© 2007 Nature Publishing Group http://www.nature.com/natureimmunology
ARTICLES
Origin of dendritic cells in peripheral lymphoid organs
of mice
Kang Liu1, Claudia Waskow1, Xiangtao Liu2, Kaihui Yao1, Josephine Hoh3 & Michel Nussenzweig1,4
Parabiosis experiments demonstrating that dendritic cells (DCs) do not equilibrate between mice even after prolonged joining by
parabiosis have suggested that DCs are derived from self-renewing progenitors that divide in situ. However, here we found that
unequal exchange of DCs between mice joined by parabiosis reflected uneven distribution of DC precursors in blood due to their
short half-life in circulation. DCs underwent only a limited number of divisions in the spleen or lymph nodes over a 10- to
14-day period and were replenished from blood-borne precursors at a rate of nearly 4,300 cells per hour. Daughter DCs
presented antigens captured by their progenitors, suggesting that DC division in peripheral lymphoid organs can prolong the
duration of antigen presentation in vivo.
Lymphoid organs contain subsets of dendritic cells (DCs) with distinct
functions1–3. In the spleen, the main conventional DC (cDC) subsets
with CD8+DEC-205+ or CD8–33D1+ cell surface protein expression
differ in their localization in the spleen, surface expression of antigen
and capacity to process and present antigens4–7. Despite those notable
differences, bone marrow transfer experiments with irradiated recipient mice indicate that all cDC subsets can be derived from a
common clonogenic myeloid progenitor that also gives rise to
macrophages; this progenitor is defined by an absence of lineage
markers (Lin–) and expression of CX3CR1 and c-Kit8–10. Plasmacytoid
DCs (pDCs) differ from cDCs in that they produce large amounts of
type I interferons through signaling by Toll-like receptors 7 and 9 and
they originate from distinct estrogen-resistant myeloid progenitors in
the bone marrow11,12.
Monocytes can give rise to cells with many of the phenotypic and
functional features of DCs when cultured in vitro with the appropriate
cytokines13–15 or in inflammatory conditions in vivo16–19. In addition,
monocytes seem to give rise to DCs in the skin and lungs18,20,21.
However, in the steady state, spleen and lymph node DCs do not seem
to be derived from monocytes, and in humans, circulating immature
DCs can be distinguished from monocytes2,17,19.
DCs were initially thought to be end-stage, nondividing cells. Rapid
incorporation of bromodeoxyuridine (BrdU) by DCs in vivo has been
attributed to dividing DC precursors labeled in the bone marrow that
give rise to short-lived (half-life, about 1.5–2.9 d) nondividing DCs in
the spleen and lymph nodes22–24. In contrast, subsequent experiments
have shown that DCs can divide for prolonged periods in vitro25 and
that in vivo about 5% of the DCs in the spleen are in the cell cycle at
any time26. Furthermore, only a fraction of the DC pool in parabiotic
mice is replaced after 6 weeks of shared circulation, suggesting that
dividing spleen DCs have the potential to maintain the DC pool by
in situ replication26. Therefore, the assumptions used to calculate the
turnover of the DC pool in peripheral lymphoid organs are no longer
applicable. Moreover, to our knowledge, the self-renewal capacity of
the DCs in spleen has not been measured directly.
Here we report data from and our conclusions regarding parabiotic
experiments designed to examine the origin and turnover of DCs in
peripheral lymphoid organs. We found that spleen and lymph node
DCs were continually replaced by blood-borne precursors that did not
equilibrate between mice joined by parabiosis (‘parabionts’) because
they were rapidly cleared from circulation. DCs underwent a limited
number of divisions in spleen and lymph node and were entirely
replaced within 10–14 d. Daughters of proliferating DCs retained
antigen as complexes of peptide and major histocompatibility complex (pMHC) that probably function to extend the duration of
antigen presentation.
RESULTS
DCs divide in situ in peripheral lymphoid organs
To examine the turnover of DCs in spleen and lymph nodes, we
measured BrdU uptake by DCs, and to analyze cell cycle distribution
among DCs, we stained cells with 4,6-diamidino-2-phenylindole
(DAPI) and for expression of the nuclear antigen Ki-67, which
indicates proliferation27. We found that 53.7% of CD8+DEC-205+
and 45.4% of CD8–33D1+ DCs in spleen and lymph nodes were
labeled with BrdU within 2 d, and these percentages increased to
91.7% and 71.6%, respectively, by day 5 of continuous BrdU supply22,23 (Fig. 1a and Supplementary Fig. 1 online). Staining with
DAPI and antibody to Ki-67 (anti-Ki-67) showed that 5% of splenic
CD11c+ cDCs were actively dividing at any given time25,26 (Fig. 1b).
1Laboratory of Molecular Immunology, The Rockefeller University, New York, New York 10021, USA. 2Department of Applied Mathematics and 3Department of
Epidemiology and Public Health, Yale University, New Haven, Connecticut 06520, USA. 4Howard Hughes Medical Institute, The Rockefeller University, New York,
NY 10021, USA. Correspondence should be addressed to M.N. ([email protected]).
