, 186: 2780–2791.
alaria infection during pregnancy constitutes a major
public health problem in malaria-endemic regions of
the world. Pregnant women, particularly those in their
cells (CBMC) ranges from ∼5% to .70% (13–18). The con-
sequences of this prenatal exposure of the infant to P. falciparum
remain poorly understood.
Several observations indicate that some newborns may become
immune tolerant to malaria blood-stage Ags in utero. Epidemio-
are more susceptible to P. falciparum infection and demonstrate
higher parasitemia compared with offspring of women without
placental malaria (19–21). Recently, we found that a subset of
newborns of women infected with malaria during pregnancy ac-
quired an immune tolerant phenotype, which persisted into
childhood, characterized by increased IL-10 production, T cell
anergy, and failure of CBMC to produce primarily IFN-g and IL-2
in response to malaria blood-stage Ags (22). Importantly, these
same children had increased risk for malaria infection compared
with children who did not acquire this tolerant phenotype. Similar
observations have been made for other human parasitic diseases
such as lymphatic filariasis and onchocerciasis (23–25).
The mechanistic basis for this tolerant phenotype acquired
in utero remains unclear. Possible explanations include clonal
deletion of or anergy in malaria-specific T cells (26, 27) due to
impaired APC function in cord blood (28–32). Alternatively, in
utero exposure to malaria blood-stage Ags may trigger activation
and expansion of regulatory T cells (Tregs) and/or increased pro-
duction of immunomodulatory cytokines such as IL-10 or TGF-b
(33–35). Recently, several studies have identified expanded pop-
ulations of CD4+T cells capable of producing IL-10 in cord blood
from offspring of women with placental malaria when compared
with those without placental malaria (36, 37). In some of these
studies, specifically CD4+CD25hicells have been shown to be
an important source of IL-10 (36, 38). Depletion of CD4+C25hi
T cells from cord blood augmented the IFN-g production of
*Center for Global Health and Diseases, Case Western Reserve University, Cleve-
land, OH 44106;†Kenya Medical Research Institute, Nairobi, Kenya;‡Division of
Vector Borne Diseases, Ministry of Health, Nairobi, Kenya; andxVeterans Affairs
Research Service, Cleveland, OH 44106
Received for publication April 12, 2010. Accepted for publication December 21,
This work was supported by Grant AI064687 from the National Institutes of Health.
M.S.M. was partially supported by a fellowship of the German National Merit Foun-
Address correspondence and reprint requests to Maria S. Mackroth and Christo-
pher L. King, Center for Global Health and Diseases, Case Western Reserve
University, WRC 4132, 2103 Cornell Road, Cleveland, OH 44106. E-mail addresses:
email@example.com and firstname.lastname@example.org
The online version of this article contains supplemental material.
Abbreviations used in this article: CBMC, cord blood mononuclear cell; cRPMI,
complete RPMI; FMO, fluorescence minus one; MSP, merozoite surface protein;
PfP0, Plasmodium falciparum phosphoriboprotein P0; Teff, effector T cell; Treg,
regulatory T cell.
CBMC cultures stimulated with either malaria blood-stage Ags or
mitogens, suggesting an immunoregulatory function of these cells
(36–38). Further characterization indicated an expanded pop-
ulation of CD4+CD25+FOXP3+T cells after in vitro culture of
CBMC with merozoites or staphylococcal enterotoxin B among
offspring of mothers with chronic or past placental malaria (37).
These studies, however, failed to isolate and fully characterize
these Tregs. Because activated nonregulatory CD4+T cells can also
express high levels of CD25, and FOXP3 expression can be in-
duced in effector T cells (Teff) upon activation in vitro (39), it
remains unclear whether these cells are activated or directly
In the current study, we focus on the potential role of Tregsin the
fetal immune response to P. falciparum Ags and whether these
regulatory cells suppress malaria Ag-driven responses by CD4+
CD25loT cells. Newborns who have been exposed and/or sensi-
tized to malaria blood-stage Ags in utero provide a unique op-
portunity to isolate and further characterize malaria-specific Tregs
because of the large number of lymphocytes often available in
cord blood. Isolation of Tregsfrom P. falciparum malaria-infected/
exposed newborns, children, or adults has not been previously
reported. In this study, we enriched for CD4+CD25hicells, the
majority of which express the Tregmaker FOXP3, and evaluated
their ability to actively suppress both spontaneous and P. falci-
parum blood-stage Ag-specific Teffresponses in vitro. We further
evaluated the frequency, phenotype, and activation of CD4+T cell
subsets among P. falciparum-sensitized versus not sensitized
Kenyan neonates (born to women with and without malaria at
delivery) relative to the frequency and phenotype of those cells in
naive North American controls.
Materials and Methods
Mothers participating in the study delivered their children at Msambweni
District Hospital (Kwale District, Coast Province, Kenya), where perennial
P. falciparum transmission occurs. Umbilical cord blood was collected
from full-term newborns immediately after parturition and was anti-
coagulated with heparin. Additionally, maternal peripheral blood and
placental intervillous blood were obtained for malaria diagnosis as de-
scribed (15). Full-thickness placental biopsies of ∼1 cm square were
obtained and stored in 10% buffered formalin. Subsequently, the sections
were embedded in paraffin, sectioned, stained with H&E and Giemsa stain,
and examined for the presence of malaria parasites in the placenta and/or
hemozoin deposits. Control cord blood was obtained from healthy North
American newborns delivered at University Hospitals, Cleveland, OH.
Ethical approval was obtained from the Human Investigations Institutional
Review Boards of University Hospitals (Case Western Reserve University,
Cleveland, OH) and the Kenya Medical Research Institute in Nairobi.
Determination of malaria infection status
Plasmodium infections were identified via two methods: 1) blood smear;
and 2) a post-PCR oligonucleotide ligation assay. Thick and thin blood
smears were stained with 4% Giemsa for 20 min and examined under oil
immersion (original magnification 3100). DNAwas extracted from 200 ml
erythrocyte pellet obtained from fetal cord blood and 200 ml whole ma-
ternal intervillous placental blood using Qiagen DNA extraction kits
(DNeasy Kit; Qiagen). The post-PCR oligonucleotide ligation assay based
on amplification of the small subunit rRNA gene was performed as pre-
viously described (40).
Ags and mitogens
Cytokine responses to two P. falciparum blood-stage Ags, merozoite sur-
face protein (MSP)-142and P. falciparum phosphoriboprotein P0 (PfP0),
were examined. rMSP-142[3D7 allele, the most common allele in the
study population (I. Malhotra and C.L. King, unpublished observations)]
was provided by Drs. C. Long, S. Singh, and D. Narum (Malaria Vaccine
Development Unit, National Institute of Allergy and Infectious Diseases,
National Institutes of Health, Bethesda, MD). Three peptides corre-
sponding to N- and C-terminal regions of PfP0 were synthesized and
purified to 70–80% (Chiron, Clayton, Victoria, Australia). The peptides
were designated N1 (DNVGSNQMASVRKSLR; codons 33–48), N2 (SV-
RKSLRGKATILMGKNT; codons 42–59), and C1 (AKADEPKKEE-
AKKVE; codons 285–299) and correspond to T cell epitopes identified
by lymphocyte proliferation responses of immunized mice (41). PHA
(Sigma-Aldrich) or anti-CD3/28–coated T cell expander beads (Dynal)
were used as positive controls.
Isolation of mononuclear cells
CBMC were isolated within 2 h of collection by standard density gradient
centrifugation on Ficoll-Paque (Amersham Biosciences). The overall
scheme for cell preparation is shown in Fig. 1. Only freshly isolated
CBMC were used in immunomagnetic cell separation steps and cell-
culture experiments including suppression experiments. Cord blood sam-
ples from which .1.2 3 108CBMC were obtained (n = 44; shown in
Table I) underwent immunomagnetic cell separation to isolate CD4+
T cells, monocytes, and CD4+CD25hicells (Fig. 1). If .1.8 3 108CBMC
were isolated, the excess CBMC were immediately cryopreserved (n = 5).
If ,1.2 3 108CBMC were obtained, they were not used for CD4+CD25hi
enrichment, and an aliquot of CBMC were cryopreserved. Irrespective of
whether cells were used for selection of CD4+CD25hi, a small subset of
freshly isolated CBMC were resuspended at a density of 106/ml in RPMI
1640 supplemented with 10% pooled human AB serum (Sigma-Aldrich),
4 mM L-glutamine, 25 mM HEPES, and 80 mg/ml gentamicin (complete
RPMI [cRPMI]; BioWhittaker, Gathersburg, MD) and cultured with
malarial Ags to detect cytokine production and lymphocyte proliferation.
For immunomagnetic cell separation, the remaining CBMC (if .1.2 3
108) were washed and resuspended in MACS buffer (PBS, 2 mmol/l
EDTA, and 0.5% BSA).
CD25hicells were isolated by immunomagnetic positive selection using
microbeads directly conjugated to anti-CD25 Abs (Miltenyi Biotec) at 2
ml/107CBMC as previously described (42). This amount of anti-CD25 is
5-fold lower than the recommended 10 ml by the manufacturer. We did this
to ensure selection of only CD25hicells, those with high expression of
CD25. This protocol reduced the recovery of CD25hicells, but increased
enrichment of CD25hiFOXP3+cells.
