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Systemic Inflammatory Priming in Normal
Pregnancy and Preeclampsia: The Role of
Circulating Syncytiotrophoblast
Microparticles
Sarah J. Germain, Gavin P. Sacks, Suren R. Soorana, Ian L.
Sargent and Christopher W. Redman
J Immunol 2007; 178:5949-5956; ;
doi: 10.4049/jimmunol.178.9.5949
http://www.jimmunol.org/content/178/9/5949
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References
The Journal of Immunology
Systemic Inflammatory Priming in Normal Pregnancy and
Preeclampsia: The Role of Circulating Syncytiotrophoblast
Microparticles1
Sarah J. Germain,* Gavin P. Sacks,* Suren R. Soorana,† Ian L. Sargent,*
and Christopher W. Redman2*
I
t has been argued that the human fetus is a natural allograft,
at risk of T cell-dependent alloimmune rejection (1). We
have presented an alternative view that is based on the presence of two fetal-maternal immune interfaces in human pregnancy
(2). In early pregnancy, interface I comprises a localized tissue
interaction in the decidua between maternal NK cells and invasive,
fetal extravillous cytotrophoblast. The syncytiotrophoblast at the
surface of the hemochorial placenta forms interface II in contact
with maternal blood. Interface I regresses in the second half of
pregnancy with loss of the invasive trophoblast (3) and the associated decidual lymphocytes (4). Interface II is activated with onset
of the uteroplacental circulation at 8 –9 wk (5) and enlarges with
placental growth to become the main maternal-fetal immune interface after 20 wk of pregnancy.
Our work has suggested that in later pregnancy the dominant
immune interaction at interface II involves the innate rather than
the adaptive systems and is characterized by a mild systemic inflammatory state (6, 7). This is evident in activation of circulating
leukocytes (6), the associated acute phase response (reviewed by
*Nuffield Department of Obstetrics and Gynaecology, University of Oxford, John
Radcliffe Hospital, Oxford, United Kingdom; and †Department of Maternal Fetal
Medicine, Imperial College School of Medicine, Chelsea and Westminster Hospital,
London, United Kingdom
Redman and Sargent in Ref. 8) and the parallel endothelial activation (9). We have shown that pre-eclampsia is associated with
more exaggerated systemic inflammatory changes (6) and argued
that pre-eclampsia is what happens when the response is so intense
as to provoke the features of the disorder (7).
Pre-eclampsia is a specific disorder of the second half of pregnancy, comprising variable combinations of signs of which hypertension and proteinuria are emphasized for clinical diagnosis. The
possible range of features is however much wider including dysfunction of clotting and of the liver. The condition can progress to
crises which can be fatal. These include convulsions and the acute
hemolysis, elevated liver enzymes, and low platelets (HELLP)
syndrome,3 which is characterized by disseminated intravascular
coagulation, acute hemolysis, and liver damage (10).
The cause of the inflammatory response is not known. Some
workers have assumed that alloimmune reactivity to the fetoplacental unit (11) is possible although the concept has evolved in
relation to murine rather than human pregnancy. There is little specific evidence for a human equivalent. Inflammatory responses are not
Ag specific and our current hypothesis is that one or more factors
derived from the placenta are the stimuli, which does not depend on,
or evolve into, Ag-specific immunity. The relevant issues are the intensity and quality of the maternal inflammatory response and its relation to the maternal syndrome of pre-eclampsia.
Received for publication August 30, 2006. Accepted for publication January 29, 2007.
The costs of publication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked advertisement in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
1
This project was supported by Action Research Grant SP3441 and by the Oxford
Radcliffe Hospitals Trust Research and Development Fund.
2
Address correspondence and reprint requests to Dr. Christopher W. Redman, Nuffield Department of Obstetrics and Gynaecology, John Radcliffe Hospital, Oxford,
U.K. E-mail address: [email protected]
www.jimmunol.org
3
Abbreviations used in this paper: HELLP syndrome, acute hemolysis, elevated liver
enzymes, and low platelets; STBM, general term for syncytiotrophoblast microparticles; mSTBM, STMsB prepared by a mechanical method; pSTBM, STMsB
prepared by perfusion of the maternal surface of the placenta; MsIgG, mouse IgG
isotype Ig.
Copyright © 2007 by The American Association of Immunologists, Inc. 0022-1767/07/$2.00
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Systemic inflammatory responsiveness was studied in normal human pregnancy and its specific inflammatory disorder, pre-eclampsia.
Compared with nonpregnancy, monocytes were primed to produce more TNF-␣ throughout normal pregnancy, more IL-12p70 in the
first and second trimesters, and more IL-18 in the first trimester only. Intracellular cytokine measurements (TNF-␣ and IL12p70)
showed little change by comparison. IFN-␥ production was suppressed in all three trimesters. In pre-eclampsia, IL-18 secretion was
increased. Secreted but not intracellular measures of TNF-␣ and IL-12p70 were also further enhanced compared with normal pregnancy. Inhibition of IFN-␥ production was lost and involved both CD56ⴙ NK and CD56ⴚ lymphocyte subsets. We determined whether
circulating syncytiotrophoblast microparticles (STBM) could contribute to these inflammatory changes. Unbound STBM could be
detected in normal pregnancy by the second trimester and increased significantly in the third. They were also bound in vivo to
circulating monocytes. Women with pre-eclampsia had significantly more circulating free but not cell-bound STBMs. STBMs prepared
by perfusion of normal placental lobules stimulated production of inflammatory cytokines (TNF-␣, IL12p70, and IL-18 but not IFN-␥)
when cultured with PBMCs from healthy nonpregnant women. Inflammatory priming of PBMCs during pregnancy is confirmed and
is established by the first trimester. It is associated with early inhibition of IFN-␥ production. The inflammatory response is enhanced
in pre-eclampsia with loss of the IFN-␥ suppression. Circulating STBMs bind to monocytes and stimulate the production of inflammatory cytokines. It is concluded that they are potential contributors to altered systemic inflammatory responsiveness in pregnancy and
pre-eclampsia. The Journal of Immunology, 2007, 178: 5949 –5956.
5950
TROPHOBLAST MICROPARTICLES AND SYSTEMIC INFLAMMATION IN PREGNANCY
Table I. Cytokine production by PBMCs
Normal Pregnancy
Maternal age (yr)
Nulliparous (%)
*Sampling GAb (wk)
*Delivery GA (wk)
a
b
Preeclampsia
Nonpregnant
(n ⫽ 10)
1st Trimester
(n ⫽ 10)
2nd Trimester
(n ⫽ 10)
3rd Trimester
(n ⫽ 10)
Nonpregnant
(n ⫽ 10)
Normal pregnant
(n ⫽ 10)
Preeclampsia
(n ⫽ 10)
32.3 (6.5)a
60
33.3 (7.1)
60
12.9 (0.7)
39.4 (1.7)
34.3 (5.8)
60
20.4 (3.0)
39.4 (1.9)
29.5 (6.7)
60
31.9 (4.1)
40.7 (1.2)
28.6 (6.3)
60
29.5 (6.7)
60
31.9 (4.1)
40.7 (1.2)
28.7 (6.6)
60
32.1 (3.8)
32.7 (3.6)
Values in parentheses are given as mean (SD) or percentage.
GA, Gestational age.
its evolution during normal pregnancy, and the changes that are
associated with pre-eclampsia. We studied the four inflammatory
cytokines TNF-␣, IL-12, IL-18, and IFN-␥, which were selected
because of their importance in the Shwartzman reaction. We determined how early during normal pregnancy circulating STBMs
can be detected, whether they circulate in maternal blood bound to
PBMCs, specifically monocytes, and whether they stimulate secretion of inflammatory cytokines from PBMCs when cultured ex
vivo. We found that an enhanced inflammatory response can be
elicited from PBMCs from the first trimester onwards, that it is
characterized by suppression of lymphocyte production of IFN-␥
and that this suppression is partially released in pre-eclampsia with
enhanced IL-18 production. STBMs are proinflammatory in culture and could contribute to the systemic inflammatory response
that we observe, both in normal pregnancy and in pre-eclampsia.
Materials and Methods
Reagents and Abs
Penicillin, streptomycin, sodium pyruvate, ␤-ME, PMA, ionomycin,
brefeldin A, saponin, and rat IgG Ig isotypes were obtained from SigmaAldrich. RPMI 1640 with L-glutamine was obtained from Invitrogen Life
Technologies. Recombinant human IFN-␥ was supplied by Cambridge
Biosciences. Monoclonal mouse anti-CD14 (conjugated with FITC), unconjugated anti-CD45, and rat anti-TNF-␣ Abs (FITC-conjugated) were
obtained from Serotec; mouse PE-conjugated anti-IL12p40/70, rat PE-conjugated anti-IL12p70, and FITC-conjugated anti-IFN-␥ were obtained from
BD Pharmingen and mouse FITC-conjugated anti-CD56 from Immunotech.
NDOG2 and ED822 were in-house mAbs to human placental alkaline phosphatase (21) and an unknown Ag expressed on the apical surface of the syncytiotrophoblast (22), respectively. Mouse IgG used in control steps was obtained from Serotech (unconjugated) or BD Pharmingen (FITC-conjugated).
PE-conjugated goat anti-mouse IgG was obtained from Dako.
Patients, normal subjects, and cell purification
To study normal gestational changes, healthy pregnant women were recruited in the first, second, and third trimesters and matched for age (⫾4
years) and parity (0, 1–3, ⱖ4) with nonpregnant women. To study preeclampsia, patients were recruited and matched for age and parity with normal pregnant women and nonpregnant women. Pregnant women were also
matched for gestation (⫾13 days). Normal pregnant women were studied in
the first (12–14 wk), second (15–25 wk), and third (26 – 40 wk) trimesters of
pregnancy. Nonpregnant women of reproductive age were recruited from staff
members. None had any significant past medical history, current illness, or
were taking regular medication (apart from folic acid and iron supplements).
Pregnant patients were not in labor at the time of sampling.
Pre-eclampsia was diagnosed by new hypertension in the second half of
pregnancy (blood pressure ⱖ140/90 mm Hg or an increase of ⬎30/15 mm
Hg from baseline at booking, on at least two occasions at least 6 h apart)
and new proteinuria (ⱖ2⫹ on dipstick testing on at least two occasions or
ⱖ500 mg protein in a 24-h urine collection, in the proven absence of a
urinary tract infection).
Case characteristics are detailed in Tables I and II. These studies were
approved by the Oxfordshire Clinical Research Ethics Committee.
PBMC, culture, and cytokine measurement
Peripheral venous blood (40 ml) was taken into sodium heparin (10 IU/ml),
and the mononuclear cells (PBMCs) were separated by density gradient
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Four cytokines are of potential importance in generating the
excessive inflammatory response of pre-eclampsia: TNF-␣, IL-12,
IL-18, and IFN-␥. They are involved in the Shwartzman reaction,
which is a lethal response to endotoxin after a prior priming injection. Pregnant animals are uniquely sensitive to the Shwartzman
reaction in that the priming administration of endotoxin is not
needed (several authors, for example; see Ref. 12). A notable feature of the HELLP syndrome is its hyperacute presentation in some
women (several authors, for example; see Ref. 13), which matches
the acute time course of the Shwartzman response. In experimental
animals the Shwartzman reaction results from a positive feedback
loop in which activated monocytes secrete IL-12 and IL-18 that
stimulate lymphocytes to produce IFN-␥, which magnifies further
the degree of activation of the monocytes (14). Hepatic necrosis,
which is a typical consequence of the HELLP syndrome, occurs in
a murine variant of the Shwartzman reaction (15), in which it has
been shown that IL-18, IL-12, IFN-␥ and TNF-␣ all contribute to
the experimental pathology. The process can be totally prevented
by prior administration of anti-IL-18 but is only partially ameliorated by anti-IFN-␥ or anti-TNF-␣ (16). We have shown that, in
human pregnancy, circulating monocytes are primed to produce
IL-12 (17) which may indicate that a human equivalent to the
priming of animals may occur.
The candidate stimulants for the systemic inflammatory response of pregnancy include circulating cytokines or antiangiogenic factors (18), products of oxidative stress (19), or subcellular
debris shed from the syncytial surface of the placenta (20). We
were particularly interested in the last, as in human hemochorial
placentation, the outermost layer of the placenta, the syncytiotrophoblast, is in direct contact with maternal blood. Any microparticulate debris from the placenta will be shed into the maternal
circulation and could interact with both maternal leukocytes and
vascular endothelium. Furthermore, the syncytiotrophoblast is
both class I and class II MHC negative and will therefore not
provoke classical T cell allograft responses.
