REPORTS

Identification of Splenic Reservoir Monocytes and Their Deployment to Inflammatory Sites

Science31 July 2009
Vol. 325 no. 5940 pp. 612-616
DOI: 10.1126/science.1175202

 

Abstract

Summary Diagram
In response to heart injury (myocardial infarction), specific subsets of monocytes are recruited from the bone marrow and spleen to remove and repair damaged tissue.

A current paradigm states that monocytes circulate freely and patrol blood vessels but differentiate irreversibly into dendritic cells (DCs) or macrophages upon tissue entry. Here we show that bona fide undifferentiated monocytes reside in the spleen and outnumber their equivalents in circulation. The reservoir monocytes assemble in clusters in the cords of the subcapsular red pulp and are distinct from macrophages and DCs. In response to ischemic myocardial injury, splenic monocytes increase their motility, exit the spleen en masse, accumulate in injured tissue, and participate in wound healing. These observations uncover a role for the spleen as a site for storage and rapid deployment of monocytes and identify splenic monocytes as a resource that the body exploits to regulate inflammation.


See Also:
Perspective | Immunology

Dispensible But Not Irrelevant

Editor's Summary:

Monitoring Monocyte Reservoirs

Monocytes are cells of the immune system that are recruited to sites of tissue injury and inflammation where they help to resolve the infection and are important for tissue repair. The bone marrow and blood are believed to be the primary reservoirs from which monocytes are mobilized after injury. Swirski et al. now demonstrate that the spleen also serves as a critical reservoir of monocytes that are recruited during ischemic myocardial injury. Monocytes in the spleen are very similar in phenotype to blood-derived monocytes and are mobilized to the injured heart, where they represent a large fraction of the total monocytes that are recruited. The chemoattractant, angiotensin II, is required for optimal monocyte mobilization from the spleen and emigration into injured tissue.

Figures and Selected Supplementary Material

 

Figure: 1A

Total number of Ly-6Chigh and Ly-6Clow monocytes in the infarcted myocardium and (2 ml) peripheral blood (means ± SEM, n = 9 to15). Monocytes were identified as CD11bhigh, Linlow, and (F4/80, I-Ab, CD11c)low. Lin refers to the combination of CD90, B220, CD49b, NK1.1, and Ly-6G monoclonal antibodies.

Author Profile

Filip Swirski
 

Filip Swirski

Center for Systems Biology,
Massachusetts General Hospital and Harvard Medical School

Filip Swirski received his PhD in 2004 in Immunology at McMaster University, Canada. In 2007, he completed his postdoctoral studies in Vascular Biology at Brigham and Women’s Hospital and Massachusetts General Hospital (MGH), and became Faculty at the Center for Systems Biology (CSB), MGH, and Harvard Medical School. The central theme of his laboratory within the Immunology Program at CSB rests on the hypothesis that leukocytes promote organ co-operation and thus are key architects of organ systems. This is fundamental to leukocytes and encompasses such distinct, if not exclusive, properties as motility, plasticity and clonality. His current interests include immune regulation in atherosclerosis.

 

Matthias Nahrendorf

Center for Systems Biology,
Massachusetts General Hospital and Harvard Medical School

Matthias Nahrendorf attended Medical School at Heidelberg University, followed by residency in Internal Medicine and fellowships at the Biophysics Department in Wuerzburg and the Center for Molecular Imaging in Boston. Since 2006, he is Faculty at the Center for Systems Biology, the Director of the Mouse Imaging Program, and a member of the Immunology Program at MGH. His laboratory focusses on imaging of molecular processes in heart failure, atherosclerosis and transplant rejection. Imaging targets are enzymes, immune cells and molecular players with a central role in cardiovascular disease. The Nahrendorf laboratory uses the entire spectrum of modalities, including MR, nuclear, optical and hybrid imaging, to gain insight into inflammation and tissue repair at a systems level, and in an undisturbed in vivo environment.

Ralph Weissleder
 

Ralph Weissleder

Center for Systems Biology,
Massachusetts General Hospital and Harvard Medical School

Dr. Weissleder is a Professor at Harvard Medical School, Director of the Center for Systems Biology at Massachusetts General Hospital (MGH), and Attending Clinician (Interventional Radiology) at MGH. Dr. Weissleder is also a member of the Dana Farber Harvard Cancer Center, an Associate Member of the Broad Institute (Chemical Biology Program) and a member of the Harvard Stem Cell Institute (HSCI) leading its Imaging Program. His work has been honored with numerous awards and he is a member of the US National Academies Institute of Medicine. Dr. Weissleder’s research interests include the development of novel molecular imaging techniques, tools for detection of early disease, and the development of nanomaterials for sensing and systems analysis.