Received 23 January; accepted 3 April; published online 22 April 2007; doi:10.1038/ni1462
578
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ARTICLES
pDCs
c
DCs
36.4
71.6
BrdU
5d
91.7
cDCs
0
Isotype
b
0
58.2
0
70.2
74.3
99.5
1.57
Graft
spleen
BrdU
pDCs
1.4
0.11
Host
CD8α
PDCA
Host
spleen
Host Graft
53.7
CD45.1
45.4
Graft DCs
99.8
1.33
CD3, CD19, NK1.1, Ter119
6.77
Host
Control
spleen
0.80
BrdU
2d
Host DCs
0
0.07
0.94
BrdU
0d
29.7
68.5
0.34
0
CD11c
CD45.2
BrdU
d
0.3
5.0
Ki-67
© 2007 Nature Publishing Group http://www.nature.com/natureimmunology
CD8+DCs
CD11c
CD8–DCs
a
Isotype
Ki-67
CD80
CD40
CD86
MHC I
MHC II
CD1d
+
Ki-67–
DAPI
Figure 1 Rapid turnover of dividing DCs in the spleen and lymph nodes. (a) Flow cytometry of BrdU uptake by CD8+ or CD8– DCs or PDCA+ DCs after 0, 2
or 5 d of continuous labeling. Numbers adjacent to outlined areas indicate percent BrdU+ cells. (b) Flow cytometry of Ki-67 versus DAPI staining of purified
spleen pDCs and cDCs. Numbers in top right quadrants indicate percent Ki-67+ dividing cells in S-G2-M phase (DAPIhi). (c) Flow cytometry of BrdU uptake
in grafted spleen at 8 d after transplantation. Left, gating on CD11chiCD3–CD19–NK1.1–Ter119–. ‘DC’ plots, percent host CD45.2+ and donor F1 DCs in
control spleen, recipient spleen and graft. Host DC and Graft DC plots, BrdU uptake. (d) Flow cytometry of CD40, CD80, CD86, MHC class I (MHC I), MHC
class II (MHC II) and CD1d staining after gating on CD11chiCD3–B220–NK1.1– Ki-67+ or Ki-67– DCs in spleen. Numbers in plots indicate percent gated
cells. Data are from one representative of two to three experiments.
In contrast, only 0.3% of the pDCs were in the cell cycle (Fig. 1b).
We obtained similar results for DCs in lymph nodes (Supplementary
Fig. 2 online).
To determine whether the dividing cells were derived from blood or
tissue, we transplanted pieces of spleen from mice expressing allotypically distinct cell surface markers (CD45.1 CD45.2 F1 mice) into
CD45.2 recipient mice and measured BrdU incorporation by F1 graft
DCs at 8 d after transplantation. Because there were no F1 graft DC
precursors in the CD45.2 recipient bone marrow or blood, F1 DCs in
the grafted spleen pieces that incorporated BrdU must have been graft
derived and hence were dividing in situ. F1 DCs in the grafted spleen
pieces incorporated BrdU at the same rate as did CD45.2 host DCs in
CD45.2 recipient spleen (Fig. 1c). The phenotype of the dividing DCs
was indistinguishable from that of other DCs in terms of the
expression of cell surface molecules involved in antigen presentation
and T cell costimulation (Fig. 1d). We conclude that mature DCs can
divide in peripheral lymphoid organs.
Unequal exchange of DCs between parabionts
To examine the kinetics and origins of DC replacement, we established
parabiosis between mice expressing the two distinct CD45 allotypes,
CD45.1 and CD45.2 (Fig. 2). The exchange of blood-borne cells or
proteins between parabiotic mice is directly related to their half-lives in
the circulation28–30. Cells and proteins with a long half-life in circulation are equally distributed in the two partners, whereas those with a
short half-life do not28–31. For example, it takes approximately
2 h for soluble dyes to reach equilibrium in both partners31, and
leptin, which has a short half-life in circulation, never reaches equilibrium30. In the parabiotic mice, B cells, T cells and stationary
NATURE IMMUNOLOGY
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monocytes in blood were completely exchanged by day 9 after the initiation of parabiosis, although B cells and polymorphonuclear leukocytes
were differentially equilibrated between CD45.1 and CD45.2 parabionts
(Fig. 2a and Supplementary Fig. 1; average chimerism, 45–50%).
Similarly, B lymphocytes and T lymphocytes in the spleen and lymph
nodes were freely exchanged between partners, reaching a plateau 9 d
after the initiation of parabiosis (Fig. 2c,d). In contrast, cells with
shorter circulating half-lives, such as polymorphonuclear leukocytes32
and inflammatory monocytes17,33, did not equilibrate in blood even
after 120 d (Fig. 2a and Supplementary Fig. 1). As reported
before28,29, the exchange of hematopoietic stem cells, defined as Lin–
Sca-1+c-Kit+ (ref. 34), in bone marrow was less than 5% (Fig. 2b).
DC exchange varied by subset and location. In the spleen, DC
exchange reached plateaus of 24.4% for CD8+DEC-205+ pDCs, 29.0%
for CD8–33D1+ pDCs and 16.7% for pDCs 30 d after surgery (Fig. 2c
and Supplementary Fig. 1). In lymph nodes at 30 d, exchange reached
plateaus of 21.7% for cDCs (CD11chiMHCint), 18.1% for pDCs and
19.7% for DCs derived from Langerhans cells (CD11c+MHChi; Fig. 2d
and Supplementary Fig. 1). Unexpectedly, those percentages of
exchange were stable for 120 d in both spleen and lymph nodes.
