Minimal Differentiation of Classical Monocytes
as They Survey Steady-State Tissues
and Transport Antigen to Lymph Nodes
Claudia Jakubzick,1,2,3Emmanuel L. Gautier,4Sophie L. Gibbings,1Dorothy K. Sojka,5Andreas Schlitzer,7
Theodore E. Johnson,2Stoyan Ivanov,4Qiaonan Duan,8Shashi Bala,4Tracy Condon,2Nico van Rooijen,9
John R. Grainger,10Yasmine Belkaid,10Avi Ma’ayan,8David W.H. Riches,1,2Wayne M. Yokoyama,5,6Florent Ginhoux,7
Peter M. Henson,1,2and Gwendalyn J. Randolph3,4,*
1Department of Pediatrics at National Jewish Health, Denver, CO 80206, USA
2Integrated Department of Immunology, University of Colorado, Denver, CO 80206, USA
3Department of Gene and Cell Medicine, Mount Sinai School of Medicine, New York, NY 10029, USA
4Department of Pathology
5Department of Medicine
6Howard Hughes Medical Institute
Washington University Medical School, St. Louis, MO 63110, USA
7SingaporeImmunology Network, Singapore(SIgN), Agency for Science, Technology and Research(A*STAR), 138648 Singapore, Singapore
8Department of Pharmacology and Systems Therapeutics, Mount Sinai School of Medicine, New York, NY 10029, USA
9Department of Molecular Cell Biology, Free University Medical Center, Amsterdam 1007, the Netherlands
10Mucosal Immunology Section, Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of
Health, Bethesda, MD 20892, USA
It is thought that monocytes rapidly differentiate to
macrophages or dendritic cells (DCs) upon leaving
blood. Here we have shown that Ly-6C+monocytes
constitutively trafficked into skin, lung, and lymph
nodes (LNs). Entry was unaffected in gnotobiotic
mice. Monocytes in resting lung and LN had similar
gene expression profiles to blood monocytes but
elevated transcripts of a limited number of genes
including cyclo-oxygenase-2 (COX-2) and major his-
tocompatibility complex class II (MHCII), induced by
monocyte interaction with endothelium. Parabiosis,
bromodoxyuridine (BrdU) pulse-chase analysis, and
intranasal instillation of tracers indicated that instead
of contributing to resident macrophages in the lung,
recruited endogenous monocytes acquired antigen
for carriage to draining LNs, a function redundant
with DCs though differentiation to DCs did not occur.
Thus, monocytes can enter steady-state nonlym-
phoid organs and recirculate to LNs without differen-
tiation to macrophages or DCs, revising a long-held
view that monocytes become tissue-resident macro-
phages by default.
In the 1960s, van Furth and Cohn used labeling with [3H]
thymidine to define kinetics of monocyte transit in the blood-
stream. Monocytes spent at most a few days in blood and
then mobilized to various tissues, including the inflamed perito-
neum (van Furth and Cohn, 1968). In their view, the data indi-
cated that monocytes continuously replenished tissue-resident
macrophages. The concept was supported by the relative ease
with which monocytes became macrophages in culture (van
Furth and Cohn, 1968), and more recent data specifically point
to the Ly-6C+subset of monocytes (Wynn et al., 2013) as
precursors for macrophages in inflammatory settings. Begin-
ning in the mid-1990s, arguments developed that the fate of
monocytes could be diverted from becoming macrophages
and redirected to that of dendritic cells (DCs), if appropriate
signals were encountered (Iijima et al., 2011; Leo ´n et al., 2007;
Nakano et al., 2009; Sallusto and Lanzavecchia, 1994;
Serbina et al., 2003; Wakim et al., 2008). This differentiation,
like the evidence for monocyte conversion to macrophages
in vivo, stemmed almost entirely from scenarios involving
After the work of Van Furth and Cohn, an alternative possibility
that macrophages renewed themselves by local proliferation
rather than dependence upon monocytes was presented (Cog-
gle and Tarling, 1984; Sawyer et al., 1982). This viewpoint was
difficult to reconcile with the concept of monocytes as precur-
sors of macrophages and it failed to gain major traction without
stronger evidence than existed at the time. Recently, definitive
studies have been carried out on the origin of tissue-resident
macrophages, starting with brain macrophages (Ginhoux et al.,
2010) and then extending to many other organs (Hashimoto
et al., 2013; Schulz et al., 2012; Yona et al., 2013). The data
clearly indicate that in most organs, resident macrophages are
derived embryonically and maintain themselves in adults by
self-renewal. An exception is observed in the intestine where
macrophages are continuously repopulated by circulating
monocytes (Zigmond and Jung, 2013).
These recent findings, therefore, alter a key aspect of the
model that the field has held on the life cycle of monocytes
Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc. 599
since the 1960s and raise new questions about the biology
of monocytes and whether they indeed extravasate constitu-
tively (Wynn et al., 2013). Here, we asked whether monocytes
extravasate into steady-state tissues. Upon finding such mono-
cytes, we addressed their differentiation status. Our results sup-
port a revised paradigm wherein extravasated monocytes in
noninflamed tissues retain much of their monocytic character,
rather than differentiating to macrophages or DCs. Even so,
they survey tissue for antigens for transport to draining lymph
Classical Monocytes Are Present in Resting
Nonlymphoid Tissues and Lymph Nodes
CD64 (FcgR1) selectively recognizes macrophages, and Mer
tyrosine kinase (MerTK) is an additional macrophage-selective
marker (Gautier et al., 2012; Tamoutounour et al., 2012). Not
previously tested in skin, MerTK identified dermal macrophages
as well, in contrast to isotype-matched control Ab (Figure S1A
available online). Costaining for MerTK and CD64 marked mac-
rophages (Figure 1A) but excluded DCs and monocytes.
Although monocytes express CD64 (Ingersoll et al., 2010),
they do not express MerTK (Figure 1B). Skin DCs were identified
as remaining MerTK?CD64?cells expressing high CD11c and
Presence of Monocytes
(A and C) For skin (A) and lung (C), doublet-cell-
excluded, live cells were plotted as CD11c versus
MerTK versus CD64 defined tissue macrophages
as MerTK+CD64+. Gated MerTK+CD64+macro-
phages were CD11c+and CD11b+(top panels).
Continuing with the remaining MerTK?CD64?
gate, DCs were identified as MerTK?CD64?
CD11c+MHCII+(second row panels), with both
CD11bhiand CD11blosubsets. Events captured in
neither the macrophage or DC gate were then
plotted to depict CD64 versus CD11b or CD11c
versus CD11b, allowing us to identify monocytes
and granulocytes (third row panels). Tissue
monocytes were low-SSC F480+Ly-6C+MHCII+
(fourth and fifth row panels).