After the first round of positive selection, the selected CD25hicells were
subjected to a second round of immunomagnetic separation that produced
two populations of CD25hicells: the double positively selected cells, which
we designate CD25hi++, and the cells remaining from the first round of
positive selection (i.e., those not positively selected in the second round),
which we designate CD25hi+(Fig. 1). CD4+CD25loT cells and CD14+
monocytes were then isolated from CD25hidiminished CBMC by negative
selectionusingtheIsolationKitII andtheCD4+TCell IsolationKitII(both
Miltenyi Biotec) following the manufacturer’s instruction (Fig. 1).
After separation, cell populations were washed and resuspended in
cRPMI and immediately used for cell cultures and add-back suppression
assays. All samples that underwent the above described immunomagnetic
cell separation and were used for suppression assays are listed in Table I.
In a subset of samples (n = 8), freshly isolated cells were directly stained
for flow cytometric analysis to verify the purity of isolated cell populations
(Fig. 2). CBMC contained 2–3% FOXP3+cells. After one round of
magnetic selection, this was reduced by 40–81% in CD25hidiminished
CBMC. CD25hi+showed an average enrichment of 62% for FOXP3 pos-
itivity (CD25hi+, range 59–70%; n = 8). The twice positively selected
CD25hi++were further enriched to an average of 76% for FOXP3 positivity
(CD25hi++, range 73–80%; n = 8).
Cell culture and suppression assay
Cell cultures were performed in round-bottom 96-well microtiter plates on
freshly isolated cells. CBMC and CBMC diminished in CD25hicells were
cultured at 1 3 106/ml, and 105CD4+T cells were cultured with 5 3 104
monocytes per well. CD25hi++or CD25hi+cells were added to CD4+
/monocyte cultures at different ratios (ratio CD4+/CD25hiat 1:0, 1:1, and
1:0.5) to assess suppressive activity of isolated CD25hipopulations. Ad-
ditional medium was added to wells, so that the final volume was 200 ml.
Lymphocytes were stimulated in separate cultures under the following
conditions: 1) with highly purified MSP-142(5 mg/ml, kindly provided by
Carole Long and David Narum at Malaria Vaccine Development Unit,
National Institutes of Health); 2) with PfP0 N1, N2, and C1 peptides (10
mg/ml); 3) with either anti-CD3/28 beads (one bead/five CD4+T cells) or
PHA (10 mg/ml) as a positive control; and 4) with medium alone (negative
control). Optimal concentrations had been determined in previous studies
and pilot experiments (14, 15). Neutralizing anti–IL-10 and/or anti–TGF-b
were added to a subset of samples to assess the role of immunosuppressive
cytokines (n = 12; samples were selected based on available number of
cells). Culture wells containing CD4+T cell/APCs with and without
The Journal of Immunology2781
CD25hi++or CD25hi+cells were supplemented with 1 mg/ml anti–IL-10
(JES3-9D7; BD Biosciences) and/or 1 mg/ml anti-TGF-b (clone MAB
1835; R&D Systems) based on the manufacturer’s recommendation.
Cultures were set up in triplicate where sufficient cell numbers were
available. Due to limitation on samples and number of isolated CD25hi
cells, not all tests could be carried out on all samples.
Quantification of cytokines and lymphocyte proliferation
Quantification of the cytokines IFN-g, IL-13, IL-5, IL-2, IL-6, and IL-10
was performed on culture supernatants collected at 120 h. IFN-g was
measured by ELISA. The Ab pair for cytokine capture and detection
(biotinylated) was as follows: M-700A and M-701B (Endogen, Cam-
bridge, MA). IL-5, IL-10, IL-2, IL-6, and IL-13 were measured using
a bead-based multicytokine immunoassay (Upstate Luminex kit) following
the manufacturer’s instruction. The lower limit of detection for the various
cytokines that were evaluated ranged from 5–10 pg/ml depending on the
cytokine (5 pg/ml for IL-5 and 10 pg/ml for IL-10, IL-2, IL-6, and IL-13).
A positive response was scored when the following criterion was fulfilled:
for both CBMC and CD4+T cell cultures, an Ag-driven cytokine pro-
duction that was at least 2-fold greater than that of parallel cultures con-
taining medium alone. If cytokine production was not detectable in the
negative control cultures, $20 pg/ml for the Ag-specific cytokine pro-
duction was considered to be a positive response. No P. falciparum Ag-
driven cytokine response was detected in test cultures of CBMC from 16
healthy North American newborns.
Lymphocyte proliferation was performed as previously described (14).
Samples were performed in triplicate. A positive response was a stimula-
tion index (cpm in test sample/cpm in cultures with medium alone) .2.
Flow cytometric analysis
To evaluate the purity of selected cells, isolated cell populations (CBMC,
CD25hidiminished CBMC, CD25hi++, CD25hi+,CD4+T cells, and
monocytes) from eight donors were washed in FACS buffer (PBS, 2%
FBS, and 0.09% sodium azide) directly after cell separation and surface-
stained with anti-CD25 (clone MA-251) and anti-CD4 (clone SK3) (both
from BD Biosciences). For FOXP3 expression analysis, intranuclear stain-
ing was conducted using the anti-human FOXP3 staining kit according to
manufacturer’s instruction (clone PCH101; eBioscience). Monocyte pop-
ulations were stained with CD14 (clone M5E2; BD Biosciences). Stained
cells were refrigerated at 4˚C and read within 12 h on a four-color flow
cytometer (FACS scan with additional second diode to allow detection of
allophycocyanin staining) at Coast Province General Hospital in Mom-
basa, Kenya. Assessment of purity could not be undertaken on all the
samples due to limited cell numbers, in particular of the CD25hipop-
ulations, and very limited access to a flow cytometer in Kenya.
For a broader characterization of CD4+T cells and Tregsurface mole-
cules and FOXP3 expression by flow cytometry, liquid nitrogen-stored
Kenyan CBMC were transferred to the United States. Based on cytokine
responses in cell culture assays and availability of sufficient frozen Kenyan
CBMC, 17 samples were selected and grouped as sensitized (n = 9) and not
sensitized (n = 8). Of the selected Kenyan CBMC, 5 samples represented
the same individuals used for CD25hienrichment experiments and sup-
pression assays (numbers 31, 37, 38, 42, and 44, Table I, Supplemental
Table I), and 12 samples were chosen from additional frozen CBMC
samples from the same population collected in the same time period
(Supplemental Table I, samples F1–F12). In addition, nine frozen North
American CBMC samples were included in experiments.
Cryopreserved cells were quick-thawed, washed with PBS, and in-
cubated in cRPMI at 37˚C for 2 h. Cells were then washed in FACS buffer
and surface stained with the following Abs: CD4 (SK3), CTLA-4 (BNI3)
(both from BD Biosciences), CD3 (UCHT1), CD45RO (UCHL1), CD25
(BC96), CD127 (eBioRDR5), HLA-DR (LN3), CD62L (DREG56) (all
from eBioescience) and intracellularly stained for FOXP3 (clone PCH101;
eBioscience). Flow cytometry was performed on an LSRII flow cytometer
(BD Biosciences) and analyzed using FlowJo software (Tree Star). Un-
stained cells and single stained cells/beads were included in all experi-
ments. Cells were first gated based on forward and side scatter to exclude
dead cells and cell debris. For characterization of CD4+T cells and
Tregs, lymphocytes were first gated for CD3+CD4+cells (Fig. 3). The gate
in the right panel of Fig. 3 shows designation for CD25+CD127locells.
CD4+T cells and CD4+CD25+CD127locells were further analyzed for
expression of CD45RO, CTLA-4, HLA-DR, and CD62L. Fluorescence
minus one (FMO) controls were used to define the gates. It is a staining
control that employs all reagents used in a flow cytometry assay except for
one fluorochrome of interest (termed FMO) to control for the contribution
of spectral overlap to the background when using multiple fluorochromes
The significance of differences among three groups was assessed using
Kruskal-Wallis testing with Dunn’s posttesting (GraphPad Software v4.0,
GraphPad). For paired comparisons between two groups, Student t test was
performed on log-transformed data. Correlation analysis was conducted
using Pearson’s rank-correlation test.
The criterion for including individual samples from the study
population for functional assays was the ability to obtain sufficient
number of lymphocytes (.1.2 3 108CBMC) for CD25hipurifi-
cation (n = 44; Table I; see Figs. 1 and 2 for CD25hipurification).
Mothers of these offspring were primarily primigravid (22 out of
42, 52%) and secundigravid (10 out of 42, 24%) women. Of 36
mother/infant pairs tested, 8 (22%) women and/or their newborns
showed evidence of active or prior malaria infection during
pregnancy as determined by blood smear, PCR, and/or placental
microscopy at delivery. Twenty-three of 44 (52%) CBMC samples
showed recall responses to one or both malaria blood-stage Ags
based on lymphocyte proliferation and/or cytokine production
(IFN-g, IL-13, IL-5, IL-6, IL-2, and/or IL-10; Table I, section 1).
There was a mixture of Th1- and Th2-type cytokine production by
cord blood lymphocytes in response to malaria blood-stage Ags.