We devised a technique of flow cytometric measurement of the
binding of syncytiotrophoblast membrane Ags to peripheral blood
monocytes. We determined whether such binding occurred in normal pregnancy and, if so, at what stages of pregnancy it could be
detected and if pre-eclampsia changed the degree of binding. In a
final set of experiments, we investigated the role of syncytiotrophoblast membrane microparticles (STBM) that are shed into maternal blood from the placental surface during pregnancy (20) and
whether they can stimulate the production of proinflammatory cytokines. For the latter purpose, microparticles were prepared from
placentas from healthy pregnancies, both by the standard mechanical
method (mSTBM) and from eluates of perfused isolated placental
cotyledons (pSTBM), which may better represent the particles shed in
vivo, and used in coculture experiments with PBMCs.
In this study, we sought to characterize the systemic inflammatory response of pregnancy and pre-eclampsia, the time course of
The Journal of Immunology
5951
Table II. Leukocyte binding of syncytiotrophoblast microparticles
Normal Pregnancy
Maternal age
Nulliparity (%)
*Sampling GAb (wk)
Delivery GA
a
b
Preeclampsia
Nonpregnant
(n ⫽ 25)
1st Trimester
(n ⫽ 8)
2nd Trimester
(n ⫽ 8)
3rd Trimester
(n ⫽ 20)
Nonpregnant
(n ⫽ 20)
Normal pregnant
(n ⫽ 20)
Preeclampsia
(n ⫽ 20)
30.0 (6.0)a
72
34.3 (6.6)
63
12.8 (0.7)
34.4 (6.6)
50
19.3 (2.3)
31.1 (5.2)
75
34.5 (3.3)
29.6 (5.3)
75
31.1 (5.2)
75
34.5 (3.3)
39.6 (1.4)
31.1 (5.2)
75
34.8 (3.1)
35.3 (2.9)
Values in parentheses are given as mean (SD) or percentage.
GA, Gestational age.
Measurement of free STBMs in peripheral blood
Free STBM in plasma samples were measured using an in-house ELISA
(23). In brief, peripheral blood samples were collected into sodium heparin
and centrifuged at 2000 ⫻ g for 15 min at room temperature. The resultant
plasma supernatant was diluted with PBS-E (at least 1/2) and ultracentrifuged (150,000 ⫻ g for 45 min at 4°C). The supernatant was discarded, and
the final pellet was resuspended in 350 ␮l of PBS-BSA and stored at
⫺80°C until use.
The capture Ab was NDOG2 (see above) which has been found to be
optimal for detecting STBM in this ELISA (24). The reporter system detected endogenous alkaline phosphatase activity on the surface of the microparticles by a colorimetric reaction (Invitrogen Life Technologies).
Plates containing triplicate samples were read on an MRX Microplate
reader (Dynex Technologies) at 490 nm, at 5 min. The sensitivity of the
assay is ⬍10 pg/ml (25).
A pooled preparation of mSTBM from five placentas was used to prepare the standards for the STBM ELISA (see below). All the placentas had
been obtained at elective cesarean section from healthy pregnant women
with no history of pre-eclampsia or intrauterine growth retardation. A standard curve was determined to measure the STBM concentration in each 350-␮l
sample in nanograms per milliliter. The readings were adjusted by a concentration factor to calculate the STBM level in the original plasma sample.
Immune cell response to mSTBM and pSTBM
mSTBMs were prepared from normal placentae by a modification of the
method of Smith et al. (26), and the protein content was determined using
a BCA protein assay kit. pSTBM were prepared using a dual placental
perfusion system as described by Eaton and Oakey (27). Placentas were
obtained at cesarean section, without labor, from healthy pregnant women
and were processed immediately. An individual lobule was isolated and
perfused with modified M-199 tissue culture medium (Medium 199 with
L-glutamine and Earle’s salts without NaHCO3, containing 0.8% Dextran
20, 0.5% BSA, 5000 U/L sodium heparin, and 2.75 g/L sodium bicarbonate, pH 7.4) at a controlled rate of 20 ml/min. The perfusion medium was
warmed in a 37°C water bath and oxygenated with 95% O2, 5% CO2. Every
2.5 min after perfusion started, a 50-ml fraction of effluent from the maternal circulation was collected, labeled, and kept on ice. The volume of
fetal effluent was measured simultaneously, and the oxygen concentration
of the maternal side perfusate monitored to ensure the stability of the preparation. Pressure monitors were used to ensure no significant deviations
from baseline during the experimental period. The 50-ml fractions of maternal side perfusate were centrifuged in a Beckman J6-M centrifuge at
2000 ⫻ g for 15 min at 4°C. Aliquots (12 ml) of the supernatants were spun
at 150,000 ⫻ g for 45 min at 4°C in a Beckman L8-80M ultracentrifuge.
Each pellet was resuspended in 350 ␮l of PBS-BSA and stored at ⫺80°C
until use. The protein content of each suspension was measured (BCA
protein assay kit) and adjusted to 1.9 mg/ml. pSTBM in each aliquot were
measured using the ELISA described previously.
To study stimulation of cytokine responses, PBMCs were isolated from
nonpregnant individuals (n ⫽ 3) and cultured for 4 and 24 h, as described
earlier. Cells were stimulated with 150 ng/ml mSTBM or pSTBM (prepared as described previously) or 1 ␮g/ml LPS and 20 ng/ml IFN-␥ (for
TNF-␣, IL-18, and IL-12) or 10 ng/ml PMA and 1 ␮M ionomycin (for
IFN-␥). For the 24-h time point, the cells were preincubated with IFN-␥
for 2 h. Secreted and intracellular levels of TNF-␣, IL-18, and IL-12p70
and IFN-␥ were measured as described previously. In addition, the ability of pSTBM to stimulate TNF-␣ secretion was titrated in a doseresponse curve (0 –300 ␮g/ml).
In vivo binding of STBM to circulating monocytes
PBMCs were prepared as described, and 20 ⫻ 106 cells/ml were resuspended in PBS-BSA. They were double-labeled with Abs against CD14
and ED822 that binds to a specific syncytiotrophoblast marker and has
been found to be optimal for flow cytometry (24). mAb to CD45 (Serotec)
was used as a positive control. Previous work (24) suggested that there is
insignificant binding of STBM to other leukocyte subtypes. Aliquots of
0.5 ⫻ 106 cells in 1.5-ml Eppendorf tubes were labeled in 5 ml of PBS-E,
Downloaded from http://www.jimmunol.org/ by guest on June 16, 2017
centrifugation using Ficoll-Paque PLUS (endotoxin tested). The final pellet
was resuspended in RPMI 1640 with L-glutamine containing 1 mM sodium
pyruvate, 50 U/ml penicillin, 50 ␮g/ml streptomycin, 0.02 mM 2-ME, and
10% FBS, at 2 ⫻ 106 cells/ml, and 1-ml aliquots were placed in a 24-well
Sarstedt plate.
To stimulate TNF-␣, IL-18, and IL-12 production, PBMCs were preincubated for 2 h with 20 ng/ml recombinant human IFN-␥, followed by 22 h
of stimulation with 1 ␮g/ml LPS plus 20 ng/ml IFN-␥. This protocol was
used after preliminary comparisons of different protocols (data not shown).
The final conditions were selected to optimize detection of all three cytokines for both intracellular and culture supernatant measurements. To stimulate IFN-␥ production, PBMCs were incubated with 10 ng/ml PMA and
0.75 ␮g/ml (1 ␮M) ionomycin for 4 h. Brefeldin A (10 ␮g/ml) was used
to inhibit PBMC cytokine secretion, for the measurements of intracellular
cytokine. The cells were incubated at 37°C in 5% carbon dioxide in air.
After the indicated time, they were resuspended and microfuged at
13,000 ⫻ g. The supernatants were stored at ⫺80°C until use.
Cytokines were measured in culture supernatants using commercial
ELISA kits (from PeproTech for TNF-␣, and from BioSource International
for the remaining assays) following the manufacturers’ instructions. The
plates were read on an MRX Microplate Reader using Revelation software.
Samples were analyzed in duplicate together with appropriate standards
and quality controls, and average values were determined.
To measure intracellular cytokines, each PBMC pellet was resuspended
in 100 ␮l of staining buffer (47.5 ml of PBS-E, 2.5 ml of 10% sodium
azide, and 250 ␮l of FBS). Cells were labeled in 1.5-ml Eppendorf tubes,
on ice, in the dark. Abs directly conjugated to CD14 (monocytes; final
dilution, 1/100) and to CD56 (NK cells; 1/50) were used to identify relevant cell types. Then, 25 ␮l of the Ab diluted in staining buffer were added
to 0.5 ⫻ 106 PBMCs. Controls samples comprised staining buffer alone,
and mouse IgG isotype Ig (MsIgG). The suspensions were incubated for 30
min, washed with 400 ␮l of staining buffer, and microfuged at 13,000 ⫻ g
for 7 s; the pellet was resuspended in 50 ␮l of fixation buffer (4% paraformaldehyde in PBS-E), and left for 20 min. Samples were again washed,
spun, and resuspended in 25 ␮l of permeabilization buffer (47.5 ml of
PBS-E, 2.5 ml of 10% sodium azide, 250 ␮l of FBS, and 50 mg of saponin)
containing the appropriate Ab and incubated for a further 30 min. Directly
conjugated Abs were used to detect TNF-␣ (final dilution, 1/5), IL-12p70
(2/5) and IL-12p40/70 (2/5) in monocytes, and IFN-␥ (1/50) in NK cells.
Control samples comprised cells in permeabilization buffer or incubated
with MsIgG and rat IgG of the appropriate isotype.
Ten thousand events were collected by flow cytometry using a Beckman
Coulter EPICS Altra, at 520 nm or 578 nm for analysis off-line. Monocytes
and lymphocytes were identified by their optical characteristics. Gates set
on single parameter histograms, using the isotype-negative controls to include ⱕ1% of events, were used to determine the positive populations at
both wavelengths. A quadrant gate was applied to a two-parameter plot
corresponding to these negative control gates. Similar gates were set for the
lymphocyte population, to measure the production of IFN-␥ by both
CD56⫹and CD56⫺ lymphocytes. The proportions of cytokine positive
monocytes or lymphocytes were measured. We were unable to develop a
similar method for detecting intracellular IL-18 by monocytes using the
commercially available mAb; therefore, only secreted IL-18 levels in culture supernatants could be measured.
5952
TROPHOBLAST MICROPARTICLES AND SYSTEMIC INFLAMMATION IN PREGNANCY
Table III. Free plasma STBMs in the study groups
Normal Pregnancy
Nonpregnant
(n ⫽ 10)
1st Trimester
(n ⫽ 10)
2nd Trimester
(n ⫽ 10)
3rd Trimester
(n ⫽ 14)
Nonpregnant
(n ⫽ 14)
Normal pregnant
(n ⫽ 14)
Preeclamptic
(n ⫽ 13)
32.3 (6.5)a
60
33.3 (7.1)
60
12.9 (0.7)
34.3 (5.8)
60
20.4 (3.0)
30.6 (5.9)
57
33.4 (4.2)
30.1 (6.3)
57
30.6 (5.9)
57
33.4 (4.2)
40.5 (1.2)
30.0 (7.2)
69
33.5 (3.9)
33.6 (3.8)
Maternal age (yr)
Nulliparity (%)
Sampling GAb (wk)
Delivery GA (wk)
a
b
Preeclampsia
Values are given as mean (SD) or percentage.
GA, Gestational age.
ters which was, however, significant only in the third trimester.
When intracellular production was analyzed, the pattern was similar, with a more sustained inhibition in the CD56⫺ lymphocytes
than in the CD56⫹ NK (Fig. 1C).
Cytokine production by monocytes and lymphocytes
in pre-eclampsia
This comparison was confined to third trimester pregnancies and
nonpregnant women. In general the trends documented in Fig. 1A
for late normal pregnancy were confirmed for secreted TNF-␣,
IL-12p70 and Il-18. In pre-eclampsia secreted TNF-␣ was slightly
Statistics
Data were compared using the Wilcoxon matched pairs test for studies with
equal numbers of matched subjects in each group, the Mann-Whitney U
test for studies with unequal numbers in the groups and the sign test for
aggregated data.
Results
Characteristics of subjects
There were different groups of subjects for the studies of cytokine
production by PBMCs (group A), of free syncytiotrophoblast microparticles (group B), and of leukocyte-bound STBM (group C).