However, when we irradiated one of the parabiotic partners before the
initiation of parabiosis, nearly all its DCs were replaced by cells from
the unirradiated partner within 14 d (Fig. 2e). Our results are
consistent with the possibility that in the steady state, DCs are replaced
mainly by organ-derived precursors26, but that irradiation allows
replacement by blood-borne precursors35–37. Alternatively, DCs
might always originate in the blood, with incomplete equilibration
of DCs in the spleen and lymph nodes of parabionts because of
incomplete exchange of precursors. The latter is the case for
579
ARTICLES
c
48.0
47.6 47.3
40
Chimerism (%)
Spleen
48.5
47.7
37.1
30
29.0
30.1
24.4
20
16.7
10
1.9
0
b 50
hi
B cells
T cells
cDCs
LT-HSC
Bone marrow
d
MHC DCs
pDCs
Lymph node
49.0
48.1
40
30
21.7
19.7
20
18.1
10
2.6
0
0
20
40
60
80
100
120
0
20
40
60
80
100
120
Time after parabiosis (d)
e
CD45.1 cells (%)
© 2007 Nature Publishing Group http://www.nature.com/natureimmunology
Blood
85.5 90.2
100
71.4
80
*
60
40
B6 (CD45.2)
SJL (CD45.1)
18.2
20
0
Control
Figure 2 Leukocyte exchange in parabiotic mice. (a–d) Percent chimerism
in parabionts at various times after surgery. Numbers along lines indicate
average chimerism of each cell population after reaching plateau, for all
parabionts between 30 d and 120 d. Each time point represents the average
of three independent pairs of parabionts (error bars, s.d.). (a) Chimerism
of peripheral blood B cells, T cells, stationary monocytes (Stat mono),
polymorphonuclear leukocytes (PMN) and inflammatory monocytes (Infl
mono). (b) Chimerism of bone marrow hematopoietic stem cells (LT-HSC;
Lin–Sca-1+c-Kit+). (c) Chimerism of spleen B cells and T cells, CD8+ DCs,
CD8– DCs, pDCs and red pulp macrophages (RP Mph). (d) Chimerism of
lymph node B cells and T cells, cDCs, MHChi DCs and pDCs. (e) Percent
CD45.1+ spleen DCs in B6 (CD45.2) and SJL (CD45.1) partners 2 weeks
after irradiation of the B6 mouse in a B6-SJL pair of parabionts.
*, irradiated mouse. Numbers above bars indicate average value. Data
are representative of two experiments (error bars, s.d. for the average of
two experiments).
Irradiated
hematopoietic bone marrow precursors, which fail to exchange even
after prolonged parabiosis28,29.
DC precursors have a short half-life in circulation
To determine the distribution of DC precursors in blood of parabionts, we transferred blood from individual parabionts into irradiated F1 recipients and determined the phenotype of the DCs arising
in the recipient’s spleen. DC precursors were unevenly distributed in
the blood of parabionts, with only 17.1–35.6% originating from the
parabiotic partner (Fig. 3a). In contrast, transferred T cells originated
equally from both parabionts (Supplementary Fig. 3 online). We
Figure 3 Blood DC precursor clearance and exchange in parabiotic mice.
(a) Flow cytometry of DCs from CD45.1 CD45.2 F1 mice that had
received 6 Gy irradiation, followed by adoptive transfer of 3 106 blood
leukocytes from one parabiont (CD45.2) or the other parabiont (CD45.1)
at 40 d after initiation of parabiosis; spleen DCs were analyzed 8 d after
transfer of leukocytes. Data are CD45.1 versus CD45.2 expression on DCs
derived from transferred blood after gating on CD3–CD19–NK1.1–Ter119–
CD11chiI-A+ DCs and exclusion of F1 host DCs (CD45.1+CD45.2+).
Numbers adjacent to outlined areas indicate percent DCs derived from the
CD45.1 or CD45.2 parabiont; numbers in red circles indicate percent DC
precursor exchange between parabionts. (b) Flow cytometry of peripheral
blood leukocytes from primary donor CD45.2 mice that were injected into
CD45.1 recipients, whose blood was collected after 1 or 60 min, then
transferred into irradiated secondary F1 recipients. Data represent spleen
DCs derived from secondary blood transfer in F1 host after the gating
described in a. Numbers adjacent to outlined areas indicate relative percent
primary donor-derived DCs (CD45.2+; red arrows) and primary recipientderived DCs (CD45.1+). Data are one representative of two experiments
with two mice per group.
conclude that DC precursors in blood do not fully equilibrate between
parabionts. Furthermore, the amount of chimerism of DC precursors
in the blood (17.1–35.6%) was similar to the chimerism of DCs in
spleen and lymph nodes in the parabionts (Fig. 2c,d), suggesting that
DCs in peripheral lymphoid organs are in equilibrium with their
precursors in the blood.
Incomplete equilibration of DC precursors between parabionts
suggests that these cells may have a short residence time (under 2 h)
in the bloodstream28–31. To estimate the blood residence time of DC
precursors, we did serial adoptive transfer experiments similar to those
used to estimate the half-life of hematopoietic stem cells in circulation28. We injected blood leukocytes from CD45.2 donors into CD45.1
recipients, collected blood from the recipients after 1 min or 1 h and
transferred the blood to an irradiated secondary CD45.1 CD45.2 F1
recipient mouse. We determined the number of DCs arising from
CD45.2 donors by flow cytometry at 8 d after the secondary transfer.
We found that 65% of the DC precursors in the blood of CD45.2 donor
mice were lost from circulation of the CD45.1 recipient mice within
1 min and over 93% were cleared within 1 h (Fig. 3b and Supplementary Fig. 3). Thus, the uneven distribution of DCs in parabionts
reflects the short half-life of DC precursors in the circulation.