Mikael Pittet
 

Mikael Pittet

Center for Systems Biology,
Massachusetts General Hospital and Harvard Medical School

Mikael Pittet obtained his PhD in 2001 at the Ludwig Institute for Cancer Research, Lausanne, Switzerland, and trained at MGH, Harvard Medical School and the Dana Farber Cancer Institute, Boston, USA. Since 2006, he is Faculty member at the MGH Center for Systems Biology. His laboratory studies responses mediated by innate and adaptive immune cells, which play a central role in the orchestration and resolution of tissue inflammation. Current interests include the role of these cells in controlling the delivery of protective immune responses to tissues, and their contributions in inflammatory diseases, including cancer. The laboratory is part of the Immunology Program at CSB, and collaborates with several immunology programs at Harvard Medical School, MGH and Massachusetts Institute of Technology.

 

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Received for publication 20 April 2009. Accepted for publication 10 June 2009.

Abstract

Summary Diagram
In response to heart injury (myocardial infarction), specific subsets of monocytes are recruited from the bone marrow and spleen to remove and repair damaged tissue.

A current paradigm states that monocytes circulate freely and patrol blood vessels but differentiate irreversibly into dendritic cells (DCs) or macrophages upon tissue entry. Here we show that bona fide undifferentiated monocytes reside in the spleen and outnumber their equivalents in circulation. The reservoir monocytes assemble in clusters in the cords of the subcapsular red pulp and are distinct from macrophages and DCs. In response to ischemic myocardial injury, splenic monocytes increase their motility, exit the spleen en masse, accumulate in injured tissue, and participate in wound healing. These observations uncover a role for the spleen as a site for storage and rapid deployment of monocytes and identify splenic monocytes as a resource that the body exploits to regulate inflammation.


See Also:
Perspective | Immunology

Dispensible But Not Irrelevant

1. Introduction

Protection of injured or infected tissue involves migratory leukocytes (1–3). Among them are blood monocytes, which consist of at least two functionally distinct subsets (4, 5). Ly-6Chigh (Gr-1+) monocytes are inflammatory and migrate to injured (6, 7) or infected (8–10) sites but also propagate chronic diseases (1113). Ly-6Clow (Gr-1) monocytes patrol the resting vasculature (14), populate normal (15) or inflammatory sites (14), and participate in resolution of inflammation (7).

2. The spleen contains a large reservoir of monocytes that resemble blood monocytes in morphology, phenotype and gene expression.

Tissue repair after myocardial infarction (MI) requires coordinated mobilization of both subsets: first, Ly-6Chigh monocytes digest damaged tissue; second, Ly-6Clow monocytes promote wound healing (7). We observed that the ischemic myocardium accumulates Ly-6Chigh monocytes in numbers that exceed their availability in circulation (Fig. 1A), which intrigued us. Although the bone marrow produces and contains numerous (pro)monocytes (16), we sought to identify compartmental reservoirs of extramedullary monocytes, as these could accommodate the demands of rapid-onset inflammation.

First, we screened candidate tissues for the presence of monocyte-like cells. Monocytes express CD11b and CD115 and are negative or low for CD90, B220, CD49b, NK1.1, and Ly-6G surface proteins. They are distinct from macrophages and dendritic cells (DCs) on the basis of the F4/80 glycoprotein, CD11c, and major histocompatibility complex molecule I–Ab. Of all organs profiled, only the spleen contained cells that met these criteria and that were present in large quantities. The population included Ly-6Chigh and Ly-6Clow subtypes in ratios similar to those in the blood (Fig. 1, B and C, fig. S1 and movie M1).

The spleen's major known functions are removal of aging erythrocytes and recycling of iron, elicitation of immunity, and supply of erythrocytes after hemorrhagic shock (18). The presence of numerous monocytes in the spleen seems paradoxical, because monocytes are distinguished from lineage descendants on the basis of residency: Monocytes are considered circulating, whereas macrophages and DCs are tissue-resident and predominantly sessile (5, 14, 19). We therefore conducted additional experiments to characterize the monocyte-like cells in the spleen.

We found that splenic monocytes resembled their blood counterparts morphologically: Ly-6Chigh monocytes were larger than Ly-6Clow monocytes, and both subsets had kidney- or horseshoe-shaped nuclei (Fig. 1D). Ly-6Chigh monocytes in blood and spleen had essentially indistinguishable transcriptomes (Fig. 1E and tables T1 and T2 ). Refined mRNA and protein analysis validated this similarity (Fig. 1F), although, as expected, Ly-6Chigh monocytes differed from their Ly-6Clow counterparts (5) (Fig. 1F and table T3). Splenic Ly-6Chigh and Ly-6Clow monocytes had phagocytic functions similar to those of their blood counterparts (Fig. 1G) and differentiated comparably into macrophages or DCs in vitro (Fig. 1H). We therefore concluded that the spleen contains a population of bona fide monocytes that coexists with, but is different from, macrophages and DCs and that outnumbers blood monocytes.