In each category, there were two subgroups for changes through
normal pregnancy and changes in pre-eclampsia. Matching ensured that the relevant groups were comparable except for pregnancy outcome which was associated with substantially earlier
gestational ages at delivery in pre-eclampsia. The patient details
are shown in Tables I–III.
Cytokine production by monocytes and lymphocytes in normal
pregnancy
Short term cytokine production by unstimulated monocytes or
lymphocytes for 24 h or less was low, without significant differences between any of the groups (data not shown). Therefore, only
data for stimulated monocytes and lymphocytes are presented.
The overall response in terms of TNF-␣, IL-12p70, and IL-18
was a significant increase in secretion relative to the nonpregnant
baseline in the first trimester. The patterns for each cytokine differed in the remaining two trimesters. IL-18 and IL-12p70 production showed a progressive decline after the first trimester, almost returning to the baseline level by the third in the case of IL-18.
TNF-␣ secretion, however, reached a plateau in the second trimester
that was sustained in the last trimester (Fig. 1A). The changes in intracellular cytokine expression (TNF-␣ and IL-12p70 only) were less
variable. There were small first trimester increases, which were significant for TNF-␣ only (Fig. 1B) and a nonsignificant decline in
IL-12p70 which followed the trend for secreted cytokine.
The production of IFN-␥ had a reciprocal pattern. Pregnancy
samples were characterized by less production in all three trimes-
FIGURE 1. Cytokine production by PBMC in normal pregnancy. NC,
Samples from healthy nonpregnant women of reproductive age), 1st, 2nd,
3rd, Samples from pregnant women in each of the three trimesters of pregnancy (n ⫽ 10 for each group). A, Secreted cytokines were measured by
ELISA in supernatants of cultured PBMCs. B, Intracellular production of
TNF-␣ and IL-12p70 were measured flow cytometrically as percent of
positive monocytes identified by labeling with FITC-conjugated antiCD14. C, Intracellular production of IFN-␥ was measured in CD56⫹ or
CD56⫺ lymphocytes. For CD56⫺ lymphocytes: p ⬍ 0.05 for NC vs first or
third trimesters; p ⫽ 0.10 for NC vs second trimester. For CD56⫹ lymphocytes, p ⫽ 0.07, 0.10, and 0.23 for first, second, and third trimesters,
respectively. ⴱ, p ⬍ 0.05; ⴱⴱ, p ⬍ 0.01.
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20 mM glucose, and 250 ␮l of normal human serum, while on ice and in
the dark. Primary labeling was with 25 ␮l of unconjugated mouse monoclonal ED822 Ab (1/200) incubated for 30 min. After a washing (1 ml of
PBS-BSA, microcentrifuged at 13,000 ⫻ g for 7 s), the sample was incubated for 30 min with secondary Ab (goat anti-mouse conjugated to PE at
1/100). After further washing, unoccupied binding sites were blocked using
90 ␮l of a 1/6 dilution of MsIgG. Monocytes were then labeled with FITCconjugated anti-CD14. Cell pellets were resuspended in 25 ␮l of the Ab
diluted 1/100 and incubated for 30 min. After a washing, each sample was
resuspended in 1 ml of PBS/BSA, transferred to flow cytometry tubes, and
kept on ice until analysis. Appropriate negative and single and double
positive controls were also prepared. Samples were analyzed as described
previously. The percentages of monocytes positive for both CD14 and a
trophoblast marker (ED822) were determined.
The Journal of Immunology
diminished and no different from nonpregnancy samples; secreted
IL-12p70 was unchanged from normal pregnancy and secreted
Il-18 increased above normal pregnancy, to a significant extent
relative to the nonpregnant baseline (Fig. 2A). Intracellular TNF-␣
and IL-12p70 were once again no different between normal pregnancy and nonpregnancy but the % positive PBMCs were significantly increased in pre-eclampsia for TNF-␣ only (Fig. 2B). The
reciprocal inhibition of IFN-␥, both secreted and intracellular was
again observed in normal pregnancy. The striking change was loss
of this inhibition in pre-eclampsia (Fig. 2A and 2C).
FIGURE 3. Circulating free and cell-bound plasma STBMs in normal
pregnancy and pre-eclampsia. Free STBMs (A) were measured by ELISA
in maternal peripheral vein plasma, and bound microparticles (B) were
measured by flow cytometry as percent of positive monocytes (identified
by labeling with FITC-conjugated anti-CD14) that bound STBM (identified by labeling with a mAb (ED822) that binds to a specific syncytiotrophoblast marker and a secondary goat anti-mouse conjugated to PE). A and
B (left) in the three trimesters of normal pregnancy (1st, 2nd, 3rd, n ⫽ 10,
10, and 14, respectively) compared with nonpregnant women (NC, n ⫽
10). A and B (right) in pre-eclamptic patients (PE, n ⫽ 13) compared with
third-trimester normal pregnant (NP, n ⫽ 14) and nonpregnant (NC, n ⫽
14) women. ⴱ, p ⬍ 0.05; ⴱⴱ, p ⬍ 0.01; ⴱⴱⴱ, p ⬍ 0.001.
tive to nonpregnancy was detected in normal pregnant ( p ⱕ 0.001)
but no further increase was apparent in pre-eclampsia (Fig. 3B).
Measurement of cytokine production from PBMCs after
stimulation by pSTBMs
mSTBM did not stimulate TNF-␣ (Fig. 4A), IL-12, IL-18, or IFN-␥
(data not shown) production by PBM from nonpregnant women. In
Free and bound circulating STBM in normal pregnancy and
pre-eclampsia
Free STBM were not detected in first trimester samples but were
present at significant levels in the second trimester and increased further in the third (Fig. 3A). In the first trimester, there was no significant binding of STBM compared with the nonpregnant controls (Fig.
3B); but significant binding was detected in the second trimester with
a further increase in the third trimester.
Similar techniques were used to compare samples from preeclamptic women with those from matched control normally pregnant
women. As we have reported before (24), free STBM concentrations
were significantly higher in pre-eclampsia than in matched control
normal pregnant women (Fig. 3A). However, this change was not
seen when bound STBM were studied. The significant increase rela-
FIGURE 4. TNF-␣ secretion by cultured PBMC from nonpregnant
women (n ⫽ 3) measured by ELISA. A, After stimulation with mechanically prepared (mS) or STBM from perfused placental lobules (pS), compared with LPS and IFN-␥ (LP/I␥, positive control) and no stimulant (N,
negative control). B, Dose responses to stimulation by increasing amounts
of pS (␮g/ml): 1 (0 –20); 2 (20 – 40); 3 (40 – 80); 4 (80 –120); 5 (120 –300);
compared with perfusion medium (P) and supernatant after removal of pS
(S, negative control). ⴱ, p ⬍ 0.05.
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FIGURE 2. Cytokine production by PBMC in pre-eclampsia (PE), compared with 3rd trimester normal pregnant (NP), and nonpregnant control
women (NC). A, Secreted cytokines were measured by ELISA in supernatants of cultured PBMCs. B, Intracellular production of TNF-␣ and IL12p70 were measured flow cytometrically as percent of positive monocytes
identified by labeling with FITC-conjugated anti-CD14. C, Intracellular
production of IFN-␥ was measured in CD56⫹ or CD56⫺ lymphocytes (n ⫽
10 for each group). ⴱ, p ⬍ 0.05; ⴱⴱ, p ⬍ 0.01.
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TROPHOBLAST MICROPARTICLES AND SYSTEMIC INFLAMMATION IN PREGNANCY
contrast, pSTBMs were able to stimulate moderate levels of TNF-␣
(Fig. 4A) with a significant increase in a dose-response curve (Fig.
4B). Not only TNF-␣ but also IL-18 and low levels of IL-12p70 and
IFN-␥ production could be detected after only 4 h of culture of
pSTBM with PBMCs (Fig. 5A; p ⬍ 0.05). These findings were confirmed by measuring intracellular production in PBMCs (Fig. 5B).
pSTBM also stimulated low intralymphocytic levels of IFN-␥ production from both CD56⫹ and CD56⫺ lymphocytes.
Discussion
The PBMC response in normal pregnancy changed in two remarkable ways, relative to nonpregnancy. The first was that inflammatory responsiveness was increased particularly in the first trimester: cultured PBMCs showed significantly enhanced production of
all the three inflammatory cytokines that were studied although
Il-18 production declined in later trimesters. The second notable
feature was that IFN-␥ was suppressed, although this was not statistically significant in the second trimester ( p ⫽ 0.10, n ⫽ 10).
The response in pre-eclampsia was also altered in important
ways, relative to matched normal pregnancy samples. IL-18 production was significantly increased and production of IFN-␥ was
no longer suppressed. These patterns are consistent with the consensus that there is a type 2 bias during normal human pregnancy
(28, 29) and extend our earlier findings in relation to normal pregnancy (6, 17).
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FIGURE 5. Cytokine production by nonpregnancy PBMC (n ⫽ 3) after
stimulation with pSTBM (ST) compared with LPS and IFN-␥ (LP/I␥, positive control) and no stimulant (N, negative control). A, Secreted cytokines
were measured by ELISA in supernatants of cultured PBMCs (p ⬍ 0.01 for
aggregated data compared with control). B, Intracellular production of
TNF-␣ and IL-12p70 were measured flow cytometrically as percent of
positive monocytes identified by labeling with FITC-conjugated anti-CD14
(significance not tested). C, Intracellular production of IFN-␥ was measured flow cytometrically in CD56⫹ or CD56⫺ lymphocytes (p ⬍ 0.025,
for aggregated data compared with control).
We studied isolated and stimulated PBMCs by measuring secreted inflammatory cytokines from cultured PBMCs and also intracellular cytokine production by flow cytometry on the same
samples. Hence, the measures were not of basal production, which
was negligible, but reflect priming and not in vivo activity. The
gestational time courses of inflammatory cytokine production differed. For TNF-␣, an early increase was sustained throughout
pregnancy; for IL-12 it peaked in the second trimester and declined
thereafter; for IL-18, the increase was only significant in the first
trimester. These changes could not be simply ascribed to changes
in capacity to produce IFN-␥ which was suppressed throughout
pregnancy. In normal pregnancy, there is a monocytosis from an
early stage (30) which might contribute to at least part of the differences, relative to the nonpregnant control women, in the different gestational patterns for secreted and intracellular TNF-␣ and to
a lesser extent for IL12p70. In pre-eclampsia, there is no consensus
about changes in monocytes counts relative to normal pregnancy
(31, 32). Monocyte counts were not available to interpret these
results; thus, this possibility cannot be discounted, but it is unlikely
given the evidence for inflammatory priming during pregnancy in
animals (12) and humans (17).
It has been previously reported that unstimulated PBMCs from
pregnant women produce less IL-12 relative to nonpregnancy, which
the authors suggested explains the type 2 predominance of pregnancy
(33, 34). These authors also observed increased spontaneous IL-18
production in the third trimester relative to nonpregnancy and argue
that IL-18 alone is predominantly a Th2 cytokine, which is converted
to a Th1 cytokine by IL-12 which induces IL-18R␤ (35).
The changes in culture were not mirrored in the measures of
intracellular TNF-␣ or IL-12, which varied little throughout pregnancy apart from a small significant increase in TNF-␣ in the first
trimester when compared with normal pregnancy. Measures of the
intensity of labeling for intracellular cytokine were similarly unremarkable (data not shown).
For TNF-␣ these results are comparable to those we previously
obtained for third trimester samples but do not replicate the evidence of priming for IL-12 production that we also demonstrated
(17). In the present study, more intense stimulation with LPS
(1000 vs 40 ng/ml) and IFN-␥ (20 vs 5.5 ␮g/ml) was used, and a
2-h prestimulation with IFN-␥ was added. In addition, monocytes
were identified differently, i.e., on optical characteristics not by
CD14 expression because CD14 is shed with monocyte activation
(36) and may then become an unreliable monocytes marker. The
more intense stimulation used in the present study may have obliterated the differences that were previously seen.
Basal production of IL-12 appears to be suppressed during normal
pregnancy but, under certain conditions of stimulation, primed for
increased production (13, 33). The most likely explanation for this
conclusion is that, in vivo, the production of IL-12 is suppressed by
the concurrent suppression of IFN-␥ production. In vitro stimulation
included exogenous IFN-␥, which would be expected to reverse the
suppressive effects that low availability of IFN-␥ in vivo causes.