Sixteen out of 42 samples tested for IFN-g (38%) produced IFN-g
in response to P. falciparum Ags, and 15 of 32 (46%) showed an
IL-13 response (Table I, section 1). Four CBMC samples produced
IL-5 (numbers 18, 34, 38, and 40). Cytokine concentrations mea-
sured for IL-5 were low, between 15 and 44 pg/ml. Of note, we
were unable to detect malaria Ag-induced IL-2 at 120 h due to its
consumption in 5-d cultures. Only one sample (number 38) pro-
duced IL-6. Kenyan newborns whose samples showed $1 positive
cytokine recall response and/or lymphocyte proliferation to P.
falciparum Ags were subsequently classified as sensitized (Table I,
section 1). The majority of sensitized samples (16 out of 23)
produced either multiple cytokines to P. falciparum Ag(s) or pro-
duced one cytokine in response to both Ags (i.e., samples 21, 39,
and 40, Table I, section 1).
Of note, 6 of 8 (75%) CBMC samples from malaria-infected
women demonstrated recall responses to malaria blood-stage
Ags, whereas CBMC from 13 of 36 (36%) malaria not-infected
women also had fetal priming to malaria Ags indicative of prior
Higher numbers of Tregsare observed in cord blood of offspring
sensitized or exposed to malaria Ags in utero
To determine whether CBMC from Kenyan newborns sensitized to
malarial Ags in utero have increased frequency of memory T cells
or lymphocytes with a Tregphenotype, we did the following: first,
we classified newborns as sensitized or not sensitized based on
cell culture secretion of cytokines in response to P. falciparum
Ags. A sample was classified as sensitized if at least one positive
cytokine response (IFN-g, IL-13, IL-5, and/or IL-10) was mea-
sured. Newborns were classified as not sensitized if none of the
cytokine measurements were positive in response to MSP-1 and
PfP0 (Supplemental Table I). We then examined expression of
Treg, memory, and activation markers by flow cytometry for: 1)
Kenyan newborns sensitized to malaria (n = 9); 2) Kenyan new-
borns not sensitized to malaria (n = 8); and 3) healthy North
American newborns (n = 9).
As expected the overall frequency of CD4+T cells that
blood, 10–12%, and was similar among the three groups (Supple-
2782 CORD BLOOD REGULATORY T CELLS TO MALARIA
memory phenotype (CD45RO+CD62Lhi). There was also no dif-
ference in HLA-DR and CTLA-4 expression on CD4+T cells
among the three groups (Supplemental Table II).
To evaluate the relative proportion of Tregsamong the three
groups, gated CD4+T cells were further gated for a subpopulation
associated with a Tregphenotype, CD25+CD127lo(gating schema
shown in Fig. 3) (45–48).
Although FOXP3 is a more robust marker of Tregs, we found
intranuclear staining of FOXP3 technically difficult to perform
simultaneously with some of our chosen surface markers, such as
CTLA-4, CD45RO, and HLA-DR. Therefore, to validate the asso-
ciation of CD25+CD127lowith FOXP3+cells, lymphocytes from
nine cord blood samples were examined with a reduced panel that
included CD4, CD25, CD127, and FOXP3.
FOXP3+cells were mainly found within the CD25+CD127lo
population. Whereas 69–80% (n = 9) of CD4+T cells staining for
CD25+CD127lowere FOXP3+, only 1 to 2% of CD4+CD252cells
and 4–6% of CD4+CD25+CD127+cells expressed FOXP3 (see
Supplemental Fig. 1 for one representative sample). Conversely,
72–83% of CD4+FOXP3+T cells were CD25+CD127lo(data not
shown). The percentages of FOXP3 and CD25+CD127lostaining
CD4+cells were therefore highly correlated (r2= 0.84, p ,
0.0001; Supplemental Fig. 1) in CBMC. Together, this suggests
that cell populations expressing the markers FOXP3 and CD25+
CD127loare very similar and representative of Tregs.
Kenyan newborns sensitized in utero to malaria blood-stage Ags
had an average of 4.7% of their CD4+cells expressing CD25+
CD127locompared with an average of 3.7% and 3.1% of cord
Table I. Study subjects of cell separation and suppression experiments
Presence of Malaria Infection Lymphocyte Sensitization to P. falciparum Blood-Stage Ags (net pg/ml)
CB Placental BiopsyIFN-g
IL-13IL-10 LP (SI) Ag Response
Samples that underwent magnetic bead separation and showed cytokine response to P. falciparum Ags (sensitized samples)
Remaining samples that underwent magnetic bead separation
Maternal parity, presence of malaria infection, and cytokine production by CBMC to P. falciparum blood-stage Ag from all study subjects undergoing magnetic bead
separation and suppression experiments.
aSample identification numbers correspond to the chronology of sample acquisition.
b2 indicates values considered to be background or zero values.
cSamples 31, 37, 38, 42, and 44 were also included in flow cytometry experiments to further characterize T cell phenotype (see Supplemental Table I).
CB, cord blood; IVPB, intervillous placental blood; LP, lymphocyte proliferation; ND, not done; SI, stimulation index.
The Journal of Immunology2783
blood CD4+cells from not sensitized newborns or unexposed
North American newborns, respectively (Fig. 4A). The frequency
of CD25+CD127locells in the malaria-sensitized group was sig-
nificantly greater compared with North Americans (p , 0.01;
Fig. 4A). Significantly more CD25+CD127locells from Kenyan
newborns expressed HLA-DR+(0.95%) compared with North
American controls (0.32%, p , 0.01; Fig. 4B), indicating greater
activation or expansion of these Tregsin Kenyan newborns (49).
By contrast, there was no difference in CTLA-4 expression among
the three groups (mean percentage was 0.24, 0.23, and 0.2% for
Kenyan-sensitized, not sensitized, and North Americans, respec-
tively) nor for the memory effector cell phenotype CD45RO+
CD62Lhi(mean percentage was 16, 16.5, and 14.5%) or for the
central memory phenotype CD45RO+CD62Llo(mean percentage
was 7.2, 7.7, and 9.6%).
Effect of CD25hidepletion on recall responses to malaria
blood-stage Ags by CBMC
To assess whether CD4+CD25hiT cells suppress malaria Ag-
induced cytokine production by CBMC, we partially depleted
CD25hiT cells from CBMC using a single round of immuno-
magnetic selection in subjects shown in Table I. Examination of
FOXP3+CD25hiin CBMC before and after depletion (CBMC
versus CD25hidiminished CBMC) showed an average reduction
of 64% (range 40–81%, n = 8; Fig. 2). Partial depletion of CD4+
CD25hiaugmented the net malaria blood-stage Ag-driven IFN-g
in 11 of 16 malaria-sensitized subjects (previously shown malaria
Ag-specific IFN-g recall response), whereas in the remaining 5
subjects, there was a decrease or no change in IFN-g release (Fig.
5A). Among the 26 subjects that were not identified as sensitized,
partial depletion of CD25hicells resulted in detection of Ag-driven
IFN-g production in three subjects (Fig. 5B), indicating that the
failure to observe Ag-driven cytokine production in some cord
blood cells may result from active suppression through Tregs. In
contrast to IFN-g, partial depletion of CD4+CD25hiresulted in
a significant decrease in IL-13 production for seven subjects, no
change in four, and an increase in four subjects (data not shown).
Comparatively fewer subjects showed malaria Ag-driven IL-5 and
IL-10 (Table I). CD4+CD25hicell depletion produced no consis-
tent effect on increased or decreased IL-5 or IL-10 production
(data not shown). Thus, CD25hicells have a variable effect on
modulating Ag-driven cytokine production in CBMC, with a
generally suppressive effect on malaria Ag-driven Th1-like cyto-
kine production, but not on Th2-type cytokine production (i.e., IL-
13 and IL-5).
In cultures with measurable spontaneous cytokine release,
partial depletion of CD25hicells increased IFN-g levels by 2.2–
13.5-fold in 6 subjects, decreased by .50% in 5 subjects, and
showed no change in the remaining 31 subjects. By contrast, no
tion schema. The flow chart presents an overview of the
experimental setup as well as the overall scheme for
CBMC cell separation and enrichment of CD25hicells.
Freshly isolated CBMC were used for immunomag-
netic bead separation. The first round of enrichment
produced CD25hicells that underwent a second round
of separation, generating two different populations: the
twice positively selected CD25hi++cells and the
remaining cells, referred to as CD25hi+. CD4+T cells
and monocytes were further enriched from CD25hidi-
minished CBMC as described in the Materials and
Methods section. Flow cytometry was performed on
a subset of these isolated cells to demonstrate purity
and FOXP3 positivity (n = 8). CBMC and isolated
lymphocyte fractions were set up for cell culture and
add-back experiments. Cryopreserved CBMC were
later used for flow cytometric analysis of regulatory
cells, memory, and activation marker expression.
Experimental overview and cell separa-
plots show FOXP3+and CD25hiexpression on one representative sample
of CBMC, CBMC diminished in CD25hi, CD25hi++, and CD25hi+cells
after gating for CD4+T cells. Cells were first gated based on forward and
side scatter to exclude dead cells and cell debris and then gated for CD4+
T cells based on side scatter and expression of CD4. Gates for FOXP3 and
CD25 were based on FMO controls. Total of 4.4% of CD4+cells in CBMC
expressed FOXP3, which was reduced by ∼50% in CD25hidiminished
CBMC after one round of CD25 selection. Total of 80% of the CD4+cells
in the CD25hi++population were FOXP3+compared with 69% in the
Cell separation and Tregisolation. The flow cytometry dot
2784CORD BLOOD REGULATORY T CELLS TO MALARIA
subject demonstrated a .2-fold change in spontaneous IL-13, IL-
5, and IL-10 production following partial removal of CD25hicells
(data not shown).