3. Splenic monocytes are located in the subcapsular red pulp.

To determine where monocytes reside in the spleen, we used Cx3cr1gfp/+ mice in which nearly all fluorescent splenocytes are monocytes or their lineage descendants (Fig. 2A and figs. S2 and S3). We detected dense populations of green fluorescent protein–positive (GFP+) cells in two regions, the marginal zone and the subcapsular red pulp. As expected, cells in the marginal zone were mostly macrophages and DCs (18): They were large and morphologically irregular; F4/80high or CD11chigh, respectively; and localized around the white pulp (Fig. 2B, fig. S3, and movie M1). The fluorescent cells in the subcapsular red pulp were mostly monocytes: They were devoid of dendrites; had kidney- or horseshoe-shaped nuclei; were CD11b+ and F4/80–/low but CD11c; and were arranged in clusters of ~20 to 50 cells along the perimeter of the organ (Fig. 2, C to E, fig. S3, and movies M2 and M3). They clustered mostly in the reticular fiber–rich pulp cords, just as iron-recycling CD163+ macrophages do (20) (Fig. S3). Also, intravital microscopy and parabiosis experiments revealed that splenic monocytes resided in the spleen, rather than simply passed through the spleen within blood (Figs. S4 and S5).

4. Splenic monocytes are mobilized to the heart in response to ischemic myocardial injury.

A hallmark of a reservoir population is its ability to deploy to distant sites. Thus, we tested whether splenic Ly-6Chigh monocytes are mobilized in response to surgically induced ischemia of the myocardium. One day after coronary ligation, we observed reduced numbers of monocytes in the subcapsular red pulp of the spleen (Fig. 3, A to C) that could not be attributed to local cell differentiation or death (Fig. S6) and, therefore, indicated exit. Entire organ enumeration after MI revealed monocyte loss in spleen, gain in blood, but no change in the bone marrow (Fig. 3D and fig. S6), which suggested mobilization of splenic, but not bone marrow, monocytes after tissue injury.

We next compared the relative contributions of the spleen and the bone marrow in response to MI by using mice in which only one of the two tissues can contribute monocytes. First, we evaluated Ccr2–/– mice, because the chemokine receptor mediates monocyte mobilization from the bone marrow (9, 21), but not from the spleen (table S4). The number of blood monocytes comparably increased—and the number of splenic monocytes comparably decreased—after MI in both wild-type and Ccr2–/– mice (Fig. 3E). The released monocytes in Ccr2–/– mice did not accumulate in the ischemic myocardium, because infiltration depends on the chemokine CCR2 [table S4 and (7)]. Second, we evaluated animals splenectomized by a procedure that preserves the bone marrow and blood monocyte pools (Fig. S7). After MI, blood monocyte numbers increased in control, but not splenectomized, animals (Fig. 3F). Analysis of the ischemic myocardium revealed a massive influx of Ly-6Chigh monocytes in mice containing the spleen. However, this number was decreased by 75% in splenectomized mice (Fig. 3, G and H). These findings indicate that the spleen mobilizes monocytes en masse after MI. Similar observations in rats argue for a generalizable existence of a splenic monocyte reservoir (Fig. S8).

To track unambiguously the fate of monocytes from the spleen to the heart, we studied CD45.1 mice that were splenectomized, given CD45.2 spleens by transplantation, and subjected to MI (Fig. S9). We observed increased numbers of donor monocytes in the blood of animals after MI, which indicated that the injury triggered the release of splenic monocytes (Fig. S10). The infarct accumulated donor (i.e., spleen-derived) and host monocytes, all of which were Gr-1+ (Ly-6Chigh) (Fig. 3I). The transplantation procedure itself reduced the number of reservoir donor monocytes by a factor of 6.1, likely because of ischemia-related cell differentiation. Correcting for this, we calculated that the spleen contributed 41% of monocytes to the ischemic myocardium. Control experiments (transplanted pancreas or transplanted spleen but no MI) showed no accumulation of donor monocytes in the recipient heart (Fig. 3J). Noninvasive fluorescence molecular tomography–magnetic resonance imaging (FMT-MRI) three-dimensional fusion imaging (22) with phagocytosis and cathepsin-activatable sensors (23) revealed attenuated activities in infarcts of splenectomized mice (Fig. 3K), which indicates that splenic monocytes are biologically active when recruited to inflamed tissue. Thus, the spleen stores Ly-6Chigh monocytes readily recruitable to augment inflammation at distant sites. The spleen does not likely produce monocytes, because donor spleens contained only host-derived cells as early as 3 weeks after transplantation (Fig. S11).

The spleen did not contribute neutrophils significantly, which suggested selective mobilization of monocytes (Fig. S12). Of these, both subsets (Ly-6Chigh and Ly-6Clow) exited the spleen in response to MI, yet after 1 day, the ischemic myocardium recruited Ly-6Chigh monocytes selectively (Fig. S13). The excluded Ly-6Clow monocytes may have dispersed to other tissues, patrolled the vasculature, or accumulated in the infarct at a later time (7).

5. Angiotensin II-dependent signaling promotes monocyte exit from the spleen and emigration to inflammatory sites.

Monocytes express a wide variety of receptors (24), some of which may trigger splenic release. We focused on angiotensin II (Ang II), because (i) it induces cytoskeletal rearrangement and migration of monocytes in vitro (25), (ii) it augments monocyte-mediated inflammation (26), and (iii) its serum levels increase after MI (27). Ang II exerts its effects by binding to the angiotensin type 1 (AT-1) and AT-2 receptors.