Our present work is, in general, consistent with data of
McCracken et al. (29) who studied isolated T cells from women
with normal third trimester pregnancies. They demonstrated that
inhibition of intracellular production of IFN-␥ was caused specifically by loss of NF-␬B activity. This, they speculated, could result
from the high levels of circulating steroids which are characteristic
of all stages of pregnancy from the second half of the first trimester
onwards. This study extends their findings in three ways: we show
1) that IFN-␥ production is reduced in all three trimesters; 2) that
it is not limited to CD56⫺ cells, which are predominantly T cells,
but involves circulating NK cells (CD56⫹); and 3) that the inhibition is reduced or lost in pre-eclampsia. The involvement of NK
The Journal of Immunology
signals (42). They are thought to have important proinflammatory
and procoagulatory roles (44, 45), including promoting interaction
between endothelial cells and monocytes (40), platelet activation
(46), and apoptosis (47). In relation to our studies, a key question
is, “Can these circulating STBMs contribute to the immunoregulation of normal pregnancy or its dysregulation in pre-eclampsia?”
STBMs, shed from perfused placentas ex vivo, induced mononuclear cell production of the proinflammatory cytokines TNF-␣,
IL-18, and IL-12, demonstrating that they have proinflammatory
potential as described for microparticles from other cellular types
(44, 45). In another study, STBMs prepared by mechanical methods increased IFN-␥ production by peripheral blood T cells purified from healthy blood donors (38). Using different techniques,
we observed a small but consistent increase in IFN-␥ production.
We did not test STBM prepared from pre-eclampsia placentas, but
others find that they activate inflammatory leukocytes significantly
more than STBM from normal pregnancies (48). This suggests a
qualitative difference, perhaps owing to oxidized lipids (49), in
addition to the quantitative increases that we document here. The
mechanism of stimulation is not known. It is possible that oxidized
lipids or other molecules carried by STBM are detected directly as
danger signals (50) to stimulate directly inflammatory responses.
An alternative explanation is that dendritic cells or monocytes process the particles, which can then be presented as small molecules
by classical or nonclassical routes.
The systemic inflammatory response was detected from the first
trimester onwards whereas circulating STBM could be detected
only during the second and third trimesters. We have previously
emphasized the two interfaces of maternal-fetal interactions (see
Introduction). When the uteroplacental circulation opens toward
the end of the first trimester (51), interface II begins to be activated. Before then, STBM cannot be released directly into the
maternal circulation, and at that time and immediately afterward
what is released is limited by the small size of the placenta and
may be undetectable with our current assays. We speculate that
earlier systemic inflammation is secondary to the intense localized
interactions between decidual immune cells (maternal) and trophoblast (fetal) at interface I. Therefore, our current data raise the
possibility that shed STBM, which become detectable in the second trimester, increase in the third and are further enhanced in
pre-eclampsia, contribute to a second phase of systemic inflammation by stimulating an increasing counter stimulus to the immunosuppression of normal pregnancy and have the potential to
overcome it. If IFN-␥ production were promoted, (for example in
pre-eclampsia or coincident infection), the capacity to over-produce IL-12 and IL-18 could be released and generate the feedforward loop that promotes the Shwartzman reaction. This may
help to explain the unique sensitivity of pregnant animals to the
Shwartzman reaction, without an additional priming dose of endotoxin. These changes probably relate to immune interface II as
discussed in the Introduction. However, the inflammatory changes
in the first trimester more likely reflect events secondary to the
intense remodeling in the decidua, which involves inflammatory
and immune processes (52). We have not studied the regression of
the systemic inflammatory changes after delivery. The little that
we do know is that detectable STBM are cleared almost completely from plasma samples within 24 h (A. Reddy, I. L. Sargent,
and C. W. Redman, unpublished data).
Acknowledgments
We are grateful to Carol Simms, Alison Wright, and Hazel Coburn who
undertook most of the patient recruitment.
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cells is consistent with our previous report that suggested that both
NK and NKT cells play a role in determining the type 1/type 2
balance in pregnancy and pre-eclampsia (2). In this study, the phenotype of the CD56⫹ cells was not investigated in detail, and the
CD56⫺ cells were not characterized other than for this attribute
and their presence in the optical gate characteristic of lymphocytes.
It was therefore not possible to explore this issue in detail, and we
cannot exclude the participation of T cells.
We investigated circulating STBMs, shed from the placenta, as
a potential stimulus for systemic inflammatory changes, particularly in pre-eclampsia. Free particles were detectable in plasmas of
women with third trimester pregnancies and were significantly increased when pre-eclampsia was confirmed, which confirms our
previous report (24). In this study, we show that the particles cannot be detected in the first trimester but increase thereafter. In
normal pregnancy, significant monocyte binding of STBMs could
also be detected by the second trimester, with a further increase in
the third trimester. In pre-eclamptic patients, there were also significant levels of monocytes binding STBMs, but no further increase compared with gestation-matched normal pregnant controls,
unlike the changes in unbound STBM.
We used mechanically prepared STBM and compared them
with preparations from perfusates of normal term placentas because the latter are considered to be more physiological and free of
contamination with microparticles derived from maternal sources
(37, 38). The optimal conditions required for leukocyte activation
by STBM remain undetermined. STBMs alone were added,
whereas in vivo the presence of one or more proinflammatory cytokines would be predicted to enhance stimulation. STBMs stimulated only low levels of IFN-␥ production in vitro, although it was
always more than the negative control for both plasma and intralymphocytic measurements. The negative control samples were
limited by the possible size of the experiments. They might have
included microparticles from other sources, such as mechanically
processed RBC membranes, eluates from perfused nonplacental
tissues, or inert synthetic particles. We did not examine microparticles prepared from pre-eclampsia placentas, although these might
better reflect the in vivo situation. A further difficulty with these
kinds of experiments is selecting an appropriate control for trophoblast microparticles as the syncytiotrophoblast is a unique tissue. A continuing problem at this stage of microparticle research is
that the different preparations and microparticle subtypes have not
been standardized (39). The importance of this is underlined by the
different effects of sSTBM and pSTBM. As stated, we standardized our preparations in terms of their protein content, but it is
clearly important that more work is done to address this problem.
A final question is whether the level of cytokine production depends on the concentration of STBM. A pSTBM dose of 150
␮g/ml was chosen for these initial experiments, which is more than
one order of magnitude greater than that detected in the circulation.
However, what is measured in the circulation is probably only a
small part of the total of shed STBM given that most will be
rapidly cleared by monocytes and the reticuloendothelial system.
Thus, clearance and concentration in the spleen and liver may create different and more reactive conditions.
The release of high levels of microparticles from the placenta
into the maternal circulation in normal pregnancy and pre-eclampsia constitutes a specialized challenge to the maternal immune system. However, it is not unique as apoptotic release of microparticulate membrane fragments into peripheral blood has been
implicated in several pathological conditions (summarized by Ahn
in Ref. 39). Circulating microparticles may be derived from leukocytes (40), endothelial cells (41), platelets (42), or RBC (43).
The particles carry bioactive molecules that deliver intercellular
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Disclosures
The authors have no financial conflict of interest.
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Zeini, M., P. G. Través, R. López-Fontal, C. Pantoja, A. Matheu, M. Serrano, L. Boscá, and S. Hortelano. 2006. Specific
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In Fig. 3B, the IAP-1 blot for wild-type (WT) cells is duplicated in error in the column for p53⫺/⫺ macrophages. The
corrected Fig. 3 is shown below.
Rutjens, E., S. Mazza, R. Biassoni, G. Koopman, L. Radic, M. Fogli, P. Costa, M. C. Mingari, L. Moretta, J. Heeney, and
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The sixth institution in the author affiliations is incorrect. The corrected list is shown below.
*Biomedical Primate Research Centre, Rijswijk, The Netherlands; †Centro di Eccellenza per la Ricerca Biomedica,
Genoa, Italy; ‡Dipartimento di Medicina Sperimentale, Università di Genova, Genoa, Italy; §Istituto Scientifico Giannina
Gaslini, Genoa, Italy; ¶Dipartimento di Oncologia Biologia e Ginecologia, Università di Genova, Genoa, Italy; 储Istituto
Nazionale per la Ricerca sul Cancro, Genoa, Italy; and #Dipartimento di Medicina Interna, Università di Genova, Genoa,
Italy
Copyright © 2007 by The American Association of Immunologists, Inc. 0022-1767/07/$2.00
www.jimmunol.org
1390
CORRECTIONS
Germain, S. J., G. P. Sacks, S. R. Soorana, I. L. Sargent, and C. W. Redman. 2007. Systemic inflammatory priming in
normal pregnancy and preeclampsia: the role of circulating syncytiotrophoblast microparticles. J. Immunol. 178:
5949 –5956.
The third author’s last name is incorrect. The correct name is Suren R. Sooranna.
Cassatella, M. A., G. Pereira da Silva, I. Tinazzi, F. Facchetti, P. Scapini, F. Calzetti, N. Tamassia, P. Wei, B. Nardelli,
V. Roschke, A. Vecchi, A. Mantovani, L. M. Bambara, S. W. Edwards, and A. Carletto. 2007. Soluble TNF-like cytokine
(TL1A) production by immune complexes stimulated monocytes in rheumatoid arthritis. J. Immunol. 178: 7325–7333.
The second author’s name is incorrect. The correct name is Gabriela Pereira-da-Silva.
Selvaraj, R. K., and T. L. Geiger. 2007. A kinetic and dynamic analysis of Foxp3 induced in T cells by TGF-␤.
J. Immunol. 178: 7667–7677.
Changes that the authors did not request were made in production after the authors returned their page proofs resulting
in publication of the article with multiple errors. The editors and staff of The Journal of Immunology apologize to the
authors and readers for this error.
The entire article is reproduced correctly on the following pages in print only. The errors have been corrected in the
online version, which now differs from the print version as originally published.
The Journal of Immunology
CORRECTIONS
A Kinetic and Dynamic Analysis of Foxp3 Induced
in T Cells by TGF-␤1
Ramesh K. Selvaraj and Terrence L. Geiger2
TGF-␤ induces Foxp3 expression in stimulated T cells. These Foxp3ⴙ cells (induced regulatory T cells (iTreg)) share functional
and therapeutic properties with thymic-derived Foxp3ⴙ regulatory T cells (natural regulatory T cells (nTreg)). We performed a
single-cell analysis to better characterize the regulation of Foxp3 in iTreg in vitro and assess their dynamics after transfer in vivo.
TGF-␤ up-regulated Foxp3 in CD4ⴙFoxp3ⴚ T cells only when added within a 2- to 3-day window of CD3/CD28 stimulation. Up
to 90% conversion occurred, beginning after 1–2 days of treatment. Foxp3 expression strictly required TCR stimulation but not
costimulation and was independent of cell cycling. Removal of TGF-␤ led to a loss of Foxp3 expression after an ⬃4-day lag. Most
iTreg transferred into wild-type mice down-regulated Foxp3 within 2 days, and these Foxp3ⴚ cells were concentrated in the blood,
spleen, lung, and liver. Few of the Foxp3ⴚ cells were detected by 28 days after transfer. However, some Foxp3ⴙ cells persisted even
to this late time point, and these preferentially localized to the lymph nodes and bone marrow. CXCR4 was preferentially
expressed on Foxp3ⴙ iTreg within the bone marrow, and CD62L was preferentially expressed on those in the lymph nodes. Like
transferred nTreg and in contrast with revertant Foxp3ⴚ cells, Foxp3ⴙ iTreg retained CD25 and glucocorticoid-induced TNFR
family-related gene. Thus, Foxp3 expression in naı̈ve-stimulated T cells is transient in vitro, dependent on TGF-␤ activity within
a highly restricted window after activation and continuous TGF-␤ presence. In vivo, a subset of transferred iTreg persist long
term, potentially providing a lasting source for regulatory activity after therapeutic administration. The Journal of Immunology,
2007, 178: 7667–7677.
I
mmunological tolerance is achieved developmentally in the
thymus as well as through peripheral mechanisms. CD4⫹
regulatory T cells (Treg)3 that express the forkhead transcription factor Foxp3 are critical for maintaining peripheral tolerance;
their deficiency leading to early-onset, fatal autoimmune inflammation (1). Foxp3 expression is not only a marker for Treg, but
appears to administer a developmental program endowing T cells
with regulatory function. Thus, CD4⫹ T cells expressing retrovirally transduced Foxp3 display regulatory properties similar to endogenous Treg (2, 3). Treg are largely produced in the thymus
(natural Treg (nTreg)) and constitute ⬃3– 6% of CD4⫹ T cells (4).
More recent studies have shown that Foxp3 may also be induced
in CD4⫹Foxp3⫺ T cells in vivo during some immune responses,
or in vitro after stimulation of Foxp3⫺ cells in the presence of
TGF-␤ (induced Treg (iTreg)) (5– 8).