Suppressive effects of enriched CD4+CD25hiTregson
Ag-driven cytokine production by CD4+CD25loT cells
To directly evaluate the ability of CD4+CD25hiTregsto suppress the
production of cytokines by malaria Ag-specific T cells from cord
blood, we first isolated CD25hilymphocytes immunomagnetically.
We adopted a strategy in which CD25hicells were positively selected
twice using anti-CD25–coated beads. After two rounds of immu-
nomagnetic separation, we obtained two populations: CD25hi++and
CD25hi+cells (Fig. 1). A total of 75–85% of the selected cells were
CD4+. In the CD25hi++population, 78–85% of the CD4+cells were
FOXP3+(Fig. 2). The CD25hi+population (positively selected in the
first round of selection, but not retained magnetically in the second
round) showed lower levels of purity, with ∼68–75% of CD4+cells
Fig. 2 shows the CD25 and FOXP3 expression of a representa-
tive sample of CD25hi++and CD25hi+populations, both gated for
CD4+cells. Of note, preliminary studies of enriched CD25hi++
show low expression of CD127 (data not shown). Cell purity of
selected monocytes and CD4+T cells was consistently .90% and
.93%, respectively. Cell recovery was generally low, typically
between 3 and 8 3 105CD25hi++cells from a total .1.2 3 108
CBMC. Enriched CD25hi++cells (cultured with CD14 positively
selected monocytes, ratio of 2:1 for CD25hi++to monocytes; n = 5)
had a phenotype characteristic of Tregs(50); they failed to pro-
liferate or produce IL-2 in response to PHA compared with strong
were first gated based on forward and side scatter to
exclude dead cells and cell debris and then gated for
CD4+T cells based on expression of CD4 and CD3.
CD4+T cells were further gated for CD25+CD127lo
cells as shown in the right panel.
CD25+CD127logating schema. Cells
CD127locells is increased in Kenyan neonates. Figures present the per-
centage of CD4+CD25+CD127locells (A) and CD4+CD25+CD127locells
expressing the activation marker HLA-DR (B) in CD4+T cells from P.
falciparum-sensitized and not sensitized Kenyan and North American
newborns. Kruskal-Wallis test with Dunn’s posttest comparisons were used
to assess the significance of differences (shown in figure). The overall p
value that includes all three groups for A is p = 0.0083 and for B is p =
Frequency and HLA-DR expression of CD4+CD25+
falciparum Ags in some individuals. The effect of partial depletion of
CD25hicells on net malaria blood-stage Ag-driven IFN-g production from
Kenyan CBMC sensitized (A) or not sensitized (B) as evaluated in Table I
is shown. The dashed lines represent the geometric mean.
Depletion of CD25hicells enhances IFN-g responses to P.
The Journal of Immunology2785
proliferation responses of similarly cultured CD4+CD25loT cells
(data not shown). Enriched CD4+CD25hi++lymphocytes did, how-
ever, spontaneously produce variable amounts of IL-10; four out
of seven individuals tested produced from 98–627 pg/ml of IL-10,
and one subject secreted 643 pg/ml IL-6. There was no sponta-
neous production of IL-5, IL-13, or IFN-g.
To evaluate the suppressive capacity of enriched CD25hi++on
CD4+CD25loT cells (subsequently referred to as CD4+T cells),
CD4+T cells were cocultured with monocytes alone as APCs or
with the addition of an equal number of enriched CD25hi++cells
(e.g., a 1:1 ratio plus APCs). Tregswere added to cultures in the
absence (spontaneous) or presence of malaria blood-stage Ag (Fig.
6). Fifteen subjects’ CBMC had detectable spontaneous IFN-g
production, of which nine (60%) showed complete suppression by
addition of Tregs, four showed partial suppression, and two showed
increased IFN-g production (Fig. 6A; p = 0.01). We next examined
the effect of adding Tregsto cultures in the presence of malaria
blood-stage Ags (Fig. 6B, 6C). Fig. 6B illustrates an experiment
from one CBMC sample. Of note, because there were insufficient
numbers of highly enriched cells to add at a lower ratio, less
highly enriched CD25hi+cells (obtained after a single round of
positive selection) were added to CD4+cells at 1:1 and 0.5:1 ratio.
Often the addition of CD25hi++resulted in complete suppression
of malaria Ag-driven IFN-g, for which levels of suppression
decreased in a dose-dependent fashion with the less enriched
CD25hi+cells added (Fig. 6B, 6C). Fig. 6C summarizes the results
of suppressive effects of Tregson all CBMC showing malaria Ag-
induced IFN-g production. Complete suppression of Ag-driven
IFN-g was observed in 12 of 16 subjects, and 3 had partial sup-
pression. Overall, the addition of CD25hi++produced a mean av-
erage suppression of 85% for malaria Ag-driven IFN-g production
by CD4+cells (p , 0.001), and with lower numbers of Tregs
(CD25hi+), the suppression was 70% (p , 0.05).
The effect of CD25hi++on suppression of Ag driven IL-13 re-
lease was less pronounced than that observed for IFN-g (Fig. 7).
Addition of CD25hi++to CD4+cells in the absence of malaria Ag
failed to suppress spontaneous IL-13 release (CD4+alone, geo-
metric mean = 10, and CD4+with CD25hi++geometric mean = 24;
Fig. 7A). Fig. 7B shows an experiment from one CBMC sample.
In all subjects with malaria Ag-driven IL-13, 4 of 15 subjects had
complete inhibition, 1 showed no inhibition, 1 individual dem-
onstrated an increase in IL-13, and the remaining 9 individuals
showed partial suppression (Fig. 7C). The overall mean level of
suppression was 61% with highly enriched CD25hi++(p , 0.05).
Addition of less enriched CD25hi+cells failed to induce significant
IL-13 suppression (p . 0.05). Although not shown, similar results
were observed for malaria Ag-induced IL-5. Thus, CD25hi++
T cells showed weaker suppression of Th2-type cytokine pro-
duction compared with Th1-type responses. Of note, IL-10 pro-
duction by CD4+T cells was not suppressed by adding CD25hi++
cells (data not shown); rather the addition augmented IL-10 pro-
duction in 5 of 6 subjects showing an IL-10 response to malaria
Ags (data not shown).
Suppression of CD4+T cells by Tregsis not dependent on IL-10
or TGF-b production
Because some isolated CD25hi++cells from Kenyan newborns
produced IL-10, and IL-10 and TGF-b are known to mediate
suppression of T cells responses (36, 51–53), we evaluated
whether these cytokines from Tregsor other IL-10/TGF-b–pro-
ducing CD4+cells may contribute to the observed immune sup-
pression. The addition of neutralizing anti–IL-10 and/or anti–
TGF-b to cultures failed to significantly augment spontaneous
IFN-g (Fig. 8A; n = 12, p = 0.14) or IL-13 production (data not
shown). Addition of CD25hi++Tregssuppressed spontaneous IFN-
g production by CD4+cells and continued to suppress CD4+cells’
spontaneous IFN-g production after addition of anti–IL-10/TGF-b
(Fig. 8A; p = 0.03). With respect to Ag-driven cytokine pro-
duction, neutralizing Abs to IL-10– and/or TGF-b–augmented
malaria Ag-driven IFN-g in some individuals (Fig. 8B; represen-
tative of 2 out of 12 studied) but not others (Fig. 8C; representative
of 2 out of 12; the remaining 8 out of 12 did not show any Ag-
CD25hicells on suppression of spontaneous (A) and malaria Ag-driven IFN-g (B, C). B shows an experiment with CBMC from newborn number 3 (Table I)
in which CD25hi++and CD25hi+cells are enriched and added back to purified CD4+T cells with APCs. This subject was shown because sufficient CD25hi+
cells were available for add back at several ratios. Cultures were performed in triplicate unless there were insufficient cells, and values represent mean + SE.
C summarizes the results of subjects for which there was detectable net malaria Ag-driven IFN-g. Each point represents mean net (Ag-induced minus
spontaneous) cytokine production for responders. Lines connect CBMC from the same individual for the three culture conditions indicated. The condition
23 CD4+/APC was included as an additional control condition. Subsequent add-back experiments are to 13 CD4 T cells. Dashed lines (A) and bars (C) in
the figures show the geometric mean cytokine production for all individuals. Kruskal-Wallis test with Dunn’s posttest comparisons were used to assess the
significance of differences (shown in figure). The overall p values that include all three groups for C is p = 0.0002.
Cord blood CD25hiTregssuppress spontaneous and P. falciparum-Ag–specific IFN-g response. The figures illustrate the effect of enriched
2786CORD BLOOD REGULATORY T CELLS TO MALARIA
driven cytokine production above background). Importantly, the
blocking of endogenous IL-10 and/or TGF-b failed to reverse the
suppressive effect of CD4+CD25hi++cells on malaria Ag-driven
IFN-g (Fig. 8B, 8C) or IL-13 (data not shown).
In humans and other primates a unique maternal–fetal interface
develops where the placenta of fetal origin is in direct contact with
maternal blood (i.e., hemochorial placenta development). This
allows for efficient gas and nutrient exchange and the trans-
placental transfer of Abs from maternal to fetal circulation.