Limited DC cell division in spleen and lymph nodes
The number of donor and recipient DCs in spleen and lymph nodes of
parabionts reached equilibrium after 30 d of parabiosis, suggesting
that their half-life in lymphoid organs may be much longer than the
a
DCs
Blood donor
B6 (CD45.2)
parabiont
17.1
b
DCs
Time after
transfer
57.9
0 min
82.9
42.1
SJL (CD45.1)
parabiont
64.4
1 min
35.6
CD45.1
a 50
CD8+ DCs
pDCs
RP Mph
B cells
T cells
–
CD8 DCs
Infl. mono
CD45.1
Stat mono
PMN
B cells
T cells
84.5
14.5
CD45.2
60 min
97.1
2.7
CD45.2
580
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26.9
25.7
19.6
17.4
7.5
Chimerism (%)
0.3
0.7
0.7
–4.5
36.9
30
20
4.2
15.3
2.2
16.4
12.6
8.8
10
7.7
4.1
0.5
0.2
0.2
Spleen pDCs
30
0
4.8
0.7
2.6
0.7
Lymph node pDCs
40
21.9
37.0
30
20
20
12.6
10
4.6
2.3
5.5
3.1
4.4
1.0
2.5
0.8
0
1.8
hi
Lymph node CD11c DCs
19.0
17.2
10
0
5.5
4.9
0
40
31.3
25.0
SJL (CD45.1) parabionts
11.9
2.4
+
Spleen CD8 DCs
40
20
B6 (CD45.2) parabionts
14.5
10.7
3.3
1.1
30
26.8
10
10
0
Chimerism (%)
20
7
14
12.4
10
7.6
0
5.4
21
5.8
1.7
5.1
0.7
0
7
14
3.4
0.7
21
Time after separation (d)
1.5–2.9 d estimated by BrdU uptake22,23 (Fig. 1a). To directly
determine the rate of DC turnover and replacement from blood, we
separated parabiotic mice after DC equilibration (33 d) and then
measured the rate of loss of parabiont-derived DCs in lymphoid
organs. As there obviously could be no further exchange of bloodderived DCs after the parabiotic mice were separated, and as exchange
of hematopoietic precursors between bone marrow of the parabionts
was negligible (Fig. 2b), the survival of parabiont-derived DCs in
separated mice after parabiosis was a direct measure of residence time
in situ. All parabiont-derived cDCs in both spleen and lymph nodes
were lost from each of the separated parabionts with similar kinetics,
reaching background by 10–14 d (half-life, 5–7 d); in contrast,
parabiont-derived pDCs reached background amounts in only 3 d
(Fig. 4). Meanwhile, most parabiont-derived B cells and T cells were
retained 21 d after separation of the parabionts (data not shown).
Therefore, the pDC pools in the spleen and lymph nodes have more
rapid turnover than do the pools of cDCs. We conclude that despite
their capacity to divide in situ, replenishment of tissue DCs from the
circulation is required for DC homeostasis.
DC division prolongs antigen presentation
Individual DCs have a short half-life (1.5–2.9 d), as assessed by BrdU
incorporation22; nevertheless, DCs that are made to present a given
Figure 5 Antigen retention by dividing DCs in the spleen. (a) Flow cytometry
of pMHC expression, assessed by staining with monoclonal antibody Y-Ae
on the surface of CD45.1+ DCs transferred into B6 (CD45.2+) recipients
without I-Ea peptide injection (top) or immediately before (middle) or 20 h
after (bottom) I-Ea peptide injection; all cells were analyzed 12 h after
transfer. Numbers adjacent to outlined areas indicate percent Y-Ae+ cells.
(b) Flow cytometry of DCs from mice that received BrdU for 12 h beginning
24 h after injection of I-Ea peptide. Top, Y-Ae staining on gated CD11c+
DCs. Numbers adjacent to outlined areas indicate percent isotype staining
background (left) or Y-Ae+ cells (right). Bottom, BrdU incorporation by
CD11c+ DCs (All DC), CD11c+Y-Ae+ DCs (Y-Ae+) or CD11c+Y-Ae– DCs
(Y-Ae–). Numbers above bracketed lines indicate percent background
staining in control mice (left) or BrdU+ cells (right). Data are representative
of two experiments.
antigen in vivo38 can do so for as long as 14 d (refs. 7,39). To determine
whether the division of DCs in situ prolongs antigen presentation
because of passive transfer of long-lived pMHC complexes to daughter
cells during each division, we measured pMHC expression on DCs that
had divided after antigen was delivered to them by experimental
means. We injected peptide from the MHC molecule I-E a-chain
(I-Ea) intravenously into mice and detected MHC complexes expressing the specific peptide with monoclonal antibody Y-Ae (which is
specific for the peptide in complex with I-Ab; ref. 40).
As anticipated, unbound I-Ea peptide was rapidly cleared from
bloodstream, as shown by the finding that exogenous DCs (CD45.1+)
transferred immediately before peptide injection had considerable
Y-Ae staining (50.8%), whereas those transferred 20 h after peptide
injection did not (1.86%; Fig. 5a). Immediately after intravenous
injection of the I-Ea peptide, 15–30% of the DCs in spleen had surface
I-Ea–MHC expression, and a large fraction of those retained surface
I-Ea–MHC complexes 20 h later (Fig. 5a). To determine whether
I-Ea–MHC expression was retained on DCs after division, we
examined Y-Ae staining on DCs that incorporated BrdU at 24–36 h
after injection of I-Ea peptide (that is, after peptide clearance from
circulation, when 7.85% of all DCs had cell surface I-Ea–MHC
expression). We found that 13.5% of all DCs, 10.4% of Y-Ae+ DCs
and 13.9% Y-Ae– DCs were BrdU+ (Fig. 5b). Thus, daughters of
pMHC-expressing DCs can retain cell surface pMHC complexes that
may function to prolong antigen presentation in lymphoid organs.
DISCUSSION
DCs in peripheral lymphoid organs were initially considered to be
terminally differentiated, nondividing cells with a short half-life22,36,37.
That idea has been challenged by experiments with parabiotic mice
that have provided data suggesting that DCs in lymphoid organs are
replaced by locally derived precursors19,26,41. Furthermore, a fraction
of DCs in spleen and lymph nodes express the proliferation marker
Ki-67, rapidly incorporate BrdU and seem to be actively cycling26.