TGF-␤ is a critical cytokine for preserving immune homeostasis
(9). TGF-␤-deficient mice or mice expressing dominant negative
TGF-␤ receptors on T cells develop spontaneous, early-onset autoimmune disease (10, 11). This results both from cell autonomous
Department of Pathology, St. Jude Children’s Research Hospital, Memphis,
TN 38105
Received for publication January 12, 2007. Accepted for publication April 9, 2007.
The costs of publication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked advertisement in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
1
This work was supported by National Institutes of Health Grant R01 AI056153
(to T.L.G.) and by the American Lebanese Syrian Associated Charities/St. Jude
Children’s Research Hospital (to T.L.G. and R.K.S.).
2
Address correspondence and reprint requests to Dr. Terrence L. Geiger, Department
of Pathology, St. Jude Children’s Research Hospital, 332 North Lauderdale Street,
D-4047, Memphis, TN 38105. E-mail address: [email protected]
3
Abbreviations used in this paper: Treg, regulatory T cell; GITR, glucocorticoidinduced TNFR family-related gene; iTreg, induced Treg; nTreg, natural Treg; LN,
lymph node; rhIL-2, recombinant human IL-2; CD62L, CD62 ligand.
Copyright © 2007 by The American Association of Immunologists, Inc. 0022-1767/07/$2.00
effects of TGF-␤ deficiency on effector T cells and from defects
in the Treg compartment. TGF-␤⫺/⫺ mice demonstrate impaired maintenance of Foxp3⫹ Treg, indicating that TGF-␤
plays a critical role in their homeostasis (12, 13). Although
deficiency in TGF-␤ dominantly leads to an autoaggressive phenotype, the immunomodulatory role of TGF-␤ is likely complex. TGF-␤ also plays a crucial role in promoting the development of inflammatory Th17 cells and plays a supportive role
in Th1 T cell development (14, 15).
Several reports have demonstrated utility in manipulating disease states by altering Treg numbers or activity in animal models
(16 –20). Increasing the number of Treg by adoptive transfer can
diminish pathologic inflammation. Acquiring adequate numbers of
Treg for treatment, however, represent a significant challenge due
to the small number of nTreg present in vivo and their anergic state
in vitro (21). An alternative and potentially simpler approach is to
produce Foxp3⫹ iTreg from CD4⫹ Foxp3⫺ T cells by stimulation
with TGF-␤, which may then be used as an immunotherapeutic
surrogate for nTreg. Indeed, iTreg are able to suppress T cell responses in vitro (22), educate alloreactive CD4⫹CD25⫺ cells to be
suppressive (5), and have shown significant potential in the treatment or prevention of graft rejection, colitis, and diabetes in animal models (23–25).
The therapeutic applicability of iTreg will not only depend on
their expression of Foxp3, but on other cellular characteristics. For
instance, one leading hypothesis is that Treg development is
guided by a high avidity for self-Ag (26, 27). iTreg, being derived
from Foxp3⫺ T cells, lack this high avidity, which may influence
their homeostatic or other properties. Differences between thymically derived nTreg and ex vivo-generated iTreg are not well studied. To better characterize iTreg, we have used single-cell analysis
to assess the kinetics and sustainability of Foxp3 after induction
with TGF-␤ in vitro and the cellular dynamics of iTreg in vivo. We
demonstrate that iTreg development requires TGF-␤ exposure
CORRECTIONS
within a narrow window after stimulation and that Foxp3 persistence requires continued exposure to TGF-␤. After adoptive transfer, iTreg predominantly and rapidly revert to Foxp3⫺ T cells. A
subset of cells, however, retain Foxp3 for a longer time. These
cells primarily localize to and possibly expand within the bone
marrow and lymph nodes. We conclude that for the majority of
iTreg, Foxp3 expression is transient and in vitro is highly dependent on exogenous TGF-␤ exposure. Some cells, however, develop
stable expression of Foxp3 in vivo and phenotypically resemble
nTreg.
Materials and Methods
Animals
Mice in which a GFP-Foxp3 fusion has been homologously inserted at the
Foxp3 locus have been described (1) and were backcrossed more than five
generations onto the C57BL/6J (Thy1.2⫹, CD45.2⫹) background before
analysis. Male mice screened for GFP-Foxp3 were used for experimentation. Some GFP-Foxp3 mice were subsequently bred with CD45.1-congenic mice to obtain CD45.1⫹ GFP-Foxp3 mice. C57BL/6J mice and congenic CD45.1 (B6.SJL-Ptprca Pepcb/BoyJ) and Thy1.1 (B6.PL-Thy1a/CyJ)
mice were purchased from The Jackson Laboratory. Experimentation was
performed in accordance with institutional animal care and use procedures.
Media, reagents, and Abs
Media for T cell cultures was prepared as described earlier (28). Unconjugated
anti-CD3, anti-CD28, and anti-CD16/CD32 Fc block (2.4G2) and fluorochrome-conjugated anti-CD4 (L3T4), anti-CTLA-4, and anti-CXCR4 were
purchased from BD Pharmingen. All other Abs used and recombinant human
TGF-␤1 were purchased from eBioscience. Sulfate latex beads (Molecular
Probes and Invitrogen Life Technologies) were coated with anti-CD3 (40 ␮g/
ml) or anti-CD28 (40 ␮g/ml) or anti-CD3/CD28 (13.3:26.6 ␮g/ml) as per the
manufacturers’ instruction.
Cell purification and cell culture
Lymph nodes (LN) and spleen cells were collected as described previously (28). CD4⫹Foxp3⫹ (nTreg) and CD4⫹Foxp3⫺ (non-Treg) cells
were isolated by flow cytometric sorting on a MoFlo high-speed sorter
(DakoCytomation) gating on CD4 and GFP (Foxp3) expression (28).
Sorted cell purity ranged from 97 to 99%. nTreg were grown in medium
supplemented with 1 ng/ml PMA, 200 ng/ml ionomycin, and 100 U/ml
recombinant human IL-2 (rhIL-2; National Cancer Institute Biological
Resources Branch Repository), which we found to optimally preserve
Foxp3 expression. CD4⫹Foxp3⫺ cells were stimulated with antiCD3/
CD28 beads at a cell:bead ratio of 1:1 and supplemented with 100 U/ml
rhIL-2 with or without TGF-␤ (10 ng/ml) for 7–9 days to obtain iTreg
or activated Foxp3⫺ cells, respectively. Cells were split into cytokinecontaining medium, as needed, to prevent overcrowding.
Foxp3 regulation in CD4⫹Foxp3⫺ cells
Sorted CD4⫹Foxp3⫺ cells (2.5 ⫻ 104 per well) were stimulated in 96-well
plates in medium supplemented with anti-CD3/CD28-coated beads (cell:
bead ratio of 1:1) and 100 U/ml rhIL-2, or as indicated. TGF-␤ (10 ng/ml)
was added at day 0 or the indicated time after TCR stimulation. For Foxp3
reversion kinetics, cells were stimulated as described above and TGF-␤
was removed at the indicated time by removing the supernatant and washing with PBS three times before adding back IL-2-containing medium. To
study Foxp3 up-regulation in memory T cells, CD4⫹CD44highCD45Rblow
Foxp3⫺ cells and control CD4⫹Foxp3⫺ cells were isolated by flow cytometric sorting, and 2 ⫻ 104 cells/well cultured in 96-well plates in medium
supplemented with anti-CD3/CD28-coated beads (cell:bead ratio of 1:1),
100 U/ml rhIL-2, with or without TGF-␤ (10 ng/ml). To prevent overcrowding in longer-term cultures, wells were examined and split every 3– 4
days into medium supplemented with the same cytokines (IL-2 with or
without TGF-␤) as initially present. Quadruplicate samples for each condition were analyzed. Foxp3 analyses were performed every 24 h or as
indicated by quantitative flow cytometry by measuring the GFP
fluorescence.
PKH26 proliferation analysis
Naive flow cytometrically purified CD4⫹Foxp3⫺ cells were dye labeled
using the PKH26 red fluorescent cell linker kit (Sigma-Aldrich) as per the
manufacturer’s instructions, except that 1 ⫻ 10⫺6 M PKH26 dye was used
for 1 ⫻ 107 cells. Cells were cultured in medium supplemented with or
without TGF-␤ (10 ng/ml) and analyzed at the indicated times by flow
cytometry.
T cell proliferation suppression assay
Flow cytometrically purified CD4⫹CD25⫺ T cells from Thy1.1-congenic
mice (responder cells) were mixed with either nTreg or iTreg at the indicated ratios and stimulated with anti-CD3/CD28 beads at a bead:cell ratio
of 1:1. Cocultures established in quadruplicate were analyzed by flow cytometry at the indicated times. For analysis of revertant Foxp3⫺ iTreg
populations, CD4⫹Foxp3⫺ cells were converted to iTreg and sorted to
⬃99% purity as described above. These iTreg were recultured with or
without TGF-␤ for 5 days, at which time the cells were again flow-sorted
for Foxp3⫹ or Foxp3⫺ cells, respectively. The iTreg or revertant Foxp3⫺
cells were then evaluated for their ability to suppress proliferation of
CFSE-labeled naive T cell responders as previously described (29). Because the iTreg express GFP, the fluorescence emission spectrum of which
cannot readily be distinguished from CFSE, Thy1.1-disparate responder
cells were used and analysis of CFSE levels in Thy1.1-gated cells was
performed.
Cytokine production profile of stimulated nTreg, iTreg,
and non-Treg
CD4⫹Foxp3⫹ and CD4⫹Foxp3⫺ cells were purified from GFP-Foxp3
knock-in mice and cultured as described above to obtain nTreg, iTreg, and
Foxp3⫺ cells. At day 7, the expanded nTreg, iTreg, and non-Treg were
flow cytometrically sorted to obtain ⬃99% pure Foxp3⫹ or Foxp3⫺ populations. Five ⫻ 104 cells were added to 96-well plates in 250 ␮l of medium in triplicate and restimulated with anti-CD3/CD28 Ab-coated beads
at a 1:1 cell:bead ratio in the presence of 100 IU/ml rhIL-2. At 48 h, murine
IL-2, IL-4, IL-5, IL-10, IL-17, IFN-␥, and TNF-␣ were measured in the
cell culture supernatant by Bio-Plex according to the manufacturer’s instructions (Bio-Rad).
Migration and survival of iTreg after adoptive transfer
CD4⫹Foxp3⫹ and CD4⫹Foxp3⫺ cells were purified from CD45.1⫹ GFPFoxp3 knock-in mice and cultured as described above to obtain CD45.1⫹
nTreg, CD45.1⫹ iTreg, and CD45.1⫹ Foxp3⫺ cells. At days 7–9, the expanded nTreg, iTreg, and non-Treg were flow-cytometrically sorted to obtain ⬃99% pure Foxp3⫹ of Foxp3⫺ populations. Four ⫻ 106 Foxp3⫹
nTreg, iTreg, or non-Treg were mixed with 1 ⫻ 106 Thy1.1⫹ CD4⫹ T cells
before adoptive transfer by retro-orbital injection into age- and sexmatched Thy1.1⫺CD45.1⫺ C57BL/6J hosts. The Thy1.1⫹CD4⫹ T cells
were purified from a LN and splenic cell suspension from Thy1.1⫹-congenic mice by anti-CD4 magnetic bead separation using the MACS separation system according to the manufacturer’s instructions (Miltenyi Biotec). Two recipient mice for each condition and time point measured were
sacrificed at days 2, 5, 12, and 28 after injection, and peripheral blood, LN,
spleen, lung, liver, and bone marrow were isolated. Cells were collected by
forced passage through a cell strainer. Lymphoid components of liver and
lung cell suspensions were further purified by centrifugation over 37.5%
Percoll. Cells were stained with the indicated Abs before flow cytometric
analysis.
Flow cytometry
Cells were stained and analyzed on a FACSCalibur (BD Biosciences) using
CellQuest software (BD Biosciences). Quantitative flow cytometry to determine total cell numbers was performed by enumerating all cells in a
culture well. The presence of Foxp3 in cells from mice expressing the
GFP-Foxp3 fusion was determined by measuring GFP fluorescence.
Statistics
Data were analyzed by ANOVA using JMP (SAS).