(B) Blood monocytes were stained with anti-
MerTK mAb but lacked reactivity. Black line, anti-
Mertk mAb; gray profile, isotype control mAb.
(D) SSC overlay of tissue and LN monocytes,
DCs, and macrophages from gates shown in (A)
Data are representative of R3 experiments. Fig-
ure S1 accompanies this figure.
1. Analysisof Tissuesfor the
major histocompatibility complex class
II (MHCII) molecules (Figure 1A). Then,
by gating on cells that were neither
macrophages nor DCs, monocytes were
identified in skin. Gating on remaining
CD11b+cells revealed two populations
of F4/80locells, with low and higher side
scatter (SSC), respectively, and a high
SSC population lacking F4/80 (Figure 1A). The latter were
neutrophils, whereas F4/80lohigh-SSC cells were eosino-
phils (Gautier et al., 2012). The low-SSC F4/80lo
were monocytes. Alternatively, gating directly on CD64+
cells aftergatingout MerTK+CD64+
MerTK?CD64?CD11chiMHCII+DCs was sufficient to identify
tissue monocytes (Figure 1A). All identified monocytes ex-
pressed Ly-6C and most expressed MHCII (Figure 1A). Use of
the same strategy in digested lymph node (LN) (Figure S1B)
and lung (Figure 1C) permitted identification of monocytes in
these locations. In all tissues, monocytes accounted for all
low-SSC, CD11bhicells with low to intermediate CD11c (Figures
1A, 1C, and 1D). Macrophages and DCs were larger and had
more granular cytoplasm resulting in greater SSC than mono-
cytes, consistent with properties expected of bona fide mono-
cytes (Figure 1D). The frequency of extracted monocytes was
roughly equal to macrophages in nonlymphoid tissues (Fig-
ure S2), whereas we extracted fewer macrophages from LNs
than monocytes. These ratios may underestimate macro-
phages, notoriously challenging to extract. LN monocytes
were one-tenth the frequency of total LN DCs (Figure S2) but
were similar in frequency to migratory, lymph-derived DCs in
resting LNs (Jakubzick et al., 2008a). Taken together, we
conclude that classical monocytes constitutively populate
noninflamed tissues, where they express MHCII.
600 Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc.
Extravascular Monocytes Resemble MHCII+Blood
Classical monocytes are thought to lack MHCII (Geissmann
et al., 2003; Ingersoll et al., 2010). Detection of MHCII+mono-
cytes in tissues and LNs prompted us to re-examine blood for
MHCII+monocytes. A minority of monocytes were MHCII+.
These were high or intermediate for Ly-6C (Figures 2A and 2B).
Because MHCII+monocytes showed a spectrum of Ly-6C
to CD43 (leukosialin) expression rather than Ly-6C (Figure 2A).
CD43 differentially defines the major subsets of monocytes in
all species examined to date (Ziegler-Heitbrock, 2007). Classical
CD43lomonocytes were divisible into three subsets (Ly-6C+
quencies in blood among total monocytes of 36.1%, 8.9%, and
1.0%, respectively (Figures 2A and 2B). CD43himonocytes were
divided into two subsets (Ly-6C?MHCII?and Ly-6C?MHCII+
[gates 4 and 5]) at frequencies of 41.9% and 8.1% (Figures 2A
and 2B) among total monocytes.
In Cx3cr1gfpmice, where blood monocyte subsets express
distinct intensities of GFP (Geissmann et al., 2003), gates 1–3
of blood CD43lomonocytes displayed a graded increase in
GFP intensity (Figure 2C) that correlated positively with MHCII
but inversely to Ly-6C. In CD43himonocytes (gates 4 and 5),
GFP intensity was characteristically high (Geissmann et al.,
2003) regardless of MHCII (Figure 2C). Monocytes in tissue
and LNs had overlapping GFP intensity with monocytes in blood
that were Ly-6C+MHCII+(gate 2) (Figure 2D). Furthermore, in
Lyz2-cre 3 RosaEGFPflox/+reporter mice, the frequency of
GFP+cells differs between monocyte subsets (Jakubzick et al.,
2008a). Analysis of the blood for MHCII+monocytes revealed
that all the CD43lomonocytes (gates 1–3), regardless of MHCII
expression, had a similar frequency of GFP+cells, and that all
CD43himonocyte subpopulations (subsets 4 and 5) shared the
same GFP+frequency (Figure 2E) but were distinct in frequency
from CD43lomonocytes (Figure 2E; Jakubzick et al., 2008a).
In tissue and LNs of Lyz2-cre 3 RosaEGFPflox/+mice, Ly-6C+
monocytes displayed similar EGFP frequency as
observed for blood Ly-6C+MHCII+monocytes (gate 2) (Fig-
ure 2F), indicating that within tissues these monocytes still
resembled blood monocytes and had not induced further
expression of lysozyme as observed in differentiated macro-
phages (Jakubzick et al., 2008a). Tissue monocytes probably
did not derive from Ly-6C?MHCII?monocytes (gate 4) because
tissue monocytes express a lower EGFP frequency than do
Ly-6C?MHCII?blood monocytes (Jakubzick et al., 2008a),
as shown by the fact that once EGFP is turned on within a
cell, it becomes irreversibly marked and cannot revert to an
Next, we compared gene expression profiles of monocytes
with macrophages and DC populations. Upregulation of MHCII
on Ly-6C+blood monocytes was accompanied by limited
change in gene expression. Only four mRNA transcripts
were R3-fold more abundant (Figure S3A; Table S1 lists
mRNA transcripts R2-fold elevated), including Tmem176b in
the CD20 family, Mrc1 encoding CD206 (mannose receptor),
the receptor tyrosine kinase Axl closely related to Mertk, and
MHCII (H2-Aa). By flow cytometry, CD206 was selectively
detected on MHCII+Ly-6Chimonocytes (Figure S3B). No tran-
scripts were significantly downregulated.
Our goal was to ask whether tissue monocytes were similar
to blood monocytes, macrophages, or DCs, because Ly-6C+
MHCII+cells in inflamed tissues have been called tumor necro-
sis factor and inducible nitric oxide-producing (Tip) DCs or
Figure 2. Analysis of Blood and Tissue
Monocytes in Cx3cr1gfpMice and Lyz2-
cre 3 Rosa26 EGFP Reporter Mice
(A)Thegating strategy leading togenerationoffive
gates for monocyte subsets is shown in dot plots
(left) and a diagram (right).