Although failure of spleen DCs to exchange between parabionts after
prolonged periods of shared circulation is consistent with replacement
by tissue-derived precursors19,26, an alternative not considered in
initial studies was that DC precursors in blood, like hematopoietic
a
CD45.1+ DCs I-Eα CD45.1 DCs
All DCs
0.068
b
0.006
0
–
12 h
31.9
0.25
7.85
Y-Ae
30
20
30
42.2
Figure 4 DC chimerism in parabiotic mice after surgical separation. Mean
spleen CD8– DC, CD8+ DC and pDC chimerism (left) and lymph node MHChi
DC, cDC and pDC chimerism (right) for CD45.2 parabionts (red) and
CD45.1 parabionts (blue) at various times after separation. CD45.2 and
CD45.1 congenic mice were surgically joined for 33 d before separation.
Numbers above data points represent average net DC chimerism of three to
six parabionts in independent pairs (error bars, s.d. of average).
Isotype
40
0
© 2007 Nature Publishing Group http://www.nature.com/natureimmunology
Lymph node MHChi DCs
0.092
50.8
Y-Ae
Chimerism (%)
Spleen CD8– DCs
50
12 h
12 h
CD11c
13.5%
Y-Ae+
0.16% +BrdU
12 h
0.10%
Y-Ae–
0.16%
13.9%
All DCs
15.3
0.029
1.86
32 h
12 h
–BrdU
BrdU
10.4%
BrdU
CD45.1
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ARTICLES
stem cells29, do not spend enough time in circulation to equilibrate
between parabionts28. Our experiments here have demonstrated that
DC precursors have a short half-life that leads to, among other things,
unequal distribution between parabionts. The degree of chimerism of
DC precursors in the blood of each parabiont (17.1–35.6%) corresponded to the degree of DC chimerism achieved in lymphoid organs
of each parabiont (16.7–29.0%). Thus, unequal DC exchange in the
spleen and lymph nodes of parabionts simply reflects the DC
precursor frequency in blood.
Our conclusion that DCs in peripheral lymphoid organs are in
continuous equilibrium with circulating blood precursors was further
supported by data demonstrating that separation of parabionts led to
the replacement of spleen and lymph node DCs in each separated
mouse after parabiosis by endogenous (bone marrow–derived) bloodborne cells within 10–14 d. We found that the individual cDC subsets
in the peripheral secondary lymphoid organs were replaced at similar
rates, whereas pDCs, which do not divide in peripheral lymphoid
organs, were replaced rapidly after the separation of parabiotic mice.
We conclude that the half-life of organ DC pools is between 5 d and
7 d, with replenishment from blood being required to maintain
steady-state DC populations.
In the steady state, DC homeostasis in lymphoid organs is maintained by means of a dynamic balance of three processes: constant
replenishment by DC precursors from blood; DC division; and DC
cell death after a limited number of divisions. For a given lymphoid
organ, by determining the number of DCs, the percent that are
dividing (about 5%)26, as well as the kinetics of BrdU labeling22 and
chimerism in individual parabionts, we have been able to solve an
algebraic equation that estimates of the rate of precursor input to be
4,236 precursor cells per hour. Thus, approximately 71 precursors
enter the spleen from blood each minute, and they then undergo a
limited number of divisions for 10–14 d. Whether DC division in situ
is restricted anatomically remains to be determined.
Priming, tolerance and maintenance of T cell memory all require
persistent antigen presentation by DCs42,43. Those immune consequences would not be feasible if DCs were nondividing cells with short
half-lives. However, a DC undergoing a limited number of divisions in
the periphery might serve as a ‘reservoir’ for pMHC complexes or
replicating pathogens that are passed down to successive daughter cells.
Our experiments have shown that pMHC complexes were retained on
the surface of DC daughters, which we speculate may extend the
duration of antigen presentation. Such ‘dilution’ of pMHC complexes
by DC division may regulate T cell fate, with more initial pMHC on
DCs stimulating the differentiation of T effector cells and less pMHC on
daughter DCs given rise to central memory or regulatory T cells44–46.
4 106 CD45.1 peripheral blood leukocytes, and the mixture was transferred
directly into irradiated F1 recipients. For peptide injection, 0.5 mg of I-Ea
peptide dissolved in 200 ml saline buffer was injected intravenously at a
different site from the injection of DCs. All mice were housed in specific
pathogen–free conditions and were treated in accordance with protocols
approved by the Institutional Animal Care and Use Committee of The Rockefeller University.
Parabiosis. Parabiosis and separation were done as reported28 with 5- to
6-week-old male B6 and B6 CD45.1+ mice that were matched for body weight.
Blood exchange was confirmed 8 d after parabiosis by injection of Evan’s blue
dye47. For each pair of joined mice, average chimerism was calculated as
follows: (percent CD45.1+ cells in CD45.2 mouse + percent CD45.2+ cells in
CD45.1 mouse) / 2. For each separated parabiont, net DC chimerism was
calculated as follows: percent donor-derived DCs – percent donor-derived
hematopoietic stem cells.