Results
TGF-␤ collaborates with TCR stimulation to up-regulate Foxp3
in CD4⫹ cells
Previous studies (6 – 8, 30) have shown that TGF-␤ up-regulates
Foxp3 in activated CD4⫹CD25⫺ T cells. To directly visualize this
event, we analyzed the induction of Foxp3 in T cells from mice
engineered to express a GFP-Foxp3 fusion protein. CD4⫹Foxp3⫺
T cells were purified, stimulated, and analyzed using quantitative
flow cytometry. Similar to the results of others, TGF-␤ up-regulated Foxp3 in CD4⫹Foxp3⫺ T cells after CD3/CD28 stimulation,
The Journal of Immunology
with conversion efficiencies of 50 –90% routinely observed (Fig. 1,
A and B). Costimulation influenced the conversion. Total numbers of
Foxp3⫹ cells were increased in the presence of anti-CD28. However,
the percentage of Foxp3⫹ cells was equivalent in cultures stimulated
with anti-CD3 in the presence or absence of anti-CD28, suggesting
that costimulation acted by promoting the expansion of the cells rather
than increasing the conversion efficiency (Fig. 1B).
Foxp3 up-regulation was dependent on the presence of both
TGF-␤- and TCR-specific signaling. As has been reported, in the
absence of TGF-␤, little or no up-regulation of Foxp3 was apparent (Fig. 1, A and B, and not plotted; after anti-CD3 and anti-CD3/
CD28 stimulation, day 5 Foxp3⫹/Foxp3⫺ cell counts were 212/
95,567 and 128/141,065, respectively). In the absence of TCR
stimulation, little conversion was also observed. This effect has
been previously noted using RT-PCR analysis for Foxp3 (8). However, cell viability is extremely poor in the absence of TCR stimulation and an alternative explanation for this finding is that the
Foxp3⫹ cells have impaired survival without TCR stimulation.
Indeed, the total number of surviving T cells cultured with IL-2/
TGF-␤ was only 1.1% of that observed in cultures also stimulated
with anti-CD3/CD28. We found, however, that culture of CD4⫹
Foxp3⫺ cells in the presence of anti-CD28 but not anti-CD3
greatly improved cell viability, with similar numbers of live cells
present at day 5 as at the start of culture. Here too though, few
of the cells (mean ⫽ 1.0%) up-regulated Foxp3. Therefore TCR
but not CD28 signaling synergizes with TGF-␤ to drive Foxp3
expression. Foxp3 up-regulation was further restricted to the
naive T cell population and was not increased in isolated CD4⫹
CD44highCD45RblowFoxp3⫺ memory cells stimulated in the
presence of TGF-␤ (Fig. 1C).
TGF-␤-induced Treg suppress the proliferation of CD4⫹ target
cells
Earlier reports (7, 31) have shown iTreg, like nTreg, possess regulatory function and suppress CD4⫹ T cell proliferation in coculture experiments. We verified that our Foxp3⫹ iTreg were similarly capable of suppressing T cell expansion using quantitative
flow cytometry (Fig. 2) as well as proliferation analysis of CFSElabeled responder cells (data not shown). In both studies, iTreg
showed an efficiency similar to that of nTreg in suppressing T cell
proliferation and expansion.
Similar cytokine profile of iTreg and nTreg
Because iTreg were as efficient as nTreg in suppressing T cell
proliferation, we wanted to examine whether their cytokine production profiles were likewise comparable (Table I). Naive CD4⫹
T cells were stimulated and expanded for 7 days without TGF-␤ or
converted into iTreg with TGF-␤. nTreg were likewise expanded.
Foxp3⫹ (iTreg, nTreg) or Foxp3⫺ (non-Treg) cells were then
flow-cytometrically sorted and stimulated. Both iTreg and nTreg
demonstrated decreased production of most cytokines when compared with non-Treg, including IL-2, IL-4, IL-5, IFN-␥, and
FIGURE 1. TGF-␤ collaborates with TCR stimulation but not costimulation to up-regulate Foxp3. CD4⫹Foxp3⫺ T cells were flow-cytometrically purified and 3 ⫻ 104 distributed per well of a 96-well plate. The cells
were stimulated with anti-CD3-, CD28-, or CD3/CD28 Ab-coated beads at
a 1:1 cell:bead ratio. rhIL-2 and TGF-␤ were added at 100 U/ml and 10
ng/ml, respectively. On pretreatment day 0 or posttreatment day 5, wells
were harvested and analyzed for expression of Foxp3 by quantitative flow
cytometry. Mean pretreatment values for Foxp3⫺ and Foxp3⫹ T cells was
29,534 and 19, respectively, per well. A, Representative Foxp3 histogram
plots are shown. B, Percent Foxp3⫹ cells is plotted. Mean absolute cell
numbers are shown. C, Foxp3 is not up-regulated in memory cells. Cells
were stimulated with anti-CD3/CD28-coated beads and IL-2 with or without TGF-␤. Total cell counts on day 7 for the different populations are
listed within parentheses. B and C, Mean ⫾ SEM of quadruplicate samples
are plotted.
CORRECTOINS
FIGURE 2. iTreg suppress the proliferation of CD4⫹ target cells. Thy1.1⫹
CD4⫹CD25⫺ responder cells were mixed with congenic Thy1.1⫺ nTreg or
iTreg in 96-well plates at the indicated ratios. Cells were stimulated with
anti-CD3/CD28-coated beads at a cell:bead ratio of 1:1 for 3 days, and
wells were then analyzed by quantitative flow cytometry for viable
Thy1.1⫹ responder numbers. Mean ⫾ SEM of quadruplicate samples are
plotted. Data are representative of two independent experiments.
TNF-␣. Although more IL-2 and IFN-␥ was produced by the iTreg
than nTreg, this was significantly diminished when compared with
non-Treg. In contrast, IL-10 was strongly produced by both the
iTreg and nTreg. Therefore, TGF-␤-induced iTreg have a cytokine
profile similar to that of nTreg, with strong expression of IL-10 and
diminished expression of other effector cytokines.
Kinetics of Foxp3 expression after anti-CD3/CD28 and TGF-␤
treatment
Considering that memory T cells did not up-regulate Foxp3 in
response to TGF-␤, (Fig. 1C), we were interested in defining the
window period after activation during which T cells were susceptible to TGF-␤. Indeed, a time dependence for the generation of
regulatory cells using TGF-␤ has been previously reported (32,
33). To test for Foxp3 induction, we stimulated CD4⫹Foxp3⫺
cells with anti-CD3/CD28 and IL-2, supplementing with TGF-␤ at
different time points after stimulation (Fig. 3A). Two effects were
notable. First, when TGF-␤ was provided at the time of TCR stimulation, up-regulation of Foxp3 protein only began after an ⬃2-day
delay. Interestingly, if TGF-␤ supplementation was provided at
later time points after TCR stimulation, Foxp3 up-regulation was
delayed by a similar ⬃2-day period from the time TGF-␤ was
administered. Second, treatment with TGF-␤ beginning up to 2
days after TCR stimulation had little impact on the ultimate percentage or number of Foxp3⫹ T cells in the culture. In contrast,
cells treated with TGF-␤ on or after day 3 showed significantly
( p ⬍ 0.01) diminished conversion into Foxp3⫹ cells. When treatment began on day 3, a peak conversion of only ⬃20% of cells was
Table I. Cytokine production pattern of nTreg, iTreg, and non-Treg
cellsa
Cytokine (pg/ml)
nTreg
iTreg
non-Treg
IL-2
IL-4
IL-5
IL-10
IL-17
IFN-␥
TNF-␣
3.9
0.8
0.5
578.5
2.3
3.6
2.7
249.9
7.1
3.1
2,135.0
19.2
487.9
4.8
11,542.4
3,614.6
46.8
501.4
27.6
2,544.7
36.2
a
Five ⫻ 104 flow-cytometrically sorted cells of the indicated type from 7-day
cultures were added to 96-well plates in 250 ␮l of medium in triplicate and stimulated
with anti-CD3/CD28 Ab-coated beads at a 1:1 cell:bead ratio. Mean cytokine production measured 48 h after stimulation is shown.
FIGURE 3. Kinetics of Foxp3 expression after TGF-␤ induction. CD4⫹
Foxp3⫺ cells (2.5 ⫻ 104) were stimulated on day 0 with anti-CD3/CD28coated beads in a 96-well plate at a cell:bead ratio of 1:1 in the presence of
100 U/ml rhIL-2. TGF-␤ (10 ng/ml) was added either at day 0 or at the time
denoted. Sample wells were harvested on different days and analyzed by
quantitative flow cytometry for Foxp3 expression by measuring GFP fluorescence. Wells were split every 3– 4 days in medium with cytokine to
prevent overcrowding. Mean ⫾ SEM of percentage of Foxp3⫹ cells (A)
and total cell numbers (B) of quadruplicate samples are plotted. Data are
representative of three independent experiments.
observed compared with ⬃80% with day 0 treatment. This difference did not result from an outgrowth of Foxp3⫺ T cells because
total cell numbers were similar in the different treatment groups
(Fig. 3B). Indeed, quantitative analysis demonstrated that absolute
numbers of Foxp3⫹ cells were significantly ( p ⬍ 0.01) higher in
the cells treated with TGF-␤ starting days 0 –2 compared with the
cells treated after day 2. Therefore, TGF-␤/TCR stimulation has a
limited window during which it can up-regulate Foxp3, and TGF-␤
supplementation leads to up-regulation of Foxp3 protein only after
a significant (⬃2-day) lag period.
TGF-␤- and CD3-induced Foxp3 up-regulation is independent
of cell cycling
Differentiation of naive T cells into Th1 and Th2 cell types occurs
only after multiple rounds of cell cycling, an event believed to be
required to relieve epigenetic repression of lineage-specific genes
(34). After a T cell is stimulated through the TCR, it begins to
cycle ⬃2 days after stimulation, consistent with the time frame for
TGF-␤-mediated up-regulation of Foxp3 (Fig. 3A). We were therefore interested whether Foxp3 up-regulation only occurred in T
cells that had divided. To test this, we labeled CD4⫹Foxp3⫺ cells
with the cell membrane-associated red fluorescent dye PKH26 and
stimulated them with anti-CD3/CD28 with or without TGF-␤ (Fig.
4). The fluorescence of PKH-26 is diminished with each cell division. In the absence of TGF-␤, Foxp3 expression was not observed in divided or undivided cells. In contrast, Foxp3 was upregulated in all cell populations treated with TGF-␤, including
The Journal of Immunology
FIGURE 4. TGF-␤-induced Foxp3 up-regulation is independent of cell cycling. Purified CD4⫹Foxp3⫺ cells were labeled with PKH26, washed, and
stimulated with anti-CD3/CD28-coated beads at a cell:bead ratio of 1:1 in the
presence of 100 U/ml rhIL-2 with or without 10 ng/ml TGF-␤. Flow cytometric analysis of cells immediately after culture (day 0) and at days 2 and 3
are shown. Each condition was analyzed in triplicate with essentially identical
results. Data are representative of two independent experiments.
those that had not divided. This demonstrates that Foxp3 up-regulation and iTreg development does not require cell cycling.
Limited persistence of Foxp3 in iTreg after TGF-␤ withdrawal
To determine whether iTreg require continuous exposure to
TGF-␤ to preserve Foxp3 expression, we stimulated CD4⫹
Foxp3⫺ cells with anti-CD3/CD28 and TGF-␤, and withdrew
TGF-␤ at various time points after stimulation (Fig. 5A). Foxp3
expression persisted at high levels in cells that received continuous
TGF-␤ treatment for the 13 days of culture. In contrast, Foxp3
expression returned to baseline by day 13 in cells that had TGF-␤
removed after initial treatment. After TGF-␤ was withdrawn,
Foxp3 persisted for another 3– 4 days, at which point expression
began to decrease ( p ⫽ 0.013). Expression was virtually undetectable in the T cells by 9 days after TGF-␤ withdrawal. Cells that
were exposed to ⬍3 days of TGF-␤ lost Foxp3 at a much higher
rate than those exposed for at least 3 days. Because total cell numbers in the wells were similar with or without withdrawal of
TGF-␤ and total numbers of cells steadily increased with culture
time (Fig. 5B), the loss of Foxp3 seemed to be primarily due to the
conversion of the Foxp3⫹ cells to Foxp3⫺ cells rather than cell
death. This conversion was confirmed in separate experiments in
which iTreg purified by flow cytometric sorting for GFP-Foxp3
and then cultured without exogenous TGF-␤ displayed a similar
loss of Foxp3 (data not shown). When sorted Foxp3⫹ iTreg were
allowed to lose Foxp3 expression, Foxp3 could not be reinduced
by restimulation, even in the presence of TGF-␤ (data not shown).
Therefore, Foxp3 can only be induced in a brief window as naive
T cells differentiate into effector/memory cells. Foxp3 persistence
is dependent on sustained TGF-␤ signaling.