However, this physiology requires development of immunologic
mechanisms whereby the semiallogeneic fetus escapes recognition
by the maternal immune system. This requirement provides for the
expression on the placenta of nonclassical MHC class I HLA-G,
which stimulates inhibitory receptors on cells of lymphoid and
myelomonocytic origin (54, 55). It also selects for the production
of immune inhibitory cytokines, PGs, and immunoregulatory
T cells (56, 57). The increasing study of CD4+CD25+FOXP3 Tregs
in cord blood has shown they are typically immature and not
activated compared with those from adult blood, yet they appear
to be more suppressive (42, 58–61). When fetal Tregsare exposed
to nonself-Ags in utero, however, they become critical in the
regulation of maternal cell interaction in the fetus, and these cells
persist well into childhood (62). Thus, the introduction of exog-
enous Ags, such as malarial products, into this potent fetal regu-
latory milieu may heighten stimulation of Ag-specific Tregs. In this
study, we show the presence of malaria blood-stage Ag-specific
Tregs suppress P. falciparum-Ag–
specific, but not spontaneous IL-13
response. Enriched CD25hicells fail
to suppress spontaneous IL-13 re-
lease from CD4+CD25lo T cells (A)
but CD25hi++do suppress malaria
Ag-driven IL-13 production (B, one
individual) and for all malaria-Ag–
reactive individuals that produced IL-
13 (C). Panels are identical to those
described in legend of Fig. 6 with the
exception that B shows results from
subject 38 in Table I. The overall p
value (for the three groups) for C is
p = 0.023, and comparison for degree
of suppression of CD4+C25hi++on
CD4+CD25locells is partial and sig-
nificant at p , 0.05 level using the
same analysis described in the legend
of Fig. 6.
Cord blood CD25hi
and malarial Ag-induced cytokine production were examined with and without neutralizing anti–IL-10 and anti–TGF-b. A shows the effects of neutralizing
anti–IL-10/anti–TGF-b on spontaneous IFN-g production and suppression by CD25hi++Tregs(n = 12). Each symbol point represents spontaneous cytokine
production for one individual. Bars in A show the geometric mean of spontaneous IFN-g production for all study samples in this experiment (n = 12). The p
values were determined using paired t test of log-transformed data. B and C, Graphs present IFN-g production by T cells plus or minus Tregsand/or
neutralizing anti–IL-10/anti–TGF-b as indicated on the x-axis. B shows results from cultures induced with PfP0 from individual 32 and C from cultures
induced with MSP-1 using CBMC from newborn 44 in Table I. Cultures were performed in triplicate unless there were insufficient cells, and values
represent mean + SE.
Anti–IL-10/anti–TGF-b do not abrogate suppression by CD25hiTregs. Suppressive effects of CD25hi++Tregson spontaneous cytokine release
The Journal of Immunology2787
CD4+CD25hiFOXP3 Tregs in cord blood from newborns of
malaria-infected or malaria-exposed women. These cells, when
purified, are directly suppressive of CD4+CD25loT cells in a dose-
dependent fashion. This suppression is much more potent for Th1
than for Th2-type cytokine production by cord blood cells, con-
sistent with generally Th2 cytokine bias of cord blood lympho-
cytes (18, 63, 64). IL-10 or TGF-b do not mediate this Treg
suppression, consistent with some but not all prior studies (36, 38).
This study differed from prior studies examining Tregs in
malaria-infected individuals or in cord blood from mothers ex-
posed to malaria in that we directly purified CD4+CD25hiregu-
latory cells. The large amount of lymphocytes collected in some of
our cord blood samples allowed such studies, which cannot be
routinely performed on peripheral blood samples, particularly in
young children. We undertook two rounds of positive selection
using a lower concentration of anti-CD25 beads than recom-
mended by the manufacturer, with the aim of preferentially
enriching for CD4+T cells that expressed very high levels of
CD25. These cells are most strongly associated with a Tregphe-
notype as determined by the presence of FOXP3. As a conse-
quence of this stringent selection, the number of CD4+CD25hi
cells recovered was low. This and practical obstacles in the field
led to the limitation that not all of the enriched samples were
checked for the purity of FOXP3+cells (those checked did show
limited variability, however, with 78–85% of CD4+cells being
FOXP3+). As a consequence, the observed differences in level of
suppression with add back among samples may have resulted from
differences in FOXP3+enrichment. Another limitation of this
purification is that not all Tregshave been depleted from whole
CBMC. This may account for the failure to augment Ag-driven
responses in some subjects following partial depletion. Only
a subset of CBMC diminished in CD25hiwas measured for the
amount of CD4+CD25hidepletion, which was highly variable due
to the lower number of beads used to remove CD4+CD25hi. There-
fore, we were unable to accurately associate depletion of CD4+
CD25hiwith the magnitude of Ag-driven IFN-g.
The enriched CD4+CD25hicells had functional characteristics
of Tregs. They failed to proliferate or to generate IL-2 with mito-
gen stimulation and produced variable amounts of IL-10 (47). Of
note, retention of functional suppression following enrichment of
CD25hiT cells required the use of fresh CBMC and a larger
number of available CBMC. This requirement, along with the
limited number of enriched cells, precluded detailed phenotypic
analysis. Therefore, we did not correlate functional activity with
phenotypic expression, a limitation of the study. Enriched Tregs
suppressed both spontaneous and malaria blood-stage Ag-driven
IFN-g production in a dose-dependent fashion. The observation
that enriched Tregssuppressed spontaneous IFN-g in the absence
of additional exogenous Ag suggests preactivation to exogenous
Ag in utero or by the process of positive selection. However, some
enriched Tregsthat showed only partial suppression in spontaneous
cultures completely suppressed Ag-induced IFN-g production
(Fig. 6), suggesting that a subset of Tregsis malaria Ag-specific.
Furthermore, not all enriched Tregssuppressed spontaneous or Ag-
induced IFN-g, supporting the notion of in utero activation in
some infants and not others.
Enriched cord blood Tregswere much more effective in sup-
pressing Th1-type as compared with Th2-type cytokine pro-
duction. Enriched Tregsconsistently failed to inhibit spontaneous
IL-13 release, whereas in the same cultures, IFN-g production was
completely suppressed. Similarly, enriched Tregs partially sup-
pressed Ag-induced IL-13 production, but to a lesser extent than
that of IFN-g in the same cultures and only at a ratio of CD4+
CD25hi++to CD4+CD25loof 1:1. Although much data supports the
observation that Tregscan suppress Th2-type responses (65, 66),
this varies depending on the culture conditions used. Prior studies
indicate that enriched CD25+Tregsshow defective suppression of
Th2-type cytokines to birch pollen, but only during the birch
pollen season (67). Similarly, Th2 clones were less susceptible to
suppression by human thymocyte-derived CD25+Tregscompared
with Th1 clones (68). The addition of IL-4 or IL-9 could further
reduce the suppressive capacity on Th2 cells, but not Th1 lines,
supporting the interpretation that Th1 clones respond to primarily
IL-2, whereas Th2 cells can respond to other growth factors such
as IL-4 and IL-9. This is consistent with one possible mechanism
of Tregssuppression, in which high expression of CD25, or IL-2R,
depletes cultures of IL-2 necessary for cell activation and growth,
especially for Th1-type cells, whereas Th2-type cells can respond
to other growth factors. This may occur under conditions that
produce greater amounts of these additional growth factors (e.g.,
IL-4 and IL-9), such as during allergy season or in the fetal en-
vironment (66, 69, 70). This observation is consistent with our
prior observations that newborns who develop a tolerant pheno-
type in utero show persistent suppression of malaria-specific Th1,
but little suppression of malaria-specific Th2-type responses in
exposed fetuses (14, 15, 17, 38), which are thought to be important
for immunoregulation (36, 38, 71). This can occur both sponta-
neously and in response to malaria blood-stage Ags, suggesting
that expansion of IL-10–producing T cells may be important for
modulating malaria Ag-specific immune responses. In malaria-
exposed neonates, CD4+CD25hicells appear to be a source of IL-
10 (36, 38), and IL-10 has been identified as a key mediator of Treg
function (along with TGF-b and IL-35); the extent to which these
cytokines mediate suppression by Tregsappears to vary greatly in
different pathogenic/hemostatic settings (72). The present studies
suggest that IL-10 or TGF-b are not important mediators of CD4+
CD25hiFOXP3+Treg-induced suppression by cord blood following
malaria exposure, suggesting that other mechanisms may be in-
volved, such as metabolic disruption of Teffby consumption of
locally produced IL-2 as mentioned above or by targeting dendritic
cells for suppression. Our observations do show, however, an im-
munoregulatory role of IL-10 whereby adding anti–IL-10 aug-
mented IFN-g production in previously unresponsive individuals,
which is consistent with previous studies (71). This phenomenon is
likely mediated through other Tregsubsets.