Reagents. The following reagents were from BD Biosciences or eBioscience:
anti-CD16-CD32 (2.4G2); biotin-conjugated anti-I-Ab (AF6-120.1) and antiCD45.2 (104); fluorescein isothiocyanate–conjugated anti-CD45.2 (104),
anti-BrdU(PRB-1), anti-4/80 (BM8) and anti-Ki-67 (B56); phycoerythrinconjugated anti-B220 (RA3-6B2), anti-CD40 (3/23), anti-CD62L (MEL-14),
anti-CD80 (16-10A1), anti-CD86 (GL1), anti-Ki-67 (B56), anti-H2-Kb
(AF6-88.5), anti-I-Ab (AF6-120.1), anti-CD1d (1B1) and anti-Sca-1 (D7);
phycoerythrin-indodicarbocyanine–conjugated anti-CD45.1 (A20); phycoerythrin–carbocyanine 5.5–conjugated anti-Gr-1(RB6-8C5); phycoerythrinindotricarbocyanine–conjugated anti-CD3 (145-2C11), anti-CD19 (1D3),
anti-NK1.1 (PK136) and anti-Ter119(TER-119); Pacific blue–conjugated
anti-Gr-1 (RB6-8C5); allophycocyanin-conjugated anti-CD11b (M1/70), antiCD11c (N418) and anti-c-Kit (2B8); Alexa Fluor 700–conjugated anti-CD3
(17A2) and anti-CD8a (53-6.7); Alexa Fluor 750–conjugated anti-CD11b (M1/
70); allophycocyanin-indotricarbocyanine–conjugated anti-CD19 (1D3), antiB220 (RB6-8C5) and anti-Ter119 (TER-119); and Cytoperm/Cytofix solution
and Perm/Wash buffer. DAPI and Pacific orange–streptavidin were from
Molecular Probes. Phycorythrin-conjugated anti–mouse pDC antigen (antiPDCA) and anti-CD11c microbeads were from Miltenyi Biotec. Monoclonal
antibody Y-Ae was from Chemicon and was biotinylated with a biotinylation
kit (Sigma). Other reagents included PBS and FBS (Gibco-BRL), CSFE
(carboxyfluorescein succinimidyl ester; Molecular Probes), ACK lysing buffer
(BioSource) and BrdU (Sigma). I-Ea peptide was synthesized at the Rockefeller
University protein facility.
METHODS
BrdU labeling and cell cycle analysis. For BrdU labeling experiments, after the
initial intraperitoneal injection of BrdU (2 mg per mouse), mice were
maintained on water containing 0.8 mg/ml of BrdU. Splenocytes were stained
according to the instruction manual from the BD BrdU staining kit with DNase
treatment. For cell cycle analysis, samples were enriched for spleen DCs with
CD11c microbeads and Miltenyi magnetic columns. After extracelluar staining
with anti-CD11c and anti-CD8, permeablization and fixation, and intracellular
staining with phycoerythrin-conjugated anti-Ki-67, cells were incubated for
30 min at 4 1C with DAPI immediately before analysis by flow cytometry with
the LSR II System (BD Biosciences) with gating on single nuclei.
Mice, renal capsule grafting, blood transfer and peptide injection. C56BL/6J
(B6), C57BL/6 Pep3b CD45.1+ (SJL) and CD45.1+CD45.2+ F1 mice 5–10 weeks
of age were from Jackson Lab or were bred at Rockefeller University. Renal
capsule grafting of spleen pieces was done according to a procedure published
online (http://mammary.nih.gov/tools/mousework/cunha001/index.html). For
peripheral blood transfer, 2 106 to 10 106 peripheral blood leukocytes were
injected intravenously into 8- to 10-week-old recipient mice that had received
sublethal irradiation (6 Gy). For analysis of blood clearance of DC precursors,
10 106 peripheral blood leukocytes from primary donor CD45.2 mice were
injected intravenously into CD45.1 recipients whose blood was collected after
1 or 60 min. The total peripheral blood leukocyte count for each CD45.1
mouse was approximately 20 106. Blood collected from the primary recipient
was injected intravenously into irradiated secondary CD45.1 CD45.2 F1
recipients, and spleen DC phenotype was analyzed after 9 d. As a control for
no clearance, 2 106 CD45.2 peripheral blood leukocytes were mixed with
Cell preparation. Spleens, peripheral (axillary, brachial and inguinal) lymph
nodes and spleen grafts were cut into small fragments and were digested for
30 min at 37 1C with mixing in 10 ml Hank’s buffer containing collagenase
(375 mU/ml; type II; Roche). For disruption of DC–T cell complexes, 200 ml of
0.5 M EDTA, pH 8.0, was added and mixing continued for 5 min. Undigested
fibrous material was removed by filtration through nylon mesh. All subsequent
steps were at 4 1C with 5% (vol/vol) FBS in PBS. In some experiments, samples
were enriched for DCs from spleen and lymph nodes by incubation with
CD11c microbeads followed by passage through a Miltenyi Biotec column.
Peripheral blood was collected into a solution containing heparin by heart
puncture. Red blood cells were lysed by the addition of 10 volume of ACK
lysis buffer and incubation for 10 min at 25 1C followed by two washes with
flow cytometry buffer. For injection in vivo, peripheral blood leukocytes were
resuspended in 100–200 ml PBS.
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© 2007 Nature Publishing Group http://www.nature.com/natureimmunology
ARTICLES
Flow cytometry. Cells were stained at 4 1C in PBS with 5% (vol/vol) FBS.
Intracellular staining with anti-Ki-67 and BrdU was done with Cytofix/
Cytoperm solution according to the manufacturer’s protocol (BD Biosciences).