(B) The frequency of blood monocyte subsets in
gates 1–5 from (A) are plotted on a log scale.
(C) GFP intensity of blood monocyte subsets,
gates 1–5, from Cx3cr1gfp/gfpmice.
(D) Overlay of GFP intensity in monocytes from
blood, lung, skin, and LNs in Cx3cr1gfp/gfpmice.
Ly-6C+MHCII?(gate 1) and Ly-6C?MHCII?(gate
4) blood monocytes were used to illustrate GFP
intensity of these subsets.
(E) The frequency of GFP+cells among blood
monocyte subsets, gates 1–5, from Lyz2-cre 3
RosaGFPfloxreporter mice. Each dot represents
one mouse from six experiments.
(F) EGFP+frequency within indicated populations
divided by the frequency of EGFP+Ly-6C+MHCII+
(gate 2) blood monocytes in the same mouse.
Blood Ly-6C?MHCII?monocytes (gate 4) and
pDCs were used as controls. Abbreviations are as
follows: LLN, lung-draining LN; SLN, skin-draining
LN; MLN, mesenteric LN.
All data are derived from three to six experiments.
Figure S2 accompanies this figure.
Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc. 601
available from Immgen (http://www.immgen.org) revealed that
LN monocytes clustered most closely with blood monocytes
migratory DCs in skin-draining LNs (SLNs) (Figure 3A), and LN-
resident CD4+and CD8+DCs (Figure 3A). Lymph-migratory
DCs from distinct sources clustered (Miller et al., 2012), but LN
monocytes did not cluster with migratory DCs or LN-resident
cytes rather than as DCs.
We extended analysis of tissue monocytes to include those in
lung. Because Affymetrix had updated their platform by the time
we made this second round of isolations, we did not combine
the new data. Instead, we reprofiled LN monocytes (from SLNs
or mesenteric LNs [MLNs]), Ly-6C+blood monocytes (gate 1),
and the two distinct CD11b?and CD11b+lung macrophages
(Figure 1C). Lung monocytes clustered with LNs and blood
(Figure 3B). A few dozen mRNA transcripts were elevated R3-
fold in tissue monocytes relative to blood. Many of these
transcripts were upregulated in common between lung and LN
monocytes, pointing to a common signature induced upon
monocyte entry into tissues (Figure 3C, Table S2). This signature
included Ptgs2 encoding cyclo-oxygenase 2 (COX-2), Il1b
encoding interleukin 1-b, Tnfaip3 encoding A20, and Itgax
encoding CD11c (Figure 3C). CD11c was not expressed on
on tissue monocytes, albeit lower than on DCs (Figure 1). Immu-
nostaining analysis revealed COX-2 in Ly-6C+mononuclear cells
in lung parenchyma (Figure 3D), indicating expression of COX-2
in lung monocytes, along with its expression on neighboring
DCs that were detected as cells expressing more MHCII in the
lung than tissue monocytes (Figure 3D). Ptgs2 mRNA is ex-
pressed by CD11b+CD24+DCs in the Immgen database, fitting
with these results. Some transcripts were upregulated only in
(Eberlein et al., 2010) whose transcripts were upregulated only in
LN and not in lung monocytes, including CCL5 and CXCL10,
stained LN monocytes more distinctly than lung monocytes
(Figure 3E). An interferon (IFN) signature was observed in LN
but not lung monocytes (CXCL9, CXCL10, etc.) (Table S2).
Neutralizing IFN-g did not reduce MHCII on blood monocytes
(Figure S3C) and mice lacking IFN-g and type I IFN receptor
also retained MHCII+blood monocytes (data not shown).
Thus, tissue monocytes derived from steady-state organs
closely resemble blood monocytes, rather than macrophages
or DCs. The persistence of monocytes as bona fide monocytes
in tissues probably applies only to steady-state tissues because
monocytes recruited to sites of inflammation differentiated
into macrophages and clustered with macrophages resident in
that organ. That is, thioglycollate-elicited monocyte-derived
peritoneal macrophages (labeled as PC Mo-MF with MHCII
[II+] or without MHCII [II?]; Figure 3A) clustered with resident
Comparing Tissue and Blood Monocytes
with Macrophages and DCs
Principle component analysis (PCA) is a mathe-
matical procedure that depicts uncorrelated vari-
ables on each axis. Percentage shown on each
axis indicate the percent of variability explained,
with the first principle component (PC1) defined
as that which explains the most variability. The
placement of data in proximity indicates a closer
relationship between the data sets.
(A) PCA of LN-resident DCs CD8+and CD4+,
migratory LN DCs, Langerhans cells (LCs), mac-
rophages (MF), and monocytes (Mo) from LN and
blood (BL) (population gates 1–5 from Figure 2;
gate numbers next to Mo symbol). Abbreviations
are as follows: PC, peritoneal cavity; CNS, central
nervous system; Sp, spleen; Lu, lung. Each pop-
ulation is depicted with a different color.
(B) PCA of lung (Lu) alveolar macrophages
(CD11b?) and interstitial macrophages (CD11b+)
and monocytes from blood, lung, MLN, and SLN.
Each population is depicted with a different color.
(C) Heat map depicts genes that are elevated R3-
fold in blood, lung, and SLN monocytes.
(D) Immunostaining for COX-2 expression in
Ly6C+monocytes and DCs of the lung paren-
(E) Intracellular staining for chemokines, CXCL10
(open black) and CCL5 (open gray), along with
isotype control (shaded gray) in the lung, skin,
Figure S3 and Tables S1 and S2 accompany this
602 Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc.
macrophages. Perhaps monocytes that enter tissues on an
ongoing steady-state basis fail to differentiate to macrophages
but remain similar to blood monocytes that have been activated,
whereas in inflammation, they differentiate to macrophages. To
monocyte life cycle in the steady state.
Life Cycle and Fate of Extravascular Monocytes
that nearly completely depleted blood monocytes for 1 day with
a gradual recovery thereafter (Figure 4A). In lung, extravascular
tissue monocytes persisted for up to 3 days after monocytes
were depleted (Figure 4A). Monocyte frequency in these tissues
was normalized to that of B cells, because B cells are not tar-
geted for depletion. When we normalized to total CD45+cells
or total cells recovered, similar results were obtained. In SLNs,
monocyte numbers also did not dip significantly for 3 days (Fig-
ure 4B). However, in MLNs, depletion of blood monocytes
reduced LN monocytes by half in 1 day (Figure 4B), with slow
recovery in parallel with blood (Figure 4B).