Atgr1a–/– animals subjected to MI did not expel splenic monocytes efficiently (Fig. 4A) and accumulated only a few monocytes in the ischemic myocardium (Fig. S14), which indicated that Ang II–AT-1 signaling contributes to expulsion in this model. Sustained exogenous administration of Ang II in wild-type mice reproduced monocyte egress; mice with Ang II concentrations comparable to those after MI (Fig. 4B) released splenic monocytes (Fig. 4C and fig. S15 ). Moreover, Ang II infusion or MI elicited AT-1 receptor dimerization in splenic monocytes in vivo (Fig. 4D), an event that stimulates a wide spectrum of intracellular responses (26). We also found that Ang II induced directional migration of splenic monocytes in vitro (Fig. 4E).These data support a direct link between Ang II, the AT-1 receptor, and splenic monocytes and prompted us to explore the effects of Ang II on cell behavior in vivo.

We developed a real-time intravital microscopy technique that allows observation of endogenousS monocytes and vessels in the subcapsular red pulp of the spleen in live animals, at depths up to 100 µm below the fibrous capsule, while preserving organ temperature and blood flow. Spleens of Cx3cr1gfp/+ mice contained three distinct GFP+ populations, based on their location and size (Fig. 4F, fig. S16, and movies M4 to M6). Real-time tracking of GFP+ cells in cluster-rich regions of the subcapsular red pulp revealed behavioral changes shortly after MI or after administration of Ang II (Fig. 4G). Splenic monocytes increased their displacement over time by more than threefold within 2 hours after either intervention (Fig. 4, G to I, movies M7 to M9 and fig. S17), which indicated that Ang II induced their migration in vivo. Conversely, splenic macrophages or DCs showed very low displacement that did not increase in response to intervention, whereas patrolling monocytes showed typically high displacement (Fig. 4I), as reported in dermal and mesenteric vessels (14). The motile splenic monocytes that responded to Ang II were more likely to encounter neighboring venous sinuses or collecting veins and to enter the blood stream to exit the spleen. Figure 4J and movie M10 show one example of a prototypical departing monocyte. The increased motility of splenic monocytes and subsequent egress led to a considerable loss of fluorescent cells in tissue (Fig. S15).

Splenic contraction after hemorrhagic shock is associated with erythrocyte expulsion (28). Our intravital microscopy data show, however, that the subcapsular red pulp did not measurably contract when monocytes were already activated 1.5 hours after treatment with Ang II (Fig. S18). A contraction-induced mechanism would also affect other leukocytes, but our neutrophil data indicate otherwise.

Our findings illuminate the body's ability to mobilize a reservoir of undifferentiated splenic monocytes in response to injury.Triggering of this reservoir likely provides a stochastic advantage for rapid monocyte accumulation, but such triggering is not necessarily desirable. Future studies should investigate the contribution of the splenic monocyte population in response to other inflammatory events and whether additional factors control monocyte migration, organization, and differentiation in the splenic environment. Understanding how an organism controls the quality and quantity of its immune players is essential to understanding homeostasis, and its perturbation and restoration following infection and tissue injury.


Received for publication 20 April 2009. Accepted for publication 10 June 2009.

Figures (collapse all 4 figures)

Figure 1 (A to H): The resting spleen contains a large reservoir of bona fide monocytes.
Figure 1 (A to H): The resting spleen contains a large reservoir of bona fide monocytes.

(A) Total number of Ly-6Chigh and Ly-6Clow monocytes in the infarcted myocardium and (2 ml) peripheral blood means ± SEM, n = 9 to 15). Monocytes were identified as CD11bhigh, Linlow, and (F4/80, I-Ab, CD11c)low. Lin refers to the combination of CD90, B220, CD49b, NK1.1, and Ly-6G monoclonal antibodies. (B) Total monocyte number retrieved from different tissues (means ± SEM, n = 3 to 5). (C) Flow cytometric analysis of blood and splenic Ly-6Chigh and Ly-6Clow monocytes, as well as CD11b+ macrophages and DCs (Mø/DCs) (n = 5). (D) Cytospin preparations of Ly-6Chigh and Ly-6Clow monocytes from blood and spleen. Insets indicate forward (FSC) and side (SSC) scatters identified by flow cytometry (means ± SEM, n = 3). (E) Microarray analysis of Ly-6Chigh monocytes from blood and spleen. Data depict the average log2-based intensity of the same probe across four replicates. Also refer to tables T1 and T2. (F) Expression of 45 genes (by quantitative reverse transcription polymerase chain reaction) and 11 proteins (by flow cytometry) for Ly-6Chigh and Ly-6Clow monocytes in blood and spleen (n = 4). Also refer to table T3. (G) Ex vivo phagocytosis of beads by Ly-6Chigh and Ly-6Clow monocytes and T cells (control) from blood and spleen [mean fluorescent intensity (MFI) ± SEM, n = 3]. (H) In vitro differentiation of blood and splenic Ly-6Chigh and Ly-6Clow monocytes to macrophages (F4/80high) and DCs (CD11chigh) in response to macrophage colony-stimulating factor (M-CSF) and granulocyte-macrophage CSF (GM-CSF) + interleukin-4 (IL-4), respectively (MFI ± SEM, n = 3 to 9). Data pool at least three independent experiments [(A) and (H)], or are from one experiment representative of at least three independent experiments [(C) and (D)].
Figure 2 (A to E): Splenic monocytes cluster in the subcapsular red pulp.
Figure 2 (A to E): Splenic monocytes cluster in the subcapsular red pulp.