An LSR II (Becton Dickinson) was used for multiparameter flow cytometry of
stained cell suspensions, followed by analysis with FlowJo software (TreeStar) or
Diva software (Becton Dickinson). Cells were gated as follows: B cells, CD11b–
NK1.1–Ter119–CD19+CD3–; T cells, CD11b–NK1.1–Ter119–CD19–CD3+; polymorphonuclear leukocytes, CD3–CD19–NK1.1–Ter119–SSChiCD11bhiGr-1hi;
stationary monocytes, CD3–CD19–NK1.1–Ter119–SSCloCD11bhiCD62L–Gr-1–;
inflammatory monocytes, CD3–CD19–NK1.1–Ter119–SSCloCD11bhiCD62L+
Gr-1int (refs. 17,48); bone marrow hematopoietic stem cells, Lin– (CD19–CD3–
CD4–CD8–NK1.1–B220–Ter119–CD11b–CD11c–)Sca-1+c-Kit+ (refs. 28,34);
splenic CD8+ DCs, CD3–CD19–NK1.1–Ter119–CD11chiI-A+CD8+; CD8– DCs,
CD3–CD19–NK1.1–Ter119–CD11chiI-A+CD8–; red pulp macrophages, CD3–
CD19–NK1.1–Ter119–CD11bloF4/80hi (ref. 48); lymph node cDCs, CD3–
CD19–NK1.1–Ter119–CD11chiI-A+; MHC-high DCs, CD3–CD19–NK1.1–
Ter119–CD11cintI-Ahi; spleen and lymph node pDCs, CD3–CD19–NK1.1–
Ter119–CD11cintI-A–PDCA+ (ref. 49).
Calculation of DC precursor input. These calculations are described in the
Supplementary Methods online.
Note: Supplementary information is available on the Nature Immunology website.
ACKNOWLEDGMENTS
We thank R. Steinman, K. Tarbell and E. Besmer for reading the manuscript.
Supported by the National Institutes of Health (M.N. and J.H.), the Verto
Institute (J.H.) and the Howard Hughes Medical Institute (M.N.).
COMPETING INTERESTS STATEMENT
The authors declare no competing financial interests.
Published online at http://www.nature.com/natureimmunology/
Reprints and permissions information is available online at http://npg.nature.com/
reprintsandpermissions
1. Liu, Y.J. Dendritic cell subsets and lineages, and their functions in innate and adaptive
immunity. Cell 106, 259–262 (2001).
2. Shortman, K. & Liu, Y.J. Mouse and human dendritic cell subtypes. Nat. Rev. Immunol.
2, 151–161 (2002).
3. Steinman, R.M. et al. Dendritic cell function in vivo during the steady state: a role in
peripheral tolerance. Ann. NY Acad. Sci. 987, 15–25 (2003).
4. Crowley, M., Inaba, K., Witmer-Pack, M. & Steinman, R.M. The cell surface of mouse
dendritic cells: FACS analyses of dendritic cells from different tissues including
thymus. Cell. Immunol. 118, 108–125 (1989).
5. Witmer, M.D. & Steinman, R.M. The anatomy of peripheral lymphoid organs with
emphasis on accessory cells: light-microscopic immunocytochemical studies of mouse
spleen, lymph node, and Peyer’s patch. Am. J. Anat. 170, 465–481 (1984).
6. Vremec, D. et al. The surface phenotype of dendritic cells purified from mouse thymus
and spleen: investigation of the CD8 expression by a subpopulation of dendritic cells.
J. Exp. Med. 176, 47–58 (1992).
7. Dudziak, D. et al. Differential antigen processing by dendritic cell subsets in vivo.
Science 315, 107–111 (2007).
8. Manz, M.G. et al. Dendritic cell development from common myeloid progenitors. Ann.
NY Acad. Sci. 938, 167–173 (2001).
9. Traver, D. et al. Development of CD8a-positive dendritic cells from a common myeloid
progenitor. Science 290, 2152–2154 (2000).
10. Fogg, D.K. et al. A clonogenic bone marrow progenitor specific for macrophages and
dendritic cells. Science 311, 83–87 (2006).
11. Harman, B.C., Miller, J.P., Nikbakht, N., Gerstein, R. & Allman, D. Mouse plasmacytoid
dendritic cells derive exclusively from estrogen-resistant myeloid progenitors. Blood
108, 878–885 (2006).
12. Colonna, M., Trinchieri, G. & Liu, Y.J. Plasmacytoid dendritic cells in immunity. Nat.
Immunol. 5, 1219–1226 (2004).
13. Sallusto, F. & Lanzavecchia, A. Efficient presentation of soluble antigen by cultured
human dendritic cells is maintained by granulocyte/macrophage colony-stimulating
factor plus interleukin 4 and downregulated by tumor necrosis factor alpha. J. Exp.
Med. 179, 1109–1118 (1994).
14. Bender, A., Sapp, M., Schuler, G., Steinman, R.M. & Bhardwaj, N. Improved methods
for the generation of dendritic cells from nonproliferating progenitors in human blood.
J. Immunol. Methods 196, 121–135 (1996).
15. Randolph, G.J., Beaulieu, S., Lebecque, S., Steinman, R.M. & Muller, W.A. Differentiation of monocytes into dendritic cells in a model of transendothelial trafficking.
Science 282, 480–483 (1998).
NATURE IMMUNOLOGY
VOLUME 8
NUMBER 6
JUNE 2007
16. Randolph, G.J., Inaba, K., Robbiani, D.F., Steinman, R.M. & Muller, W.A. Differentiation of phagocytic monocytes into lymph node dendritic cells in vivo. Immunity 11,
753–761 (1999).
17. Geissmann, F., Jung, S. & Littman, D.R. Blood monocytes consist of two principal
subsets with distinct migratory properties. Immunity 19, 71–82 (2003).
18. Ginhoux, F. et al. Langerhans cells arise from monocytes in vivo. Nat. Immunol. 7,
265–273 (2006).
19. Naik, S.H. et al. Intrasplenic steady-state dendritic cell precursors that are distinct
from monocytes. Nat. Immunol. 7, 663–671 (2006).
20. Merad, M. et al. Langerhans cells renew in the skin throughout life under steady-state
conditions. Nat. Immunol. 3, 1135–1141 (2002).
21. Varol, C. et al. Monocytes give rise to mucosal, but not splenic, conventional dendritic
cells. J. Exp. Med. 204, 171–180 (2006).