Two recent studies in adults have shown that FOXP3+Tregsare
expanded in infected versus noninfected adults as well as exposed
versus nonexposed adults (73, 74). Similar to these recent studies
in adults, previous studies with cord blood have examined whether
Tregsare more likely to be obtained from mothers who have evi-
dence of current or prior malaria. The history of prenatal exposure
to malaria has been typically surmised by the presence of malaria
in the placenta at the time of delivery, either by direct detection of
parasites (PCR, blood smear) or histologically. The detection of
hemozoin (malaria pigment) in the placenta indicates prior in-
fection, resulting in classification of placental malaria as acute,
chronic, or past infection (no evidence of active infection at de-
livery). Whether this histological classification accurately reflects
prenatal exposure to malaria is unknown. Using this criterion,
however, one study demonstrated CD4+CD25hiTregs are more
prevalent in cord blood from offspring of women with placental
malaria (36). A second study showed an expansion of CD4+
CD25+FOXP3+cells only after in vitro stimulation with mer-
ozoites or staphylococcal enterotoxin B in offspring of women
with chronic or past but not active malaria (37). A third study
found no association with placental malaria and Tregs in cord
2788CORD BLOOD REGULATORY T CELLS TO MALARIA
blood (38), similar to the lack of association with malaria in-
fection in women for the current study. Because lack of evi-
dence of malaria in the placenta or peripheral blood at delivery
does not exclude prior malaria exposure, a better negative control
is cord blood cells from newborns living in an area not endemic
for malaria. Using this control, we found the proportion of CD4+
cells expressing a Tregphenotype was 30–40% higher in cord
blood from Kenyans (many of whom have been exposed to
malaria), as compared with cord blood from North Americans.
More striking is the finding that Tregsfrom Kenyan CBMC were 3-
fold more likely to express the activation marker HLA-DR com-
pared with Tregsfrom North American newborns. By contrast, we
found no difference in markers on Tregsfrom Kenyan and North
American newborns with respect to the memory marker CD45RO,
the immunoregulatory marker CTLA-4, or markers suggestive
of effector memory (CD45RO+CD62L2) or central memory
(CD45RO+CD62L+) cells. Together, these observations suggest
activation and expansion of Tregsin Kenyan newborns exposed to
malaria and other parasite Ags in utero. It is likely that only
a small subset of Tregsare strongly activated at any point, espe-
cially to malaria, and they may be difficult to detect by flow
In contrast to other studies, we also classified newborns with
malaria exposure in utero based on whether they developed recall
have developed at any time during pregnancy, even if the mother
was found to be negative for malaria at delivery. Using this
classification, there was a trend toward greater numbers and fre-
quency of activated Tregsin cord blood from malaria-sensitized
versus nonsensitized children; however, the numbers were too
small to show significant differences. This is not surprising for
three reasons. First, the nonsensitized offspring may include
a putatively tolerant group (i.e., prior malaria exposure) that may
have expanded Tregs, but lack a conventional Ag recall response
(20). Second, using only two purified Ags, MSP-1, and PfP0 in our
experiments, it is likely that not all potentially sensitized offspring
were detected. Third, Tregsmay also have been expanded and
activated in response to other Ags in utero. Pregnant women living
in malaria endemic areas are often coinfected with various hel-
minth, bacterial, and viral infections that can stimulate immune
responses in the fetus.
In our study, suppressive Tregscould be isolated from most, but
not all newborns, and the suppressive capacity varied among
individuals. This observed variation may be related more to dif-
ferences in purification than real functional differences. Addi-
tionally, the proportion of Tregsthat were malaria specific was
difficult to assess. Once Tregsare activated either to malaria or
other Ags, however, they can suppress nonspecifically (75, 76). It
unlikely that Tregdifferences in cord blood could have arisen from
maternal contamination because we have previously shown in our
population significant admixture occurs infrequently (10).
What is clear from this and other studies is that Tregsare ex-
panded and activated in cord blood from newborns living in
malaria endemic areas, either as a consequence of in utero ex-
posure to malaria or to Ags of chronic blood-borne infections
found in pregnant women in these areas. A better understanding of
Tregsfunction in utero will require the use of more accurate bio-
markers for their presence and function, as well as a better way to
correlate timing of malaria infection during pregnancy.
Recent studies show that Tregs can modify susceptibility to
disease (77, 78) and contribute to whether the host immune
responses are protective or pathological in response to infection
with P. falciparum parasites (73, 79–82). The nature of the initial
exposure to malaria Ags likely affects the potentially diverse roles
assumed by Tregsin malaria infection. For some individuals, this
first experience appears to occur in utero (10, 12, 14, 17, 83, 84).
This may have an important impact on the subsequent de-
velopment of an individual’s immune response to malaria and
potentially to other Ags. How this prenatal exposure shapes the
subsequent immune response is only now beginning to be studied.
The current and several previous studies indicate that the gener-
ation of Tregsis an important component of the response; however,
further study of factors that determine how Tregsare generated in
utero, how they function, and whether they persist into infancy
and childhood as a reservoir of preactivated regulatory cells is
needed. Such studies are important, as increasing efforts are made
to intensively control malaria during pregnancy, such as with
prophylactic drugs and through immunization programs once an
effective malaria vaccine emerges. The subsequent lack of expo-
sure to malarial Ags in utero will surely affect malarial morbidity
and mortality in childhood, but in ways that we are only beginning
Charles NgaNga for technical help in conduction of the immunological
assays. We also thank the maternity nurses at Msambweni Hospital for help
with collection of cord blood and the women residing in the Msambweni
area for participation in the study.
The authors have no financial conflicts of interest.
1. Steketee, R. W., B. L. Nahlen, M. E. Parise, and C. Menendez. 2001. The burden
of malaria in pregnancy in malaria-endemic areas. Am. J. Trop. Med. Hyg. 64(1-
2, Suppl): 28–35.
a new look at the numbers. Am. J. Trop. Med. Hyg. 64(1-2, Suppl): iv–vii.
3. Fried, M., and P. E. Duffy. 1996. Adherence of Plasmodium falciparum to
chondroitin sulfate A in the human placenta. Science 272: 1502–1504.
4. Duffy, P. E., and M. Fried. 2003. Plasmodium falciparum adhesion in the pla-
centa. Curr. Opin. Microbiol. 6: 371–376.
5. Maubert, B., N. Fievet, G. Tami, C. Boudin, and P. Deloron. 2000. Cytoadher-
ence of Plasmodium falciparum-infected erythrocytes in the human placenta.
Parasite Immunol. 22: 191–199.
6. Uneke, C. J. 2007. Impact of placental Plasmodium falciparum malaria on
pregnancy and perinatal outcome in sub-Saharan Africa: II: effects of placental
malaria on perinatal outcome; malaria and HIV. Yale J. Biol. Med. 80: 95–103.
7. Desai, M., F. O. ter Kuile, F. Nosten, R. McGready, K. Asamoa, B. Brabin, and
R. D. Newman. 2007. Epidemiology and burden of malaria in pregnancy. Lancet
Infect. Dis. 7: 93–104.
8. Jakobsen, P. H., F. N. Rasheed, J. N. Bulmer, M. Theisen, R. G. Ridley, and
falciparum-infected women and high concentrations of soluble E-selectin and
a circulating P. falciparum protein in the cord sera. Immunology 93: 264–269.
9. Malhotra, I., A. Dent, P. Mungai, E. Muchiri, and C. L. King. 2005. Real-time
quantitative PCR for determining the burden of Plasmodium falciparum para-
sites during pregnancy and infancy. J. Clin. Microbiol. 43: 3630–3635.
10. Malhotra, I., P. Mungai, E. Muchiri, J. J. Kwiek, S. R. Meshnick, and C. L. King.
2006. Umbilical cord-blood infections with Plasmodium falciparum malaria are
acquired antenatally in Kenya. J. Infect. Dis. 194: 176–183.
11. Falade, C., O. Mokuolu, H. Okafor, A. Orogade, A. Falade, O. Adedoyin,
T. Oguonu, M. Aisha, D. H. Hamer, and M. V. Callahan. 2007. Epidemiology of
congenital malaria in Nigeria: a multi-centre study. Trop. Med. Int. Health 12:
12. Redd, S. C., J. J. Wirima, R. W. Steketee, J. G. Breman, and D. L. Heymann.
1996. Transplacental transmission of Plasmodium falciparum in rural Malawi.
Am. J. Trop. Med. Hyg. 55(1, Suppl)57–60.
13. Fievet, N., P. Ringwald, J. Bickii, B. Dubois, B. Maubert, J. Y. Le Hesran,
M. Cot, and P. Deloron. 1996. Malaria cellular immune responses in neonates
from Cameroon. Parasite Immunol. 18: 483–490.
14. Malhotra, I., A. N. Wamachi, P. L. Mungai, E. Mzungu, D. Koech, E. Muchiri,
A. M. Moormann, andC. L.
neonatal lymphocytes to an abundant malaria blood-stage antigen: epitope
mapping of Plasmodium falciparum MSP1(33). J. Immunol. 180: 3383–3390.
15. Malhotra, I., P. Mungai, E. Muchiri, J. Ouma, S. Sharma, J. W. Kazura, and
C. L. King. 2005. Distinct Th1- and Th2-Type prenatal cytokine responses to
Plasmodium falciparum erythrocyte invasion ligands. Infect. Immun. 73: 3462–
King. 2008.Finespecificity of
The Journal of Immunology2789
16. King, C. L., I. Malhotra, A. Wamachi, J. Kioko, P. Mungai, S. A. Wahab,
D. Koech, P. Zimmerman, J. Ouma, and J. W. Kazura. 2002. Acquired immune
responses to Plasmodium falciparum merozoite surface protein-1 in the human
fetus. J. Immunol. 168: 356–364.