Figure 4. Turnover of Tissue Monocytes
(A and B) Frequency of tissue monocytes, identi-
fied in tissues as in Figure 1A, was analyzed after
liposomal clodronate injection i.v. and normalized
to B cells in blood and lung (A) and LNs (B). Data
are representative of four experiments, two to four
mice per group.
tissues after anti-CD45 was given i.v. 2 min before
(D) BrdU pulse-chase analysis of tissue mono-
cytes, n = 5.
(E) Plots depict chimerism between WT congenic
parabionts at 2 weeks (left) and WT chimerism in
Ccr2?/?hosts parabiosed with a WT partner for 1
Each bar includes two to eight parabionts and
depicts mean ± SEM. Abbreviations are as fol-
accompanies this figure.
We next pulsed mice with bromodeox-
yuridine (BrdU) i.p. To be sure that tissue
monocytes evaluated were not mixed
with blood monocytes that remained
within the vasculature of tissues as they
were processed, we injected anti-CD45
mAb i.v. 2 min before euthanasia. This
technique was effective in skin and LNs
lingering blood monocytes but occasion-
ally a minor fraction (Figure 4C). The
method was not useful in lung; resident
DCs were labeled as were lung mono-
cytes (Figure 4C), indicating that anti-
CD45 mAb readily accessed the highly
vascularized lung. However, persistence
of lung monocytes after i.p. injection of
liposomal clodronate and the general
we found no
lack of blood monocyte contamination in other tissues sug-
gested that lung monocytes were not heavily contaminated. Tis-
sue monocytes did not proliferate in situ; there were very few
(Figure 4D). Peak labeling occurred in blood at 24 hr when nearly
half of monocytes were BrdU+(Figure 4D). At 48 hr, the fre-
quency of BrdU+lung and MLN monocytes mirrored that of
blood (Figure 4D). These sites showed only a slight delay in
BrdU+monocyte clearance (slope of line between 48 and
96 hr), suggesting that the residence time for monocytes was
brief, %1 day. Indeed, a rapid turnover of monocytes in the
MLN was predicted from use of liposomal clodronate (Figures
4A and 4B). Turnover of skin monocytes was different. Labeling
of skin and SLN monocytes lagged, consistent with the finding
that monocytes in SLNs were more persistent after liposomal
clodronate. SLN monocytes labeled more quickly than skin
monocytes, suggesting that some SLN monocytes arose from
blood monocytes passing into SLNs across HEV (Nakano
et al., 2009). However, clearance of BrdU+monocytes was
also slowest for SLNs and consistent with the possibility that
Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc. 603
SLN monocytes might be in part derived from skin monocytes
emigrating to the LN from skin lymphatics (Figure 4D). The fre-
quency of BrdU labeling in macrophages from skin and lung
CD11b?or CD11b+macrophages was distinct from monocytes
and consistent with a low, basal rate of proliferation that was
somewhat higher in CD11b+macrophages compared with the
more prominent CD11b?lung macrophage population (Fig-
ure S5). Because by day 10, all BrdU+monocytes had entered
and been cleared from each tissue, we were confident that we
had identified a sufficient frame of time (10 days) to track a full
tissue monocyte life cycle in skin, lung, and different LNs.
We next wondered whether some monocytes in these tis-
sues differentiated to macrophages. Few macrophages were
extracted from LNs (Figure 1C), so we focused on macrophages
in the lung, where two subpopulations are clearly evident (Fig-
of lung macrophages is embryonically derived (Hashimoto et al.,
2013; Yona et al., 2013), the life cycle of the CD11b+lung
macrophages has not been studied in detail. We carried out
parabiosis between CD45.1 and CD45.2 WT partners or WT
and Ccr2?/?partners, where WT monocytes efficiently repopu-
late the Ccr2?/?host (Hashimoto et al., 2013), examining chime-
rism at 2 weeks and 1 year after parabiosis. Neither macrophage
population (CD11b?or CD11b+) were replenished by circulating
cells, although tissue monocytes between parabionts efficiently
exchanged (Figure 4E). When we separately analyzed macro-
phages in alveolar lavage, the same outcome was observed. In
skin, by contrast, macrophages were partially replenished by
circulating precursors derived from the partner parabiont at
2 weeks (Figure 4F, left graph) and nearly fully by 1 year (Fig-
ure 4E, right graph), suggesting that monocytes are critical pre-
cursors for skin macrophages. We conclude that monocytes
enter some organs like lung constitutively for purposes other
than to replenish macrophages. Recent data also suggest they
do not give rise to DCs in the lung (Satpathy et al., 2012). Thus,
the life cycle of monocytes can include trafficking into tissues
without obligatory differentiation to macrophages or DCs. These
observations raised the question as to whether monocytes
failing to become macrophages complete their life cycle as local
monocytes that eventually die or whether they might move out of
Monocyte Trafficking to Tissues Is Not Driven
by Commensal Bacteria
The frequency of monocytes in MLNs and SLNs was greater
than in lung-draining mediastinal LNs (Figure 5A). Therefore,
we wondered whether tissues in contact with microbiota (skin
and gut) recruited Ly-6C+MHCII+monocytes resulting from
mild commensal-driven inflammatory signals. However, the fre-
quency of Ly-6C+MHCII+monocytes in theLN (Figure 5B) or skin
(Figure 5C) was similar between germ-free and conventionally
housed mice. Extravascular neutrophils were also found in skin
(Figure 1A), but in contrast to monocytes, their recruitment was
significantly reduced in germ-free mice (Figure 5D). Finally,
monocyte subset frequency in the blood of germ-free mice
showed a greater proportion of Ly-6C+monocytes (Figure 5E),
though total monocyte numbers were not different between
germ-free and conventionally housed mice, indicating that the
development of Ly-6C?monocytes from Ly-6C+monocytes
Tissue and LNs
(A) The frequency of monocytes relative to all live cells in lung-draining LNs
(LLN), skin-draining LNs (SLN), and mesenteric LNs (MLN) was quantified.
(B) Monocyte frequency in SLNs and MLNs of conventionally housed (CH) and
(B and C) Monocyte (B) and neutrophil (C) frequency in skin of CH and germ-
free (GF) mice.
(D) Blood monocyte count in CH or GF mice.
Each symbol represents data from an individual mouse with mean ± SEM
shown by bars within each data set.
604 Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc.
was at least partially regulated by microbial signals. Overall,
however, the mobilization of monocytes into tissues was unaf-
fected by the presence or absence of microbial colonization.