Loss of suppressive potency in iTreg that have down-regulated
Foxp3
Considering that removal of TGF-␤ resulted in loss of Foxp3 in
iTreg (Fig. 5A), we were interested in whether loss of Foxp3 also
FIGURE 5. Limited persistence of Foxp3 in iTreg after TGF-␤ withdrawal. Purified CD4⫹Foxp3⫺ cells were stimulated with anti-CD3/CD28coated beads at a cell:bead ratio of 1:1 in the presence of 100 U/ml rhIL-2.
No or 10 ng/ml TGF-␤ was added at day 0. TGF-␤ was removed by
washing the cells at the indicated time. To prevent overcrowding during the
extended culture, cells were split every 3– 4 days into medium with IL-2
and with or without TGF-␤. Cells were analyzed by quantitative flow cytometry at the indicated time points. Mean ⫾ SEM of percentage of
Foxp3⫹ cells (A) and total cell numbers (B) of quadruplicate samples are
plotted. Data are representative of three independent experiments.
leads to a loss of suppressive activity. To test this, TGF-␤ was
either added to or excluded from cultures of flow-cytometrically
purified Foxp3⫹ iTreg. Five days later, Foxp3⫹ or Foxp3⫺ cells
from the respective cultures were flow-cytometrically isolated. The
iTreg or revertant Foxp3⫺ cells were then added to naive, Thy1.1disparate, CFSE-labeled T cells, and the proliferation of the naive
population to anti-CD3/CD28 was measured by loss of CFSE (Fig.
6). As in Fig. 2, iTreg that retained Foxp3 strongly suppressed
naive cell proliferation. In contrast, iTreg that lost Foxp3 showed
a substantially reduced ability to suppress T cell proliferation.
Thus, loss of Foxp3 expression is accompanied by a loss of suppressive activity.
Migration and survival of iTreg after adoptive transfer
Our in vitro studies suggested that the persistence of iTreg is dependent upon exogenous TGF-␤ and that Foxp3 is lost within a
several-day period after TGF-␤ removal. The transient nature of
Foxp3 expression in iTreg implies that iTreg would not be suitable
for immunotherapeutic application. Yet, studies have now documented that iTreg are effective in treating model alloimmune and
autoimmune diseases (23–25). To determine whether iTreg persist
in vivo, we adoptively transferred flow-cytometrically purified
GFP-Foxp3⫹ iTreg derived from CD45.1⫹ mice into CD45.1⫺
congenic recipients and followed their migration and survival (Fig.
7). As controls, equivalent numbers of either Foxp3⫺ cells from
CORRECTIONS
FIGURE 6. Suppression of naive T cell proliferation by Foxp3⫺ revertant iTreg. Thy1.1⫺ iTreg were flow-cytometrically sorted for Foxp3⫹ cells
7 days after induction with TGF-␤. These cells were recultured for 5 days
with or without TGF-␤. The cells grown with TGF-␤ were then sorted
again for expression of Foxp3, whereas the revertant Foxp3⫺ cells were
sorted from the population grown in the absence of TGF-␤. Thy1.1⫹ naive
T cells were CFSE labeled, mixed with the Foxp3⫹ or Foxp3⫺ populations
at the indicated ratios, and stimulated with anti-CD3/CD28-coated beads.
At 72 h, cultures were stained for Thy1.1, and CFSE expression on the
Thy1.1⫹ population was analyzed by flow cytometry.
cultures stimulated in the absence of TGF-␤ or Foxp3⫹ nTreg
from similarly expanded cultures were also transferred into naive mice. The CD45.1-congenic background permitted identification of the transferred cells regardless of their expression of
GFP-Foxp3. To control for adoptive transfer efficiency among
mice, control freshly purified Thy1.1⫹CD4⫹ cells were admixed at a 1:4 ratio with each of the adoptively transferred cell
populations before transfer into congenic Thy1.1⫺CD45.1⫺
C57BL/6 hosts (Fig. 7C).
At selected time points mice were sacrificed, cell suspensions
were prepared from different organs, and the cells were analyzed
by flow cytometry for CD4, Thy1.1, CD45.1, and Foxp3 expression. Similar numbers of Thy1.1⫹ cells were routinely observed in
the recipient mice, indicating equivalent adoptive transfer efficiencies (data not shown). CD4⫹ cells were gated to distinguish the
CD45.1⫹ transferred population and CD45.1⫺ host cells, and the
transferred cells were then further segregated into Foxp3⫹ and
Foxp3⫺ populations (Fig. 8, A and B). Numbers of CD45.1⫹
Foxp3⫹ or Foxp3⫺ cells were normalized to endogenous CD4⫹
CD45.1⫺Thy1.1⫺ cell numbers and plotted (Fig. 8C).
FIGURE 7. Migration and survival of iTreg after adoptive transfer. A,
Scheme for the preparation and adoptive transfer of iTreg, nTreg, or nonTreg. B, Flow cytometric analysis of expanded iTreg, nTreg, and non-Treg
populations before flow purification. C, Flow cytometric analysis of adoptively transferred cell populations, which include purified CD4⫹Thy1.1⫹
injection controls and CD4⫹CD45.1⫹Thy1.1⫺Foxp3⫹ or ⫺ experimental
populations. Data are representative of three independent experiments.
Transferred flow-cytometrically purified Foxp3⫹ iTreg largely
disappeared within the first 2 days after adoptive transfer (Fig. 8C).
This seemed to result from down-modulation of Foxp3 since large
numbers of CD4⫹CD45.1⫹Foxp3⫺ cells were simultaneously observed in several organs, including the spleen, liver, blood, and
lung. Indeed, in support of this interpretation, numbers of Foxp3⫺
cells detected after transfer of purified Foxp3⫹ iTreg were similar
to those observed after adoptive transfer of equal numbers of purified CD4⫹Foxp3⫺ non-Treg T cells. Transferred CD4⫹Foxp3⫺
cells were also found in similar locations as Foxp3⫺ former iTreg,
specifically the spleen, liver, blood, and lung, within the first week
after adoptive transfer. By 4 wk after transfer, virtually all of the
Foxp3⫺ cells from either the iTreg or control non-Treg transfer
had disappeared. One organ where Foxp3 was retained on the
transferred iTreg was the LN. Indeed, of the organs analyzed at
day 2 after iTreg transfer, greater numbers of transferred Foxp3⫹
cells then Foxp3⫺ cells were only detected within the LN.
By 2– 4 wk after iTreg transfer, when the revertant Foxp3⫺ cells
had largely disappeared, increasing numbers of transferred
Foxp3⫹ cells were observed, particularly in the bone marrow and
LN. Interestingly, only small numbers of Foxp3⫹ cells remained in
the spleen, suggesting differential localization or inadequate support of Foxp3 expression at this site. Because the transferred
The Journal of Immunology
FIGURE 8. Migration and survival of iTreg, nTreg, and non-Treg after adoptive transfer. A, iTreg, nTreg, and non-Treg prepared as in Fig. 7 were
adoptively transferred by retro-orbital injection into C57BL/6J mice. After 2 days, the mice were sacrificed, and cells were isolated and stained with CD45.1,
Thy1.1, and CD4. Sample flow cytometry plots from forward scatter/side scatter-gated LN cells are shown, demonstrating identification of the transferred
CD4⫹CD45.1⫹ cells among endogenous CD45.1⫺ cells. Numbers in the right side quadrants indicate percentage of CD45.1⫹ and CD45.1⫺ cells among
the total CD4⫹ population. B, Histogram plots of gated CD4⫹CD45.1⫹ cells (oval gates in A) analyzed for the expression of Foxp3 are shown. C, Mice
were sacrificed at the indicated time points and analyzed for the presence of adoptively transferred cells. Numbers of adoptively transferred CD4⫹CD45.1⫹
Foxp3⫹ or Foxp3⫺ cells were normalized to 25,000 host CD4⫹CD45.1⫺Thy1.1⫺ cells in the indicated organ and plotted. Mean ⫾ SEM of two mice per
condition are plotted. Data are representative of three independent experiments.
CD45.1⫹ cells were ⬃99% Foxp3⫹ (Fig. 7C), this indicates that a
subset of iTreg persisted and possibly expanded, surviving for at
least 4 wk and primarily localizing within the bone marrow
and LN.
The survival and localization of iTreg showed significant differences though also similarities with that of adoptively transferred
preactivated nTreg. nTreg retained their Foxp3 to a much greater
extent than iTreg. All organs analyzed showed a predominance of
Foxp3⫹ cells after nTreg transfer, yet this was only observed for
the LN after iTreg transfer. Similar to the Foxp3⫹ cohort after
iTreg transfer, the nTreg persisted 4 wk after transfer and at late
time points also resided predominantly in the LN and bone marrow. One additional feature of nTreg transfers was that despite the
transfer of equivalent numbers of nTreg as non-Treg, fewer nTreg
were detected in the different organs than non-Treg. For example
⬃3,000 and 4,000 non-Treg were identified in the LN and spleen
per 25,000 endogenous CD4⫹ T cells on day 2 after transfer,
whereas ⬃400 and 130 nTreg were detected, respectively. This
may reflect an increased rate of death or decreased proliferation of
the preactivated nTreg vs non-Treg after transfer, or alternatively
differential migration to other organs not analyzed. In summary,
transferred Foxp3⫹ iTreg yielded large numbers of Foxp3⫺ cells
that entered the blood, lungs, and liver and then disappeared by
2– 4 wk after transfer. A portion of iTreg, however, retained
Foxp3. These persevered initially in the LN and then, as for nTreg,
increasingly in the bone marrow. Thus, although many iTreg act in
vivo as in vitro, losing Foxp3 after removal from TGF-␤, a subset
of these cells persist longer as Foxp3⫹ iTreg.
Transferred iTreg phenotypically resemble nTreg
nTreg constitutively express several T cell activation markers, including CD25 and glucocorticoid-induced TNFR family-related
gene (GITR) (35). These markers were also nearly uniformly upregulated on both iTreg and activated non-Treg before adoptive
transfer (Table II).
Table II. Surface marker expression on iTreg, nTreg, and non-Treg
cellsa
Percentage of Cells
Naive
In vitro stimulated
Surface
Marker
nTreg
Foxp3⫺
nTreg
iTreg
Non-Treg
GITR
CD25
CD62L
CXCR4
89.9
88.6
38.2
2.2
2.2
5.1
55.2
1.3
100
99.3
83.2
2.9
98.3
96.6
26.0
2.8
99.9
98.8
29.8
1.9
a
Freshly isolated CD4⫹Foxp3⫺ or Foxp3⫹ cells were stimulated as described for
7– 8 days to obtain Foxp3⫹ iTreg, Foxp3⫺ non-Treg, and Foxp3⫹ nTreg. Expression
of the indicated markers before and after culture is shown.
CORRECTIONS
FIGURE 9. Transferred iTreg phenotypically resemble nTreg. CD4⫹
CD45.1⫹ adoptively transferred iTreg, nTreg, and non-Treg were identified within the LN of recipient mice and analyzed for Foxp3 expression as
in Figs. 7 and 8. These were additionally analyzed for surface expression
of CD25 (A) or GITR (B). Percentage of CD4⫹CD45.1⫹ cells of the indicated cell type positive for CD25 or GITR is plotted. Data points are
missing for some cell types/conditions at some time points due to an inadequate number of cells in the organs at those times for definitive expression analysis. Sample histogram plots from the day 2 time point of transferred iTreg populations in LN that lost Foxp3 (left) or retained Foxp3
(right) are also shown. Mean ⫾ SEM of two mice per condition are plotted.
Data are representative of two independent experiments.
To determine whether expression of CD25 and GITR persisted after adoptive transfer, cells from mice receiving activated and expanded iTreg, nTreg, or non-Treg were analyzed
for CD25 and GITR expression. Foxp3⫹ iTreg, like nTreg, retained CD25 and GITR expression in the LN (Fig. 9, A and B)
and all other organs analyzed (data not shown) throughout the
28-day time course of the experiment. In contrast, the majority
FIGURE 10. CXCR4 is preferentially expressed on adoptively transferred cells in the bone marrow and CD62L on cells in the LN. Data were
acquired and analyzed as in Figs. 7 and 8. A, Percentage of indicated
adoptively transferred populations positive for CXCR4 on day 5 after
transfer is plotted. Sample histogram plots demonstrating CXCR4 expression on Foxp3⫹ iTreg in the LN or bone marrow (BM) are also shown. B,
Percentage of the indicated adoptively transferred population positive for
CD62L on day 5 or day 12 after transfer is plotted. Sample histogram plots
demonstrating CD62L expression on Foxp3⫹ iTreg in the LN or bone marrow
are also shown. Mean ⫾ SEM of two mice per condition are plotted. Data are
representative of two independent experiments. ND, No data.
of either transferred non-Treg or revertant iTreg-derived
Foxp3⫺ cells lost CD25 and GITR expression within the first
2–5 days after transfer. The small fraction of nTreg that lost
The Journal of Immunology
Foxp3 (Fig. 8) also lost surface expression of CD25 and GITR
(data not shown). This indicates that Foxp3⫹ iTreg phenotypically resemble nTreg in vivo, while the revertant Foxp3⫺ iTreg
resemble Foxp3⫺ cells. Foxp3 protein expression is associated
with the sustained expression of CD25 and GITR in vivo.