(A) Histograms depict flow cytometric analysis of GFP fluorescence of splenic leukocytes in Cx3cr1gfp/+ mice (n = 3). The pie chart shows the relative proportions of various GFP+ populations. Also refer to Figs. S2 and S3. (B and C) Multiphoton microscopy micrographs of Cx3cr1gfp/+ mice show GFP+ Mø/DCs (green) in the marginal zones (B) and GFP+ monocytes (green) in the subcapsular red pulp (C). Collagen fibers appear in blue. (D) A single optical section shows GFP+ monocytes (green) in the subcapsular red pulp. The dense collagen network (blue) of the splenic capsule indicates the boundary of the organ. (E) An intravital micrograph of the spleen subcapsular red pulp shows GFP+ monocytes (green) organized in clusters and topographically distinct from blood vessels (red). All data are from one experiment representative of at least two independent experiments.
Figure 3 (A to K): Splenic reservoir monocytes emigrate from the subcapsular red pulp & populate inflammatory sites.
Figure 3 (A to K): Splenic reservoir monocytes emigrate from the subcapsular red pulp & populate inflammatory sites.

(A) Splenic sections from mice without MI and 1 day after MI stained with hematoxylin and eosin. (B) Splenic sections stained with CD11b-specific antibodies (red) and 4',6'-diamidino-2-phenylindole (DAPI) (blue) depict the subcapsular red pulp and the marginal zone from mice without MI and 1 day after MI. (C) Enumeration of CD11b+ cells in the subcapsular red pulp and marginal zone (means ± SEM, n = 10 high-power fields). (D) Total number of monocytes in the spleen, blood, or bone marrow (tibia) in control mice (no MI) or 1 day after MI (means ± SEM, n = 6 to 15). (E) Total number of monocytes in the spleen or blood in wild-type (WT) and Ccr2-/- mice in response to MI (n = 3 to 6). (F) Total number of blood monocytes in splenectomized animals (-Spleen) and sham-operated controls (+Spleen) without MI or 1 day after MI (n = 3 to 6). (G and H) Accumulation of cells in heart in the same groups of mice as measured by flow cytometry [(G), n = 3] or by counts per mg tissue [(H), means ± SEM, n = 9]. (I) Accumulation of monocyte subsets originating exclusively from spleen as measured by flow cytometry. Dot plots (CD45.1 versus CD45.2) of gated monocytes and histograms (Gr-1) of each positive population. (J) CD45.1 versus CD45.2 profile of gated monocytes in a mouse receiving a control transplant (pancreas) and in a mouse not subjected to MI. Also refer to Fig. S7. (K) FMT-MRI of in vivo phagocytic and proteolytic activities in infarcts. Fluorochrome concentration is shown in the infarcted area, on the basis of MRI-derived anatomy (n = 6). FI, fluorescence intensity. *P < 0.05; **P < 0.005. Data pool at least three independent experiments (D to F, and H), or are from one experiment representative of two [(I) and (J)] or at least three independent experiments [(A) to (C), and (G)].
Figure 4 (A to J): Ang II-AT-1 receptor signaling promotes splenic monocyte motility and tissue emigration.
Figure 4 (A to J): Ang II-AT-1 receptor signaling promotes splenic monocyte motility and tissue emigration.

(A) Percentage of monocytes lost from the spleen 1 day after MI or Ang II infusion in WT and Atgr1a-/- mice (means ± SEM, n = 4 to 9). (B) Serum Ang II concentrations in the steady-state (control), and 1 day after MI or infusion of Ang II (n = 6 to 9). (C) Total number of splenic monocytes (Ly-6Chigh and Ly-6Clow) in the groups of mice mentioned above. (D) Western blot analysis of monomeric and dimeric forms of the AT-1 receptor on control splenic monocytes in the steady state (control), and 1 day after MI or infusion of Ang II, n = 3. (E) In vitro migration of splenic monocytes in response to Ang II (1 µM) (means ± SEM, n = 6). (F) Intravital microscopy of GFP+ cells (green) in the spleen subcapsular red pulp of an Ang II-treated Cx3cr1gfp/+ mouse. Images show a splenic monocyte, an Mø or DC, and a patrolling monocyte. Blood is shown in red. Tracks indicate the position of cell centroids at 15-s intervals (time in min:s). (G) (Top) Intravital micrographs of GFP+ cells in control mice and 1 day after MI or infusion of Ang II at the initial recording time point. (Bottom) Tracks for all GFP+ cells in the field of view and over 1 hour. Some cells entered or exited our imaging area during the recording and thus were followed for a shorter duration. V, vessel; P, parenchyma. (H) Average displacement over time of all GFP+ splenic monocytes [means ± SEM, n = 143 (control), 163 (MI), and 125 (Ang II) cells]. (I) Displacement over time of single splenic monocytes (left), splenic macrophages of DCs (middle), and patrolling monocytes (right). (J) Recording of a departing splenic monocyte. Tracks indicate the position of the cell centroid. *P < 0.05, **P < 0.01, ***P < 0.001. Data pool two [(A) to (C), and (E)] or at least three independent experiments [(H) and (I)], or are from one experiment representative of at least three independent experiments [(D), (F), and (G)].