Monocytes Use CD62L to Enter Nonlymphoid and
Lymphoid Tissue and to Transport Antigens to LNs via
the Lymphatic Vasculature in the Steady State
and molecules involved. Ccr2?/?mice had greatly reduced
monocytes in skin (Figure 6A) and LNs (Figure 6B). By contrast,
LN Ly-6C+MHCII+monocytes were elevated in plt/plt (mice lack-
ing CCR7 ligands) and Ccr7?/?mice, consistent with previous
studies (Figure 6B; Nakano et al., 2009). Monocyte frequency
was unaffected in LNs and tissue of Cx3cr1gfp/gfpand Sell?/?
mice (Figure 6B). However, when chimeric mice were generated
byusinglethally irradiated recipientmicereconstituted with a1:1
mixture of CD45.1+WT and CD45.2+Sell?/?bone marrow (BM)
cells (WT:Sell?/?), CD45.1+WT and CD45.2+Ccr7?/?BM cells
(WT:Ccr7?/?), or CD45.1+WT and CD45.2+WT BM cells
(WT:WT), Sell?/?-derived monocytes selectively showed a
modest disadvantage in gaining access to skin-draining LNs
(Figure 6C). A disadvantage for Sell?/?monocytes was further
observed in their arrival to skin (Figure 6D), although Sell?/?
Ly-6C+MHCII+monocytes modestly accumulated relative to
WT counterparts in blood (Figure 6E). Although CD62L encoded
by Sell is well known for its role in mediating passage of leuko-
cytes across HEV, it also mediates passage into skin (Leo ´n
and Ardavı ´n, 2008). Thus, the advantage of WT-derived mono-
cytes present in the LN over Sell?/?-derived monocytes may
be due to a passage of monocytes across HEV from the blood-
stream and also an impact on monocyte trafficking first into skin
and subsequently to LNs through afferent lymphatics. To inves-
tigate the migration of undifferentiated monocytes to LNs
through afferent lymphatics, we adoptively transferred purified
Ly-6C+MHCII?, Ly-6C+MHCII+, or Ly-6C?MHCII?blood mono-
cytes into skin. Notably, only Ly-6C+MHCII+monocytes (not
MHCII?or Ly-6C?monocytes) migrated to draining LNs (Fig-
ure 6F). In LNs, these cells had lost Ly-6C expression, seemingly
different than our observations for endogenous monocytes, but
that may have stimulated internalization of surface Ly-6C.
Emigration of transferred MHCII+monocytes from skin to
SLNs (Figure 6F) and data from BrdU labeling consistent with
this concept (Figure 4E) led us to wonder whether steady traf-
ficking monocytes might survey tissues and transfer antigens
to LNs, all the while retaining close resemblance to monocytes
(Figure 3). We used an experimental approach in which FITC-
conjugated lipopolysaccharide (LPS)-free chicken ovalbumin
(OVA) administered i.n. quantitatively tracks trafficking of DCs
to LNs in the steady state (Jakubzick et al., 2008b). We asked
whether monocytes also acquired FITC-OVA in the lung and
transported it to the lung-draining lymph node (LLN) in the
absence of inflammation, how this transport compared quantita-
tively to migration of DCs transporting FITC-OVA, and how
inflammation in the lung affects transport of FITC-OVA by mini-
mally undifferentiated monocytes. For these assays, we relied
on previous insight that some sources of chicken OVA induce
inflammation but other sources do not perturb steady-state
leukocyte trafficking (Jakubzick et al., 2008b). Approximately
Figure 6. Tissue Monocytes Migrate to and from Tissue to LNs via
plots). Data are representative of two independent experiments.
(B) Scatter plot of MLN monocyte frequency in WT, Cx3cr1gfp/gfp, Ccr2?/?,
Sell?/?, plt/plt, and Ccr7?/?mice.
(C and D) Mice were reconstituted with 1:1 mixture of CD45.1+WT and
CD45.2+Sell?/?bone marrow (BM) cells (WT:Sell?/?); CD45.1+WT and
CD45.2+Ccr7?/?BM cells (WT:Ccr7?/?); or CD45.1+WT and CD45.2+WT BM
cells (WT:WT) as control. The frequency of LN (C) and skin (D) CD45.1
Ly-6C+MHCII+WT monocytes was normalized to the frequency of CD45.1
Ly-6C+MHCII+blood monocytes in the same mouse.
(E) In WT:Sell?/?chimeric mice, the frequency of blood monocyte subsets
derived from Sell?/?BM cells was plotted.
(F)WTCD45.2+blood monocytes weresortedtogeneratepurifiedpopulations
of Ly-6C+MHCII?, Ly-6C?MHCII?, and MHCII+monocytes (mix of Ly-6C+and
Ly-6C?) (top plots). Sorted CD45.2 monocytes were adoptively transferred
into the back skin of CD45.1 mice, and 24 hr later, SLNs were analyzed for
(G) Left: bar graph shows the frequency of OVA+monocytes from total
monocytes in the LLNs. Right: bar graph shows the ratio of the total number of
OVA+DCs over OVA+monocytes in the LLNs. Bars depict mean ± SEM. Data
are representative of three independent experiments.
Figure S5 accompanies this figure.
Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc. 605
6% of monocytes gated in LNs were FITC-OVA+at 24 hr after
FITC-OVA was administered i.n. (Figure 6G), thus indicating
that monocytes in the LN at least are partly sustained through
lymphatic input of monocytes. Furthermore, these data show
that monocytes participate in antigen surveillance in upstream
peripheral tissues and emigrate with this antigen to LNs without
differentiating to cells other than monocytes (such as DCs or
macrophages). To get a quantitative sense as to how this basal
transport compared to DCs, we quantified the ratio of FITC-
OVA+DCs in the same LN as FITC-OVA+monocytes. OVA+
DCs still outnumbered monocytes by ?10-fold. Nonetheless,
the data indicate that monocytes have a role in basal antigen
transport to the steady-state LN. These monocytes may stimu-
late T cell responses in LNs, as we observed the stimulation of
naive OT-I transgenic T cells in vitro when blood monocytes,
especially the MHCII+subset of Ly-6C+monocytes, were puri-
fied, pulsed with OVA ex vivo, and cultured with T cells for
60 hr (Figure S6). When inflammation-inducing OVA was used
to analyze antigen-trafficking monocytes to the draining LN,
the fraction of labeled monocytes approximately doubled but
still left DCs dominant in overall antigen transport (Figure 6G).
To provide an infectious context with more robust inflamma-
tion, we inoculated anthrax spores into the airway. CFSE was
delivered i.n. 2 or 3days after spore delivery. CFSE administered
i.n. does not enter freely into the LNs (Jakubzick et al., 2008b).