17. Brustoski, K., M. Kramer, U. Mo ¨ller, P. G. Kremsner, and A. J. Luty. 2005.
Neonatal and maternal immunological responses to conserved epitopes within
the DBL-gamma3 chondroitin sulfate A-binding domain of Plasmodium falci-
parum erythrocyte membrane protein 1. Infect. Immun. 73: 7988–7995.
18. Metenou, S., A. L. Suguitan, Jr., C. Long, R. G. Leke, and D. W. Taylor. 2007.
Fetal immune responses to Plasmodium falciparum antigens in a malaria-
endemic region of Cameroon. J. Immunol. 178: 2770–2777.
19. Mutabingwa, T. K., M. C. Bolla, J. L. Li, G. J. Domingo, X. Li, M. Fried, and
P. E. Duffy. 2005. Maternal malaria and gravidity interact to modify infant
susceptibility to malaria. PLoS Med. 2: e407.
20. Schwarz, N. G., A. A. Adegnika, L. P. Breitling, J. Gabor, S. T. Agnandji,
R. D. Newman, B. Lell, S. Issifou, M. Yazdanbakhsh, A. J. Luty, et al. 2008.
Placental malaria increases malaria risk in the first 30 months of life. Clin. Infect.
Dis. 47: 1017–1025.
21. Le Hesran, J. Y., M. Cot, P. Personne, N. Fievet, B. Dubois, M. Beyeme ´,
C. Boudin, and P. Deloron. 1997. Maternal placental infection with Plasmodium
falciparum and malaria morbidity during the first 2 years of life. Am. J. Epi-
demiol. 146: 826–831.
22. Malhotra, I., A. Dent, P. Mungai, A. Wamachi, J. H. Ouma, D. L. Narum,
E. Muchiri, D. J. Tisch, and C. L. King. 2009. Can prenatal malaria exposure
produce an immune tolerant phenotype? A prospective birth cohort study in
Kenya. PLoS Med. 6: e1000116.
23. Elson, L. H., A. Days, M. Calvopin ˜a, W. Paredes, E. Araujo, R. H. Guderian,
J. E. Bradley, and T. B. Nutman. 1996. In utero exposure to Onchocerca vol-
vulus: relationship to subsequent infection intensity and cellular immune re-
sponsiveness. Infect. Immun. 64: 5061–5065.
24. Malhotra, I., P. L. Mungai, A. N. Wamachi, D. Tisch, J. M. Kioko, J. H. Ouma,
E. Muchiri, J. W. Kazura, and C. L. King. 2006. Prenatal T cell immunity to
Wuchereria bancrofti and its effect on filarial immunity and infection suscepti-
bility during childhood. J. Infect. Dis. 193: 1005–1013.
25. Malhotra, I., J. H. Ouma, A. Wamachi, J. Kioko, P. Mungai, M. Njzovu,
J. W. Kazura, and C. L. King. 2003. Influence of maternal filariasis on childhood
infection and immunity to Wuchereria bancrofti in Kenya. Infect. Immun. 71:
26. Gammon, G., K. Dunn, N. Shastri, A. Oki, S. Wilbur, and E. E. Sercarz. 1986.
Neonatal T-cell tolerance to minimal immunogenic peptides is caused by clonal
inactivation. Nature 319: 413–415.
27. Gammon, G. M., A. Oki, N. Shastri, and E. E. Sercarz. 1986. Induction of
tolerance to one determinant on a synthetic peptide does not affect the response
to a second linked determinant. Implications for the mechanism of neonatal
tolerance induction. J. Exp. Med. 164: 667–672.
28. Schwartz, R. H. 2003. T cell anergy. Annu. Rev. Immunol. 21: 305–334.
29. Taylor, S., and Y. J. Bryson. 1985. Impaired production of gamma-interferon by
newborn cells in vitro is due to a functionally immature macrophage. J. Immunol.
30. Mahnke, K., J. Knop, and A. H. Enk. 2003. Induction of tolerogenic DCs: ‘you
are what you eat’. Trends Immunol. 24: 646–651.
31. Mahnke, K., Y. Qian, J. Knop, and A. H. Enk. 2003. Induction of CD4+/CD25+
regulatory T cells by targeting of antigens to immature dendritic cells. Blood
32. Jonuleit, H., E. Schmitt, G. Schuler, J. Knop, and A. H. Enk. 2000. Induction of
interleukin 10-producing, nonproliferating CD4(+) T cells with regulatory
properties by repetitive stimulation with allogeneic immature human dendritic
cells. J. Exp. Med. 192: 1213–1222.
33. Rieger, M., and I. Hilgert. 1977. The involvement of a suppressor mechanism in
neonatally induced allograft tolerance in mice. J. Immunogenet. 4: 61–67.
34. Fernandez, M. A., F. K. Puttur, Y. M. Wang, W. Howden, S. I. Alexander, and
C. A. Jones. 2008. T regulatory cells contribute to the attenuated primary CD8+
and CD4+ T cell responses to herpes simplex virus type 2 in neonatal mice. J.
Immunol. 180: 1556–1564.
35. Rainsford, E., and D. J. Reen. 2002. Interleukin 10, produced in abundance by
human newborn T cells, may be the regulator of increased tolerance associated
with cord blood stem cell transplantation. Br. J. Haematol. 116: 702–709.
36. Brustoski, K., U. Moller, M. Kramer, F. C. Hartgers, P. G. Kremsner, U. Krzych,
and A. J. Luty. 2006. Reduced cord blood immune effector-cell responsiveness
mediated by CD4+ cells induced in utero as a consequence of placental Plas-
modium falciparum infection. J. Infect. Dis. 193: 146–154.
37. Flanagan, K. L., A. Halliday, S. Burl, K. Landgraf, Y. J. Jagne, F. Noho-Konteh,
J. Townend, D. J. Miles, M. van der Sande, H. Whittle, and S. Rowland-Jones.
2010. The effect of placental malaria infection on cord blood and maternal
immunoregulatory responses at birth. Eur. J. Immunol. 40: 1062–1072.
38. Bisseye, C., M. van der Sande, W. D. Morgan, A. A. Holder, M. Pinder, and
J. Ismaili. 2009. Plasmodium falciparum infection of the placenta impacts on the
T helper type 1 (Th1)/Th2 balance of neonatal T cells through CD4(+)CD25(+)
forkhead box P3(+) regulatory T cells and interleukin-10. Clin. Exp. Immunol.
39. Allan, S. E., S. Q. Crome, N. K. Crellin, L. Passerini, T. S. Steiner, R. Bacchetta,
M. G. Roncarolo, and M. K. Levings. 2007. Activation-induced FOXP3 in hu-
man T effector cells does not suppress proliferation or cytokine production. Int.
Immunol. 19: 345–354.
40. Mehlotra, R. K., K. Lorry, W. Kastens, S. M. Miller, M. P. Alpers, M. Bockarie,
J. W. Kazura, and P. A. Zimmerman. 2000. Random distribution of mixed species
malaria infections in Papua New Guinea. Am. J. Trop. Med. Hyg. 62: 225–231.
41. Chatterjee, S., S. Singh, R. Sohoni, V. Kattige, C. Deshpande, S. Chiplunkar,
N. Kumar, and S. Sharma. 2000. Characterization of domains of the phosphor-
iboprotein P0 of Plasmodium falciparum. Mol. Biochem. Parasitol. 107: 143–
42. Godfrey, W. R., D. J. Spoden, Y. G. Ge, S. R. Baker, B. Liu, B. L. Levine,
C. H. June, B. R. Blazar, and S. B. Porter. 2005. Cord blood CD4(+)CD25
(+)-derived T regulatory cell lines express FoxP3 protein and manifest potent
suppressor function. Blood 105: 750–758.
43. Roederer, M. 2001. Spectral compensation for flow cytometry: visualization
artifacts, limitations, and caveats. Cytometry 45: 194–205.
44. Roederer, M. 2002. Compensation in flow cytometry. Curr. Protoc. Cytom.
Chapter 1: Unit 1.14.
45. Liu, W., A. L. Putnam, Z. Xu-Yu, G. L. Szot, M. R. Lee, S. Zhu, P. A. Gottlieb,
P. Kapranov, T. R. Gingeras, B. Fazekas de St Groth, et al. 2006. CD127 ex-
pression inversely correlates with FoxP3 and suppressive function of human
CD4+ T reg cells. J. Exp. Med. 203: 1701–1711.
46. Seddiki, N., B. Santner-Nanan, J. Martinson, J. Zaunders, S. Sasson, A. Landay,
M. Solomon, W. Selby, S. I. Alexander, R. Nanan, et al. 2006. Expression of
interleukin (IL)-2 and IL-7 receptors discriminates between human regulatory
and activated T cells. J. Exp. Med. 203: 1693–1700.
47. Shevach, E. M. 2001. Certified professionals: CD4(+)CD25(+) suppressor
T cells. J. Exp. Med. 193: F41–F46.