Supplementary Figures (expand all 18 supplementary figures)

Figure S1 (A to D): Co-existence of macrophages/DCs and undifferentiated monocytes in the spleen.
Figure S1 (A to D): Co-existence of macrophages/DCs and undifferentiated monocytes in the spleen.

Leukocytes were retrieved from the spleen and labeled with antibodies as described in Methods. (A) Gating on leukocytes based on CD11b, Lin (i.e. CD90/B220/CD49b/NK1./Ly-6Glow), F4/80 and CD11c expression identifies at least eight different populations, two of which (CD11b+ Linlow F4/80low CD11c- Ly-6Chigh/low) are monocytes. (B) Enumeration of the eight populations reveals different numbers of monocytes, macrophages (Mø; defined by F4/80 expression), DCs (DC; defined by CD11c expression) or other cells (many of which are lymphocytes) (n=3). (C) Enumeration of all monocytes vs all macrophages or DCs in the spleen (n=3). (D) CD11bhigh Linlow monocytes correspond to CD11bhigh CD115+ monocytes. The dot plot overlay shows that 97.5% of gated CD11bhigh CD115+ cells (blue dots) overlay with CD11bhigh Linlow monocytes. All data are from one experiment representative of at least three independent experiments.
Figure S2: Flow cytometry identification of various immune cell types obtained from the spleen of Cx3cr1gfp/+ mice.
Figure S2: Flow cytometry identification of various immune cell types obtained from the spleen of <em>Cx<sub>3</sub>cr1<sup>gfp/+</sup></em> mice.

Monocytes were defined as CD11bhigh Linlow (F4/80/CD11c/I-Ab) low and were further divided into Ly-6Chigh/low subsets. DC were defined as CD11c+ cells; Mø as F4/80+; NK cells as (NK1.1/CD49b)high; T cells as CD3+; B cells as B220high; and neutrophils as SSChigh Ly-6Ghigh. EGFP expression was analyzed for each cell type; mean fluorescence intensities (MFI) are shown on the right (n=3). Of note, 6±3% NK cells were EGFP+, whereas virtually all monocytes were EGFP+. When taking into account the abundance of each cell population and the fractions that are EGFP+, our flow cytometry data indicate that EGFP+ monocytes outnumber EGFP+ NK cells by > 12 to 1 in the spleen (i.e., 92% EGFP+ monocytes for 8% EGFP+ NK cells). All data are from one experiment representative of at least two independent experiments.
Figure S3 (A to C): Monocytes reside in the subcapsular red pulp.
Figure S3 (A to C): Monocytes reside in the subcapsular red pulp.

(A) Immunofluorescence panels of spleens from Cx3cr1gfp/+ mice flash-frozen and stained with anti-CD11b, F4/80, CD11c, CD163 or CD49b (red) antibodies and DAPI (blue), and co-localized with GFP (green). The subcapsular red pulp (left three panels) and marginal zones (right three panels) are shown; srp = subcapsular red pulp, mz = marginal zone, wp = white pulp. We used the same exposure times to capture fluorescence of a given Ab in SRP and MZ. Monocytes were GFPhigh, CD11b+, F4/80low, CD11c- and CD163- and were found predominantly in the subcapsular red pulp. GFP- CD11bhigh population represents neutrophils. Very few CD49b+ GFP+ NK cells were observed in the subcaspular red pulp. CD163 identifies iron-recycling macrophages that reside predominantly in the SRP. (B) Negative controls were generated by staining with the appropriate secondary antibodies only. (C) Signal intensity of GFP, CD11b, F4/80 and CD11c in various regions of the spleen. The numbers were generated by drawing same-area regions of interest with ImageJ on regions either in the subcapsular red pulp or the marginal zone that were positive for each signal, so as to calculate intensity of a given signal as a measure of expression for that particular marker (note that exposure times were kept the same for any given marker in all regions). The data show that GFP+ cells are most bright in the subcapsular red pulp, in keeping with flow cytometry data in Fig. 2A that show that monocytes are brightest compared to macrophages. Also note that F4/80 and CD11c expression is brightest in the marginal zone, in keeping with flow cytometry data of Fig. S2, which show that mature macrophages express F4/80 higher than monocytes (notably, monocytes are F4/80-/low) and mature DCs express CD11c higher than monocytes (monocytes are predominantly CD11c-; see Figure 1H). CD11b and CD163 expression are similarly bright in both regions. *P < 0.05; **P < 0.001 (n=10 high power fields). All data are from one experiment representative of at least two independent experiments.
Figure S4 (A to B): Splenic monocytes reside in the spleen and thus do not traffic through the spleen while in the blood stream.
Figure S4 (A to B): Splenic monocytes reside in the spleen and thus do not traffic through the spleen while in the blood stream.