Four days after spore and CFSE delivery, CFSE+monocytes
tory DCs (Figure S7). Here, again, DCs dominated as the main
emigrating CFSE+cell type, but monocytes represented a sub-
stantial fraction of the total CFSE+migratory cells. Thus, mono-
cytes participate in steady-state tissue surveillance and antigen
transport that involves homeostatic monocyte recruitment to
various organs followed by emigration to LNs through lymphatic
vessels. In this process, monocytes undergo only limited differ-
entiation including the upregulation of MHCII, but still most
closely resemble monocytes.
Ly-6C+MHCII–Monocytes in the Blood Are Precursors
for Blood and Tissue MHCII+Monocytes
Finally, we compared the fate of Ly-6C+MHCII?versus Ly-6C+
MHCII+or Ly-6C?MHCII?blood monocytes that were isolated
and sorted from Cx3cr1gfpmice and intravenously transferred
into mice bearing congenic CD45, thus providing two criteria
for subsequent identification of rare transferred cells: GFP and
congenic CD45 alleles. Immediately after monocyte transfer,
recipient mice were given an injection of 2 mg of LPS intrader-
mally (i.d.) to enhance the attraction of grafted monocytes to
skin. In the blood, a minority of Ly-6C+MHCII?monocytes had
converted to Ly-6C+MHCII+monocytes (Figure 7A), whereas
the transferred Ly-6C+MHCII+monocytes or Ly-6C?MHCII?
monocytes had scarcely changed from their phenotype at orig-
inal sorting, although some of the transferred Ly-6C+MHCII+
monocytes had lost Ly-6C expression (Figure 7A).
Ly-6C?monocytes failed to enter SLNs (Figure 7B) but trans-
ferred Ly-6C+MHCII?blood monocytes and their separately
sorted blood MHCII+counterparts entered. Such monocytes
were MHCII+and Ly-6C+, whereas transferred Ly-6C+MHCII+
monocytes had lost Ly-6C (Figure 7B). Thus, entry into the LN
is not restricted to MHCII+blood monocytes. Transferred cells
were difficult to retrieve from the skin, but those that were
retrieved from transfer of Ly-6C+MHCII?monocytes also
showed induction of MHCII after, or in concert with, their arrival
in the skin (Figure 7C), whereas Ly-6C?monocytes did not enter
skin (Figure 7C). Taken together, these transfer data suggest
that Ly-6C+MHCII?monocytes are the precursors for blood
and tissue Ly-6C+MHCII+monocytes therein.
Figure 7. Conversion of Ly-6C+MHCII–to Ly-6C+MHCII+Monocytes
In Vivo and in Culture with Endothelium
CD115+GFP+blood monocyte subsets Ly-6C+MHCII+, Ly-6C+MHCII?, and
were sorted from CD45.2 Cx3cr1gfp/gfp
monocytes 1 3 106Ly-6C+MHCII?, 3 3 105Ly-6C+MHCII+, or 1 3 106
Ly-6C?MHCII?were injected i.v. into CD45.1+WT mice, followed by i.d.
injection of 2 mg LPS.
(A–C) At 18 hr, (A) blood, (B) skin-draining LNs (SLN), and (C) skin were
analyzed for monocyte migration and differentiation. Dot plot overlays display
totalCD11b+cells (gray) and CD45.2 GFP+adoptively transferred cells (black).
(C) Skin illustrates two separate experiments (exp 1, exp 2) adoptively trans-
(D) Purified Ly-6C+MHCII?and Ly-6C+MHCII+blood monocytes were injected
intradermally into skin and recovered 18 hr later to assess MHCII expression.
(E) Splenic Ly-6C+MHCII?monocytes, depleted completely of MHCII+cells,
were enriched by negative selection and cocultured with various cell types as
shown. Ly-6C+monocytes were gated before and after the cocultures and the
induction of surface MHCII was assessed and plotted (on the right). Each data
point represents one experiment.
606 Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc.
That Ly-6C+MHCII?monocytes upregulated MHCII after their
arrival in skin or SLN (Figures 7A–7C) seemed at odds with the
finding that injecting these cells directly into skin did not lead to
their ability to migrate to LNs, because MHCII+monocytes were
able to mobilize to LNs (Figure 6). Emigration and MHCII expres-
sion are known to be coupled (Faure-Andre ´ et al., 2008), so we
wondered whether direct injection of Ly-6C+MHCII?monocytes
in the skin might be associated with a failure of the monocytes
to upregulate MHCII. Indeed, Ly-6C+MHCII?monocytes puri-
fied from the blood and injected into skin for retrieval 18 hr later
did not upregulate MHCII (Figure 7D), in contrast to their ability
to do so if reinjected in the blood (Figure 7C), raising the possi-
bility that conversion of Ly-6C+MHCII?monocytes to Ly-6C+
MHCII+monocytes requires migration through or interaction
with the endothelium. In support of this possibility, mouse
Ly-6C+MHCII?monocytes cultured with endothelial cells, but
not fibroblasts, supported a robust upregulation of MHCII
(Figure 7E). Thus, the appearance of classical monocytes
expressing MHCII in tissues arises from the recruitment of
MHCII?rather than MHCII+monocytes from blood and the
induction of MHCII may be stimulated by transendothelial
migration. These monocytes in turn have the ability to survey
for antigens in the tissue and emigrate to lymph nodes, all the
while remaining as monocytes rather than differentiating to
macrophages or DCs.
Compelling analysis of lineage indicates that most, if not all,
organs contain a population of macrophages derived not from
monocytes but from embryonic macrophage progenitors that
develop in the embryo before definitive hematopoiesis occurs
(Hashimoto et al., 2013; Schulz et al., 2012; Yona et al., 2013).
Schulz et al. (2012) further concluded that all fetal organs con-
tained a second macrophage pool that was of bone marrow
characterized but was defined as a population with lower F4/80
and expression of MHCII, CCR2, and CD11b. Our data indicate
that this second pool in organs like lung are not macrophages
but rather monocytes that do not contribute to the resident
macrophage pool. By contrast, skin macrophages, like intestinal
macrophages (Zigmond and Jung, 2013), appear to derive at
least partly from monocytes.
subset earlier claimed to constitutively traffic into tissues from
blood (Geissmann et al., 2003), as indicated by the fact that we
did not identify Ly-6C?CD43himonocytes in extravascular tis-
sues, but instead we found classical Ly-6C+monocytes. Indeed,
the concept that resident tissue macrophages are seeded prior
to birth raises new questions about the biology of monocytes
in the absence of infection or sterile inflammation. Wynn et al.