22. Kamath, A.T. et al. The development, maturation, and turnover rate of mouse spleen
dendritic cell populations. J. Immunol. 165, 6762–6770 (2000).
23. Kamath, A.T., Henri, S., Battye, F., Tough, D.F. & Shortman, K. Developmental kinetics
and lifespan of dendritic cells in mouse lymphoid organs. Blood 100, 1734–1741
(2002).
24. Ruedl, C., Koebel, P., Bachmann, M., Hess, M. & Karjalainen, K. Anatomical origin of
dendritic cells determines their life span in peripheral lymph nodes. J. Immunol. 165,
4910–4916 (2000).
25. Zhang, M. et al. Splenic stroma drives mature dendritic cells to differentiate into
regulatory dendritic cells. Nat. Immunol. 5, 1124–1133 (2004).
26. Kabashima, K. et al. Intrinsic lymphotoxin-b receptor requirement for homeostasis of
lymphoid tissue dendritic cells. Immunity 22, 439–450 (2005).
27. Bogunovic, M. et al. Identification of a radio-resistant and cycling dermal dendritic cell
population in mice and men. J. Exp. Med. 203, 2627–2638 (2006).
28. Wright, D.E., Wagers, A.J., Gulati, A.P., Johnson, F.L. & Weissman, I.L. Physiological
migration of hematopoietic stem and progenitor cells. Science 294, 1933–1936
(2001).
29. Abkowitz, J.L., Robinson, A.E., Kale, S., Long, M.W. & Chen, J. Mobilization of
hematopoietic stem cells during homeostasis and after cytokine exposure. Blood
102, 1249–1253 (2003).
30. Harris, R.B., Zhou, J., Weigle, D.S. & Kuijper, J.L. Recombinant leptin exchanges
between parabiosed mice but does not reach equilibrium. Am. J. Physiol. 272,
R1800–R1808 (1997).
31. Nishizawa, Y. & Bray, G.A. Evidence for a circulating ergostatic factor: studies on
parabiotic rats. Am. J. Physiol. 239, R344–R351 (1980).
32. Lagasse, E. & Weissman, I.L. Bcl-2 inhibits apoptosis of neutrophils but not their
engulfment by macrophages. J. Exp. Med. 179, 1047–1052 (1994).
33. van Furth, R. & Cohn, Z.A. The origin and kinetics of mononuclear phagocytes. J. Exp.
Med. 128, 415–435 (1968).
34. Morrison, S.J. & Weissman, I.L. The long-term repopulating subset of hematopoietic
stem cells is deterministic and isolatable by phenotype. Immunity 1, 661–673
(1994).
35. Steinman, R.M., Lustig, D.S. & Cohn, Z.A. Identification of a novel cell type in
peripheral lymphoid organs of mice. 3. Functional properties in vivo. J. Exp. Med.
139, 1431–1445 (1974).
36. Steinman, R.M. & Nussenzweig, M.C. Dendritic cells: features and functions. Immunol. Rev. 53, 127–147 (1980).
37. Pugh, C.W., MacPherson, G.G. & Steer, H.W. Characterization of nonlymphoid cells
derived from rat peripheral lymph. J. Exp. Med. 157, 1758–1779 (1983).
38. Hawiger, D. et al. Dendritic cells induce peripheral T cell unresponsiveness under
steady state conditions in vivo. J. Exp. Med. 194, 769–779 (2001).
39. Bonifaz, L.C. et al. In vivo targeting of antigens to maturing dendritic cells via the
DEC-205 receptor improves T cell vaccination. J. Exp. Med. 199, 815–824
(2004).
40. Rudensky, A., Rath, S., Preston-Hurlburt, P., Murphy, D.B. & Janeway, C.A., Jr. On the
complexity of self. Nature 353, 660–662 (1991).
41. Diao, J. et al. In situ replication of immediate dendritic cell (DC) precursors contributes
to conventional DC homeostasis in lymphoid tissue. J. Immunol. 176, 7196–7206
(2006).
42. Redmond, W.L., Hernandez, J. & Sherman, L.A. Deletion of naive CD8 T cells requires
persistent antigen and is not programmed by an initial signal from the tolerogenic APC.
J. Immunol. 171, 6349–6354 (2003).
43. Zammit, D.J., Cauley, L.S., Pham, Q.M. & Lefrancois, L. Dendritic cells maximize the
memory CD8 T cell response to infection. Immunity 22, 561–570 (2005).
44. Bullock, T.N., Mullins, D.W. & Engelhard, V.H. Antigen density presented by dendritic
cells in vivo differentially affects the number and avidity of primary, memory, and recall
CD8+ T cells. J. Immunol. 170, 1822–1829 (2003).
45. Catron, D.M., Rusch, L.K., Hataye, J., Itano, A.A. & Jenkins, M.K. CD4+ T cells that
enter the draining lymph nodes after antigen injection participate in the primary
response and become central-memory cells. J. Exp. Med. 203, 1045–1054 (2006).
46. Kretschmer, K. et al. Inducing and expanding regulatory T cell populations by foreign
antigen. Nat. Immunol. 6, 1219–1227 (2005).
47. Ansel, K.M., Harris, R.B. & Cyster, J.G. CXCL13 is required for B1 cell homing, natural
antibody production, and body cavity immunity. Immunity 16, 67–76 (2002).
48. Gordon, S. & Taylor, P.R. Monocyte and macrophage heterogeneity. Nat. Rev. Immunol.
5, 953–964 (2005).
49. Blasius, A. et al. A cell-surface molecule selectively expressed on murine natural
interferon-producing cells that blocks secretion of interferon-a. Blood 103, 4201–
4206 (2004).
583