It was interesting that the greatest concentration of transferred
activated Foxp3⫹ iTreg or nTreg were found in the bone marrow
and LN. Bone marrow localization of transferred nTreg has been
previously shown to be associated with expression of the CXCR4
chemokine receptor, which is also expressed on other cell types
localizing within the marrow (36). Indeed, blockade of CXCR4
can mobilize cells from the marrow into the blood (37). Analogously, nTreg expression of CD62L, a receptor for glycan ligands
expressed on high endothelial venules, is associated with LN homing of nTreg as well as other T cells (38). To determine whether
receptor expression correlates with tissue localization of iTreg, as
it does for nTreg, we analyzed CXCR4 and CD62L expression on
the transferred cells. At the time of transfer, ⬍3% of iTreg expressed CXCR4 and only 26% expressed CD62L (Table II). By 5
days after transfer, ⬃90% of Foxp3⫹ iTreg or nTreg in bone marrow expressed CXCR4, while the same population in other organs
showed very little CXCR4 expression (Fig. 10A). Similar results
were seen at later time points (data not shown). This is consistent
with CXCR4 being the bone marrow homing receptor for iTreg, as
for nTreg.
At day 5, CD62L was discriminately expressed on cells within
the LN; however, the differential expression was not as dramatic as
for CXCR4 (Fig. 10B). By day 12, segregation of cells based on
CD62L expression was more prominent. Approximately 80% of
LN Foxp3⫹ cells (both nTreg and iTreg) expressed CD62L at day
12 (Fig. 10C), whereas only ⬃10% of Foxp3⫹ cells in the bone
marrow expressed CD62L. This indicates that the CD62L⫹
Foxp3⫹ iTreg population preferentially localizes to LN. Thus,
adoptively transferred iTreg segregated into populations with distinct localization patterns that correlate with the expression of specific chemokine receptors or adhesion molecules.
Discussion
It is now well established that TGF-␤ can up-regulate Foxp3 in
CD4⫹Foxp3⫺ T cells. Using mice in which a GFP-Foxp3 fusion
protein has been homologously inserted into the Foxp3 locus, we
have been able to assess on a single-cell basis the kinetics of upregulation of Foxp3 in vitro, as well as the maintenance of Foxp3
expression in vitro and in vivo. Our in vitro data are consistent
with and adds to previous results. Foxp3 protein was up-regulated
only after ⬃48 h of TGF-␤ treatment. This lag period in single-cell
protein expression corresponds to a similar ⬃48-h lag in Foxp3
mRNA expression observed in human T cells (7). TGF-␤-induced
Smad activation and nuclear translocation occurs within minutes,
and SMAD-DNA complexes can be observed as early as 10 min
after TGF-␤ addition (39). This ⬃2-day delay suggests that Foxp3
up-regulation involves a complex developmental program rather
than an immediate effect of SMAD-DNA interactions. Indeed, recent identification of the involvement of CTLA-4 and cbl-b in
TGF-␤-induced Foxp3 up-regulation (40, 41) supports the idea
that complex combinatorial or sequential signals are involved in
Foxp3 induction.
We further find that the lag in Foxp3 expression is independent
of the time from initial TCR stimulation as when TGF-␤ is added
24 or 48 h after stimulation, Foxp3 up-regulation shows an identical ⬃2-day delay. This implies that a TCR-dependent factor is
not rate-limiting in inducing Foxp3 protein expression, but rather
TGF-␤ induces a sequence of events requiring this time frame
to up-regulate Foxp3. Up-regulation does not require T cell cycling
and is therefore unlikely to involve cell cycle-dependent
modifications.
We find that TCR stimulation is needed for Foxp3 expression.
Viability of naive T cells cultured in the absence of TCR stimulation is poor and this viability loss may prevent examination of
Foxp3 up-regulation in unstimulated populations. By stimulating
cells with anti-CD28 in the absence of anti-TCR Ab, we were able
to preserve viability and thereby clearly demonstrate this requirement. At 48 –72 h after TCR stimulation, however, T cells become
refractory to the effects of TGF-␤. Therefore, a TCR-induced program is initially required for, although later restricts the ability of
TGF-␤ to up-regulate Foxp3.
Previous studies have led to different conclusions on the role of
CD28 costimulation in Foxp3 up-regulation. One study showed a
marked diminishment in Foxp3 mRNA up-regulation after TGF-␤
induction of peripheral T cells stimulated in the presence of antiCD28 Abs (7). In contrast, a second study found that CD28 costimulation was important for Foxp3 expression in TGF-␤-treated
CD4⫹CD25⫺ thymocytes, primarily by enhancing cell survival
(42). In multiple experiments using highly purified Ab-stimulated
T lymphocytes, we failed to observe either a beneficial or deleterious role for CD28 costimulation in Foxp3 up-regulation. The
percentage conversion of Foxp3⫺ T cells to Foxp3⫹ cells was
similar regardless of the presence of costimulation. However, costimulation increased total cell numbers of both Foxp3⫺ and
Foxp3⫹ cells, suggesting that in our system it primarily promotes
cell survival or proliferation rather than conversion per se. One
important role of CD28 costimulation is the induction of IL-2 production in activated T cells (43). Since our cultures contained exogenous IL-2, which is an important facilitator of Foxp3 induction,
this role of costimulation on Foxp3 up-regulation may have been
masked in our system. Indeed, recent studies have emphasized the
requisite role for IL-2 in Foxp3 up-regulation and in one case
supported a role for CD28-induced IL-2 in this regard (44, 45).
We further analyzed the sustainability of Foxp3 after removal of
TGF-␤. Interestingly, continued Foxp3 expression in vitro requires
continued exposure of iTreg to exogenous TGF-␤. Foxp3 is lost
from cells beginning several days after TGF-␤ removal and is
eventually fully lost. Whether endogenous TGF-␤ expression can
replace this exogenous TGF-␤ in sustaining Foxp3 is unclear. In
our system, this was not the case. Data showing that TGF-␤-induced T cells can educate naive T cells to develop suppressive
properties in a TGF-␤- and IL-10-dependent manner, however,
suggest that under some conditions a self-sustaining process promoting continued Treg development may take place (5). In those
experiments, persistent Ag stimulation was required to sustain regulatory function and Foxp3 expression, and it may be hypothesized
that this difference in stimulation conditions results in the induction of TGF-␤.
The down-regulation of Foxp3 in iTreg was accelerated when
the iTreg were adoptively transferred in vivo. By day 2 after transfer, few of the cells retained Foxp3. Interestingly, the tissue localization of Foxp3⫺ former-iTreg was similar to that of transferred
control Foxp3⫺ T cells and different from that of transferred
Foxp3⫹ nTreg (Fig. 8). The reversion of iTreg into Foxp3⫺ T cells
has potentially significant implications for their therapeutic utility
because it may limit the intrinsic regulatory activity of the transferred cells. Indeed, we observed that iTreg that lost Foxp3 had
diminished regulatory activity in a T cell proliferation suppression
assay. In contrast to transferred iTreg, transferred activated nTreg
showed limited loss of Foxp3 expression. Therefore iTreg and
nTreg, although similar in suppressive function in vitro, differentially preserve Foxp3 after adoptive transfer.
CORRECTIONS
Maintenance of nTreg requires several factors, including TGF-␤
and IL-2, in vivo (13, 46, 47). nTreg that lack IL-2R or that are
from TGF-␤⫺/⫺ mice have a diminished peripheral life span. The
differential maintenance of Foxp3 in nTreg and iTreg may suggest
that transferred nTreg and iTreg have different access to sites
where these cytokines are produced. An alternative or complementary possibility may lie in the distinct TCR repertoire of iTreg and
nTreg. nTreg contain a skewed representation of TCR that is biased toward high affinity for self when compared with Foxp3⫺ T
cells (27, 48). This self-specificity may alter the interactions of
nTreg with resident APC compared with Foxp3⫺ T cells, leading
to the distinct homeostatic properties of the different cell types.
Because iTreg are derived from Foxp3⫺ T cells, we conjecture that
they will lack the inherent self-specificity present in nTreg. Further
studies analyzing the repertoire of stable Foxp3⫹ iTreg vs revertant Foxp3⫺ cells are ongoing to explore this possibility.
Despite the loss of Foxp3 in the majority of transferred iTreg, a
proportion of cells maintain Foxp3. Interestingly these cells concentrated primarily in two sites, the bone marrow and the LN. The
number of transferred Foxp3⫹ cells increases in these locations
over a 4-wk time period, suggesting that they expand there. Indeed,
by day 28 after transfer, the numbers of Foxp3⫹ iTreg in the bone
marrow and LN are equivalent to those of similarly transferred
nTreg. Cellular localization of Foxp3⫹ T cells follows adhesion
molecule expression. CXCR4, which is associated with bone marrow localization in a variety of cell types, is prominently expressed
in the bone marrow resident population. Indeed, recent data have
demonstrated that purified and transferred CXCR4⫹ nTreg primarily localize to the bone marrow (49). Likewise, we find that the
majority of Foxp3⫹ iTreg inhabiting the LN are CD62Lhigh, a
marker with a well-established role in LN homing (38).
It would be anticipated that the distribution of adoptively transferred iTreg should mimic that of endogenous Treg. Endogenous
Treg freely distribute, although modestly different proportions may
be detected in different lymphoid compartments. However, even 1
mo after transfer, we observed a skewed distribution of transferred
cells to the LN and bone marrow. Our data on CXCR4 and CD62L
expression suggest that this is due to preferential homing and localization to these organs; however, preferential survival or expansion of Foxp3⫹ cells in these sites cannot be excluded. Interestingly, few nTreg or iTreg expressed CXCR4 before transfer
(Table II). Whether de novo up-regulation of this receptor occurred
in vivo or the localization of the transferred cells was predetermined by their receptor expression after in vitro stimulation will be
important to examine. The homing and localization of adoptively
transferred therapeutic populations may impact their therapeutic
qualities and understanding these processes will be important for
any future application of in vitro-expanded regulatory cells.
Like nTreg, iTreg that retained Foxp3 after adoptive transfer
also maintained CD25 and GITR surface expression. Our results
suggest that these Foxp3⫹ cells, which were primarily found in the
LN and bone marrow, play an important role in the amelioration of
immunopathologic diseases after iTreg transfer. However, defining
the role of the different cell populations in regulating disease will
be important and it remains to be determined whether revertant
Foxp3⫺ cells have regulatory properties as well. Analyzing this
may be complex since the transferred cells may not uniquely possess regulatory function. In one study, transferred iTreg were able
to induce regulatory activity in endogenous populations of T cells
(23). We have observed a similar infectious tolerance with transferred nTreg (50).
In summary, we demonstrate that iTreg are dependent on TGF-␤
signaling for the preservation of Foxp3 expression. Loss of Foxp3
occurs rapidly after in vivo transfer, with a predominance of
Foxp3⫺ cells appearing within 2 days. The brief life span of Foxp3
in most transferred iTreg contrasts with that of transferred nTreg,
which largely preserve Foxp3 expression. Although a subset of
Foxp3⫹ iTreg is maintained, the distinct cellular dynamic properties of the iTreg and nTreg populations may lead to differential
effects on immune responses, and this, in addition to the impact of
the large numbers of Foxp3⫺ cells forming after iTreg transfer,
must be considered in potential applications of the different cell
types. iTreg, although phenotypically homogeneous after expansion in vitro, further segregate into different populations based on
their expression of different homing/adhesion molecules, including
CXCR4 and CD62L. Deciphering the role of the multiple cell
types generated after iTreg transfer in the induction and preservation of immune tolerance will be important for understanding their
therapeutic potential.
Acknowledgments
We thank Richard Cross and Jennifer Smith for assistance with flow cytometry, cell sorting, and Bio-Plex assays; Phuong Nguyen and Rajshekhar
Alli for technical support; and Sasha Rudensky for providing the GFPFoxp3 knock-in mice.
Disclosures
The authors have no financial conflict of interest.
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