48. Baecher-Allan, C., J. A. Brown, G. J. Freeman, and D. A. Hafler. 2001. CD4+
CD25high regulatory cells in human peripheral blood. J. Immunol. 167: 1245–
49. Baecher-Allan, C., E. Wolf, and D. A. Hafler. 2006. MHC class II expression
identifies functionally distinct human regulatory T cells. J. Immunol. 176: 4622–
50. Stephens, L. A., C. Mottet, D. Mason, and F. Powrie. 2001. Human CD4(+)
CD25(+) thymocytes and peripheral T cells have immune suppressive activity
in vitro. Eur. J. Immunol. 31: 1247–1254.
51. Belkaid, Y. 2008. Role of Foxp3-positive regulatory T cells during infection. Eur.
J. Immunol. 38: 918–921.
52. Maynard, C. L., and C. T. Weaver. 2008. Diversity in the contribution of
interleukin-10 to T-cell-mediated immune regulation. Immunol. Rev. 226: 219–
53. Wohlfert, E., and Y. Belkaid. 2008. Role of endogenous and induced regulatory
T cells during infections. J. Clin. Immunol. 28: 707–715.
54. Hunt, J. S., and D. L. Langat. 2009. HLA-G: a human pregnancy-related im-
munomodulator. Curr. Opin. Pharmacol. 9: 462–469.
55. Hunt, J. S., D. K. Langat, R. H. McIntire, and P. J. Morales. 2006. The role of
HLA-G in human pregnancy. Reprod. Biol. Endocrinol. 4(Suppl 1): S10.
56. Koch, C. A., and J. L. Platt. 2007. T cell recognition and immunity in the fetus
and mother. Cell. Immunol. 248: 12–17.
57. Trowsdale, J., and A. G. Betz. 2006. Mother’s little helpers: mechanisms of
maternal-fetal tolerance. Nat. Immunol. 7: 241–246.
58. Wing, K., A. Ekmark, H. Karlsson, A. Rudin, and E. Suri-Payer. 2002. Char-
acterization of human CD25+ CD4+ T cells in thymus, cord and adult blood.
Immunology 106: 190–199.
59. Cupedo, T., M. Nagasawa, K. Weijer, B. Blom, and H. Spits. 2005. Development
and activation of regulatory T cells in the human fetus. Eur. J. Immunol. 35: 383–
60. Wing, K., P. Larsson, K. Sandstro ¨m, S. B. Lundin, E. Suri-Payer, and A. Rudin.
2005. CD4+ CD25+ FOXP3+ regulatory T cells from human thymus and cord
blood suppress antigen-specific T cell responses. Immunology 115: 516–525.
61. Takahata, Y., A. Nomura, H. Takada, S. Ohga, K. Furuno, S. Hikino,
H. Nakayama, S. Sakaguchi, and T. Hara. 2004. CD25+CD4+ T cells in human
cord blood: an immunoregulatory subset with naive phenotype and specific
expression of forkhead box p3 (Foxp3) gene. Exp. Hematol. 32: 622–629.
62. Mold, J. E., J. Michae ¨lsson, T. D. Burt, M. O. Muench, K. P. Beckerman,
M. P. Busch, T. H. Lee, D. F. Nixon, and J. M. McCune. 2008. Maternal
alloantigens promote the development of tolerogenic fetal regulatory T cells in
utero. Science 322: 1562–1565.
63. Zaghouani, H., C. M. Hoeman, and B. Adkins. 2009. Neonatal immunity: faulty
T-helpers and the shortcomings of dendritic cells. Trends Immunol. 30: 585–591.
64. Forsthuber, T., H. C. Yip, and P. V. Lehmann. 1996. Induction of TH1 and TH2
immunity in neonatal mice. Science 271: 1728–1730.
65. Ozdemir, C., M. Akdis, and C. A. Akdis. 2009. T regulatory cells and their
counterparts: masters of immune regulation. Clin. Exp. Allergy 39: 626–639.
66. Pace, L., C. Pioli, and G. Doria. 2005. IL-4 modulation of CD4+CD25+
T regulatory cell-mediated suppression. J. Immunol. 174: 7645–7653.
67. Grindebacke, H., K. Wing, A. C. Andersson, E. Suri-Payer, S. Rak, and
A. Rudin. 2004. Defective suppression of Th2 cytokines by CD4CD25 regula-
tory T cells in birch allergics during birch pollen season. Clin. Exp. Allergy 34:
68. Cosmi, L., F. Liotta, R. Angeli, B. Mazzinghi, V. Santarlasci, R. Manetti,
L. Lasagni, V. Vanini, P. Romagnani, E. Maggi, et al. 2004. Th2 cells are less
susceptible than Th1 cells to the suppressive activity of CD25+ regulatory
thymocytes because of their responsiveness to different cytokines. Blood 103:
69. Pace, L., S. Rizzo, C. Palombi, F. Brombacher, and G. Doria. 2006. Cutting
edge: IL-4-induced protection of CD4+CD25- Th cells from CD4+CD25+
regulatory T cell-mediated suppression. J. Immunol. 176: 3900–3904.
70. Scheffold, A., J. Hu ¨hn, and T. Ho ¨fer. 2005. Regulation of CD4+CD25+ regu-
latory T cell activity: it takes (IL-)two to tango. Eur. J. Immunol. 35: 1336–1341.
71. Brustoski, K., U. Mo ¨ller, M. Kramer, A. Petelski, S. Brenner, D. R. Palmer,
M. Bongartz, P. G. Kremsner, A. J. Luty, and U. Krzych. 2005. IFN-gamma and
2790CORD BLOOD REGULATORY T CELLS TO MALARIA
IL-10 mediate parasite-specific immune responses of cord blood cells induced by
pregnancy-associated Plasmodium falciparum malaria. J. Immunol. 174: 1738–
72. Vignali, D. A., L. W. Collison, and C. J. Workman. 2008. How regulatory T cells
work. Nat. Rev. Immunol. 8: 523–532.
73. Minigo, G., T. Woodberry, K. A. Piera, E. Salwati, E. Tjitra, E. Kenangalem,
R. N. Price, C. R. Engwerda, N. M. Anstey, and M. Plebanski. 2009. Parasite-
dependent expansionofTNFreceptorII-positiveregulatoryTcellswith enhanced
suppressive activity in adults with severe malaria. PLoS Pathog. 5: e1000402.
74. Finney, O. C., D. Nwakanma, D. J. Conway, M. Walther, and E. M. Riley. 2009.
Homeostatic regulation of Teffector to Treg ratios in an area of seasonal malaria
transmission. Eur. J. Immunol. 39: 1288–1300.
75. Shevach, E. M., R. A. DiPaolo, J. Andersson, D. M. Zhao, G. L. Stephens, and
A. M. Thornton. 2006. The lifestyle of naturally occurring CD4+ CD25+ Foxp3+
regulatory T cells. Immunol. Rev. 212: 60–73.
76. Jonuleit, H., E. Schmitt, M. Stassen, A. Tuettenberg, J. Knop, and A. H. Enk.
2001. Identification and functional characterization of human CD4(+)CD25(+)
T cells with regulatory properties isolated from peripheral blood. J. Exp. Med.
77. Torcia, M. G., V. Santarlasci, L. Cosmi, A. Clemente, L. Maggi, V. D. Mangano,
F. Verra, G. Bancone, I. Nebie, B. S. Sirima, et al. 2008. Functional deficit of
T regulatory cells in Fulani, an ethnic group with low susceptibility to Plas-
modium falciparum malaria. Proc. Natl. Acad. Sci. USA 105: 646–651.
78. Todryk, S. M., P. Bejon, T. Mwangi, M. Plebanski, B. Urban, K. Marsh,
A. V. Hill, and K. L. Flanagan. 2008. Correlation of memory T cell responses
against TRAP with protection from clinical malaria, and CD4 CD25 high T cells
with susceptibility in Kenyans. PLoS ONE 3: e2027.
79. Scholzen, A., G. Minigo, and M. Plebanski. 2010. Heroes or villains? T regu-
latory cells in malaria infection. Trends Parasitol. 26: 16–25.
80. Walther, M., D. Jeffries, O. C. Finney, M. Njie, A. Ebonyi, S. Deininger,
E. Lawrence, A. Ngwa-Amambua, S. Jayasooriya, I. H. Cheeseman, et al. 2009.
Distinct roles for FOXP3 and FOXP3 CD4 T cells in regulating cellular im-
munity to uncomplicated and severe Plasmodium falciparum malaria. PLoS
Pathog. 5: e1000364.
81. Hansen, D. S., and L. Schofield. 2010. Natural regulatory T cells in malaria: host
or parasite allies? PLoS Pathog. 6: e1000771.
82. Finney, O. C., E. M. Riley, and M. Walther. 2010. Regulatory T cells in malaria
—friend or foe? Trends Immunol. 31: 63–70.
83. Perrault, S. D., J. Hajek, K. Zhong, S. O. Owino, M. Sichangi, G. Smith,
Y. P. Shi, J. M. Moore, and K. C. Kain. 2009. Human immunodeficiency virus
co-infection increases placental parasite density and transplacental malaria
transmission in Western Kenya. Am. J. Trop. Med. Hyg. 80: 119–125.
84. Tena-Toma ´s, C., M. K. Bouyou-Akotet, E. Kendjo, M. Kombila, P. G. Kremsner,
and J. F. Kun. 2007. Prenatal immune responses to Plasmodium falciparum
erythrocyte membrane protein 1 DBL-alpha domain in Gabon. Parasitol. Res.
The Journal of Immunology 2791