(2013) have raised this question and suggested that monocytes
may not enter tissues constitutively, assuming that the raison
rophages derive embryonically. We have shown they do, but not
becausetheyareobliged todifferentiate tomacrophages orDCs
after diapedesis. They come into tissues, a step partially depen-
dent upon CD62L, at least in part to survey the environment and
then emigrate to LNs, and they are able to do this with only min-
imal differentiation, such that we are left to best classify them
simply as tissue monocytes.
Ly-6C+MHCII+monocyte-derived or monocyte-like cells have
been given many different names in the literature, including
myeloid-derived suppressor cells (Ochando and Chen, 2012),
DCs (Nakano et al., 2009), inflammatory monocytes (Soudja
et al., 2012), and effector monocytes (Leiria ˜o et al., 2012). Typi-
cally in these scenarios when they are studied for their role in
antigen presentation, it is in the context of inflammatory reac-
tions (Iijima et al., 2011; Leo ´n et al., 2007; Nakano et al., 2009;
Serbina et al., 2003; Soudja et al., 2012; Wakim et al., 2008). In
LNs, they have been argued to enter through the HEV (Nakano
et al., 2009), with no information on how, after entering via
such a route, that they would gain access to the antigen they
appear to present. Soluble antigen is largely sequestered in LN
conduits, and it appears that sampling of antigen from conduits
et al., 2003). In agreement with Ardavin and colleagues (Leiria ˜o
et al., 2012), we have shown here that monocytes retaining fea-
tures of minimally differentiated monocytes transport molecules
from the nonlymphoid tissues to draining LNs, but we have
extended this observation to showing its occurrence in the
steady state, with all the while the cells retaining surface and
gene expression profiles of monocytes. Consistent with this
concept, studies in rat lymph have identified monocytes in
resting lymph (Yrlid et al., 2006), but largely ignored them while
focusing on the smaller fraction of those that could be called
bona fide DCs.
Therefore, monocytes appear to share with DCs the charac-
teristic of tissue surveillance even without becoming DCs. How-
ever, the effector mechanisms used by DCs and monocytes
need not be the same. Monocytes may support antigen presen-
tation and adaptive immunity by means that do not require they
act as the antigen-presenting cell per se. Indeed, when directly
analyzed, monocytes displaying Ly-6C, MHCII, and CD11c,
perhaps similar to the cells studied here, are poor presenters
of antigen (Drutman et al., 2012), and alternative mechanisms
have been identified as to how monocytes potently support
adaptive immunity (Soudja et al., 2012). Thus, we believe that
the literature, when combined with our findings, supports a
model of the monocyte as a surveillance and effector cell in tis-
the role of lymph-homing DCs.
Although this work suggests that constitutively trafficking
monocytes are not obliged to fully differentiate to macrophages
or DCs after they enter tissues, we underscore that monocytes
can differentiate to macrophages. Thioglycollate-elicited mono-
cytes, which rapidly lose Ly-6C expression after emigrating from
the blood, cluster not with monocytes as do Ly-6C+extravas-
cular monocytes but with resident peritoneal macrophages
and they upregulate the genes universally associated with
macrophages (Gautier et al., 2012). It seems from our data that
differentiation to macrophages is not obligatory subsequent to
monocyte entry, and triggers may be necessary to drive such
differentiation. Indeed, the organs wherein macrophages are
derived from monocytes in the steady state—skin and intes-
tine—are those that receive stimulation from microbiota contin-
uously. Although infiltration of skin by monocytes was not
Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc. 607
reduced in germ-free mouse colonies, future studies will be
needed to address whether the differentiation of monocytes to
macrophages in skin does depend upon microbial stimuli.
The retention of Ly-6C expression in extravascular monocytes
may beaconvenient signature ofcellsretainingmonocyte status
in extravascular tissues. Although more profiling will be needed
to establish Ly-6C as such a transitional marker, it is very inter-
esting that recent work in the intestine has tracked the fate of
monocytes in different disease scenarios by means of Ly-6C,
MHCII, andCD64expression. Inthiscase,inhighly inflammatory
scenarios (e.g., colonic inflammation, Crohn’s disease), mono-
cytes fail to differentiate beyond the Ly-6C+MHCII+stage,
whereas in healthy colon they fully downregulate Ly-6C and
become IL-10-producing macrophages (Bain et al., 2013;
Tamoutounour et al., 2012).
Gene expression profiles were not identical between blood
and extravascular LN monocytes, though they were similar.
Prominent among upregulated genes was a common signature
between the two tissue monocytes—those from lung and
SLN—that we were able to sort in sufficiently large numbers.
This included upregulation of mRNA transcripts that are often
associated with inflammation and inflammasome or cytokine
receptor activation and transcripts often associated with DCs
(CD11c, MHCII, CD83). The mRNA transcript encoding COX-2
(Ptgr2), known to be an activation-induced transcript, was
expressed in situ, arguing that the common tissue monocyte
signature was not a result of activation during isolation and
flow cytometric cell sorting. COX-2 has been associated with
the monocyte-derived cell’s ability to produce PGE2 during
infection with Toxoplasma gondii (Grainger et al., 2013). Such
production protects the host against neutrophil-mediated dam-
age (Grainger et al., 2013) and maintenance of tissue integrity.
The emergence of COX-2 in the common tissue monocyte
signature might suggest that tissue monocytes fulfill this same
role, albeit at lower magnitude, in the steady state.
In conclusion, we have identified an extravascular population
of classical monocytes in resting tissues that in some organs
may account for the previously proposed second population of
tissue macrophages (Schulz et al., 2012). Studies in various
reporter mice and gene expression profiling support the concept
that in naive tissues, extravascular monocytes differentiate only
minimally in the steady state and do not become bona fide
macrophages by molecular criteria. Considering that the major
populations of resident macrophages do not rely on monocytes
viewed not just as precursors for either macrophages or DCs but
to macrophages in response to cues that remain to be defined.
Mice and Treatments
C57BL/6 mice were used at 6–8 weeks of age according to protocols
approved by Mount Sinai School of Medicine, University of Colorado, NIAID,
or Washington University. Strains were from Jackson Laboratory, except
germ-free (GF) mice that were bred at the Washington University Digestive
Diseases Core Center or maintained in isolators at the NIAID gnotobiotic facil-
ity. Parabiosis was carried out as described (Hashimoto et al., 2013). For BrdU
labeling, mice were injected with 1 mg of BrdU (Life Technologies Corporation)
i.p. and then euthanized at the indicated time points. BrdU incorporation into
DNA was detected with the BrdU Staining Kit (eBiosciences).