(A) Intravital microscopy investigation of splenic monocytes in mice devoid of blood leukocytes. (1) Cx3cr1gfp/+ mice received an intravenous injection of a fluorescent blood pool agent (AngioSense-680). (2) Intravital imaging of the spleen subcapsular red pulp shows blood vessels (red), as well as monocyte clusters (green) outside the vessels. (3) Blood from these mice was then removed by injecting 20 ml saline into the left ventricle. Blood was allowed to exit on the venous side through a small excision performed in the right atrium. The inserts on the lower right of the quadrant show pictures of venous samples that were obtained before and after the flushing procedure. Note that the samples obtained after the flushing procedure are clear and virtually devoid of leukocytes (flow cytometry analysis indicates that the few remaining cells are all red blood cells). Thus, the perfused mice did not contain circulating blood monocytes. (4) The same mice were subjected again to imaging. As expected, the extensive flushing of the vessels with saline removed the blood pool agent in the spleen vessels. Importantly, however, the subcapsular red pulp retained splenic monocytes. The topography of these cells is indistinguishable from the one observed before perfusion. (B) Flow cytometry analysis of blood samples and spleen in mice that were flushed with 20 ml saline or not. Blood data show the number of monocytes per ml of blood (before flush), or per ml of saline (after flush). Spleen data show the total number of monocytes in spleens of animals that were flushed or not (n=3-5).
Figure S5: Monocyte exchange in parabiotic mice during parabiosis and after surgical separation.
Figure S5: Monocyte exchange in parabiotic mice during parabiosis and after surgical separation.

The cartoons illustrate different groups of mice that were subjected to parabiosis. Parabiosis was established between mice expressing two distinct CD45 allotypes (CD45.1 and CD45.2). Mice were divided into 3 groups (4 mice per group). The first and second groups were sacrificed 2 and ~30 days post joining. Blood mixing initiates at day ~2 and thus is useful to measure monocyte exchange between parabionts. The percent chimerism of monocytes has reached a plateau by day ~30 (37). The third group of mice involved separation of parabionts at day ~30 and sacrifice 18h later. Separation of mice prevents further exchange between parabionts and thus is useful to measure the rate of monocyte turnover and replacement. The graphs at bottom depict the percent chimerism of monocytes in blood and spleen of each parabiont.The fold difference in percent chimerism between blood and spleen is shown on top of each graph (e.g., 13:1 means that the percent chimerism is 13-times higher in blood than in spleen). The experiment reveals that, early after joining, the percent chimerism of monocytes was lower in spleen than in blood, indicating that monocytes actively seeded the splenic niche. Monocytes also resided longer in spleen than in blood, because the percent chimerism of monocytes was higher in spleen than in blood in long-term parabionts that had been separated recently. All data pool at least two independent experiments.
Figure S6 (A to C): The decreased number of splenic monocytes after MI is not linked with increased cell death or with an increased number of splenic Mø/DC.
Figure S6 (A to C): The decreased number of splenic monocytes after MI is not linked with increased cell death or with an increased number of splenic Mø/DC.

(A) Total number of splenic monocytes in resting mice ('no MI'), or 1 day post MI. (B) Percentage of splenic monocytes that are Annexin-V+ or Propodium Iodide+. (C) Total number of splenic Mø/DC. The results suggest that after MI, splenic monocytes exit the spleen rather than die or mature into Mø/DC locally (n=3). **P < 0.005.
Figure S7 (A to B): Splenectomized animals show normal monocyte and lymphocyte counts in blood and bone marrow.
Figure S7 (A to B): Splenectomized animals show normal monocyte and lymphocyte counts in blood and bone marrow.

(A) Number of monocytes per ml of blood (left) or of bone marrow monocytes per tibia (right) in control mice('+Spleen') or in splenectomized animals ('-Spleen'). (B) Number of blood and bone marrow lymphocytes in the same group of mice. Mice were analyzed by flow cytometry. Splenectomized mice were sacrificed 1 day after surgery (n=3).
Figure S8 (A to F): Analysis of monocytes in rats subjected to MI.
Figure S8 (A to F): Analysis of monocytes in rats subjected to MI.

(A) Flow cytometry analysis of spleens from control rats and from rats subjected to MI. (B) Total count of CD11b+ cells in spleen as identified in panel A. (C) Flow cytometry analysis of enzymatically-digested hearts 1 day after MI in rats with or without spleen. (D) Total count of CD11b+ cells in hearts as in panel C. n=3-4. (E) Fluorescence reflectance imaging (FRI) of explanted hearts 1 day after MI obtained from rats with or without spleen. Rats received an intravenous injection of CLIO-VT680 at the time of MI (e.g., 1 day before sacrifice). FRI at 680 nm wavelength measures fluorescent nanopartice uptake; FRI at 488 nm wavelength measures autofluorescence. (F) Target to background ratio in the infarcted hearts shown in panel E (n=3-4). *P < 0.05; ***P < 0.0005.
Figure S9: Spleen transplantation with vascular anastomosis.
Figure S9: Spleen transplantation with vascular anastomosis.