Tissue Preparation and Flow Cytometry
Perfused tissues were minced. Lung and LNs were digested with 2.5 mg/ml
collagenase D (Roche) for 30 min at 37?C. Skin was digested in RPMI 1640
plus 1.75 mg/ml Liberase TM (Roche) at 37?C for 25 min. These mAbs and
isotype-matched control mAbs purchased from Biolegend were used for
flow cytometry staining: Pacific blue-conjugated mAbs to I-A and I-E or Gr1;
PE-conjugated mAbs to CD115, Siglec F, CD11b, CD8, CD103, or I-A and
I-E; PerCP-Cy5.5-conjugated mAbs to Ly-6C, CD11b; PE-Cy7-conjugated
mAb to CD43, CD11c; allophycocyanin-conjugated mAbs to Ly-6C, CD93,
CD103, Gr1, CD64, or I-A and I-E; and allophycocyaninCy7-conjugated
mAbs to B220, NK1.1; FITC-conjugated mAbs to CD43 or CD36. F480-FITC
expression was analyzed as described (Eberlein et al., 2010).
As part of ImmGen, for cell sorting, we followed procedures are outlined in
extensive detail at http://www.immgen.org. In brief, to sort blood monocytes,
staining for CD115, CD43, Ly-6C?MHCII was done to identify monocyte sub-
sets. FSC-W and dump-channel staining for NK1.1, CD3, and B220 were used
to improve purity. Lung and LN monocytes were stained similarly but with
MerTK, CD64, F4/80, and CD11b instead of CD115, which is lost after collage-
at 37?C and cells were stained with conjugated mAbs. Blood was collected by
cardiac puncture and red blood cells were lysed and remaining leukocytes
stained for cell sorting. All cells sorted were B220?and NK1.1?to exclude B
cellsand NKcells thatreadily attach tomonocytes.Propidiumiodide excluded
dead cells. The entire procedure from the time mice were euthanized to direct
collection of cells after the second sort in TRIZOL was <3 hr.
Immunostaining of tissue sections was carried out with appropriate isotype-
matched controls or biotinylated anti-Ly-6C mAb and I-A and I-E mAbs from
Biolegend, along with anti-COX-2 mAb (clone 33) from Sigma.
Bone Marrow Transplants
8-week-old mice F1 for CD45.1 and CD45.2 congenic alleles were lethally
irradiated with two doses of 650 rad 18 hr apart. After the second irradiation,
recipient mice received 5 3 106donor bone marrow cells i.v. comprised
of 1:1 mixtures of the following genotypes: WT CD45.1:WT CD45.2, WT
CD45.1:Sell?/?CD45.2, or WT CD45.1:Ccr7?/?CD45.2. Mice were studied
8 weeks later.
Microarray Analysis, Normalization, and Data Set Analysis
Whole-mouse genome Affymetrix gene arrays were performed with the
ImmGen. RNA was amplified and hybridized on Gene Chip Mouse 1.0 ST
Arrays (data sets in Figures 3A and S3) or Gene Chip Mouse 2.0 ST Arrays
(data sets in Figures 3B and 3C). Raw data were normalized with the robust
multiarray algorithm. A common threshold for positive expression at 95% con-
fidence across the data set was determined tobe 120. Differentially expressed
probe sets between populations were selected by a Student’s t test with Bon-
alized by means of the ‘‘Heat Map Viewer’’ module of GenePattern (http://
www.broadinstitute.org/cancer/software/genepattern/). Pathway enrichment
signatures were identified with List2Network software (http://amp.pharm.
mssm.edu/lachmann/upload/register.php). Principal component analysis
(PCA) was conducted with MATLAB with RMA-normalized and log2-trans-
formed expression data.
Monocyte-Endothelial Cell Interactions
Ly-6C+MHCII-monocytes were isolated from Cx3cr1gfp/gfpspleens via nega-
tive selection with anti-CD3, anti-B220, anti-MHCII, anti-CD11c, anti-Ly6G,
and anti-siglec F magnetic beads (Miltenyi). Analysis of monocyte purity was
performed by flow cytometry analysis. A bovine type-1 collagen (Advanced
Biomatrix) bed was prepared in microplates.Bend.3 endothelial cells, kindly
provided by J. Jacobelli at National Jewish Health, were placed atop the
collagen to achieve confluency. 1 3 106enriched monocytes were added to
endothelial-collagen wells and cocultured for 18 hr at 37?C. Monocytes were
608 Immunity 39, 599–610, September 19, 2013 ª2013 Elsevier Inc.
retrieved by adding collagenase D to digest the collagen before staining and
flow cytometry analysis with gating on monocytes. Control wells used 3T3
and primary lung fibroblasts instead of endothelium
Statistical analysis by one-way ANOVA or two-tailed Student’s t test was
conducted with InStat and Prism software (GraphPad Software). All results
are expressed as mean ± SEM. A value of p < 0.05 was considered significant.
The microarray data are available in the Gene Expression Omnibus (GEO)
database (http://www.ncbi.nlm.nih.gov/gds) under the accession number
Supplemental Information includes five figures and two tables and can
be found with this article online at http://dx.doi.org/10.1016/j.immuni.2013.
WethankC.Benoistand theImmunologicalGenomeConsortium(Immgen) for
their roles in the gene expression analysis. We thank J. Faith and J. Gordon for
gnotobiotic mice, R. Schreiber for anti-IFN-g mAb H22, L. Yang for technical
assistance and advice with parabiosis, and D. Homann for assistance and
reagents used in chemokine staining. This work was supported by National
Institutes of Health grant AI049653 to G.J.R. with a Primary Caregiver’s Sup-
plement to C.J. C.J. and P.M.H. were supported by NHLBI-HL81151 and
NHLBI-HL115334. D.W.H.R. and T.C. were supported by the U.S. Department
of Defense grant DOD W81XWH-07-1-0550-Mason. E.L.G. was funded in part
by the AHA (10POST4160140). D.K.S. is supported by Training in Cancer
Biology grant T32CA009547. A.S. and F.G. were supported by a Singapore
Immunology Network core grant. The Immunological Genome project is
funded by R24 AI072073 to C. Benoist. Funding related to the use of gnotobi-
otic mice was through Washington University Digestive Diseases Research
Core Center (DDRCC) grant DK052574.
Received: October 22, 2012
Accepted: June 13, 2013
Published: September 5, 2013
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