A spleen transplant model was developed to quantitatively track splenic monocytes after induction of myocardial infarction. (1) The spleen was harvested from a CD45.2+ mouse after inhibition of clotting with heparin. Before explantation, the donor animal was perfused with saline until no blood remained in circulation. (2) The donor spleen was then carefully exposed using micro-dissection tools and a surgical microscope. To facilitate vascular anastomosis to the recipients' circulation, the splenic artery and vein were prepared while they remained connected to the abdominal aorta and the portal vein. The use of this vasculature with a much higher caliber as a conduit facilitated vascular anastomosis of the splenic artery and vein to the recipient. Side branches and other major vessels were carefully ligated to avoid bleeding. (3) The organ was then transplanted into the intraperitoneal cavity of the CD45.1+ recipient, and arterial and venous anastomoses were produced using microsurgical techniques. End-to-side anastomoses of the aortic cuff to the recipient descending aorta, and of the portal vein to the recipient inferior vena cava were established while these vessels were clamped. After unclamping, flow through the transplanted organ was verified visually, followed by splenectomy of the orthotopic CD45.1 recipient spleen. Vascular anastomosis of splenic vessels is shown on the digital image of the surgical field on the right. To better visualize the anatomy, the spleen was flipped over to the right side of the animal. (4) The abdomen was then closed and myocardial infarction was induced in the recipient by coronary ligation through a left lateral intercostal thoracotomy. (5) 12 hours later, the infarct was harvested and prepared by enzymatic digestion. (6) The resulting cell suspension was stained with an antibody cocktail including antibodies for CD45.1 and 2, which reported the source of recruited monocytes by flow cytometry. CD45.2+ cells originated from the donor spleen, whereas CD45.1+ cells were from the recipient animal.
Figure S10: Number of monocytes originating from a transplanted donor spleen and released into blood of the recipient.
Figure S10: Number of monocytes originating from a transplanted donor spleen and released into blood of the recipient.

The data show the number of total monocytes in 2 ml blood. Mice were analyzed in transplanted animals that were subjected to MI or not (n=2).
Figure S11: The spleen is not a site of monocyte production.
Figure S11: The spleen is not a site of monocyte production.

CD45.2+ donor spleens were implanted in the omentum of CD45.1+ mice and removed for analysis 1 day or 21 days after transplantation. Data show representative contour plots depicting the relative distribution of donor CD45.2+ and host CD45.1+ cells (n=3). *P < 0.05.
Figure S12 (A to G): Splenic monocytes and neutrophils have distinct reservoir capacities.
Figure S12 (A to G): Splenic monocytes and neutrophils have distinct reservoir capacities.

(A) Flow cytometric enumeration of neutrophils in single cell suspensions obtained from the entire spleen, and either in the absence of MI or 0.5 and 1 d after MI. The number of neutrophils is statistically unchanged after MI, which indicates that the spleen does not lose a significant population of neutrophils (n=3-6). (B, C) Immunofluorescence analysis of neutrophils (NIMP-R14+ cells) in the subcapsular red pulp in the absence of MI or 1 d after MI. Panel B shows a similar pattern of neutrophil residency, and Panel C shows similar neutrophil counts 1 day after MI when compared to controls. (D) Flow cytometric analysis of neutrophil counts in the blood in the absence of MI or 0.5 and 1 day after MI in mice with or without spleen. The results show no significant contribution of the spleen to the number of circulating neutrophils (n=6). (E) Flow cytometric analysis of neutrophils in the heart either in the absence of MI or 1 day after MI in mice with or without spleen. (F) Enumeration of neutrophils in the same mice shows a slight, but insignificant decrease of neutrophils in mice without spleen. n=3-6. (G) CD45.2+ spleens were transplanted immediately after MI to CD45.1+ splenectomized mice. Data show that 1 day after MI spleenderived neutrophils do not contribute to the neutrophil population in the heart. Data pool two experiments (A, D, F), or are from one experiment representative of two independent experiments (B, C, E, G).
Figure S13: Redistribution of monocyte subsets after MI.
Figure S13: Redistribution of monocyte subsets after MI.

Total number of Ly-6Chigh and Ly-6Clow monocytes lost from the spleen (left) and gained in the heart (right) 1 day after MI. Mean±SEM (n=6-15). Data pool at least three independent experiments.
Figure S14: Accumulation of monocytes in heart of Atgr1a-/- mice.
Figure S14: Accumulation of monocytes in heart of <em>Atgr1a<sup>-/-</sup></em> mice.

Control (WT) and Atgr1a-/- mice were subjected to MI. The number of monocytes per mg tissue was determined 1 day later (n=3). *P < 0.05.
Figure S15 (A to C): CD11b+ cells emigrate the subcapsular red pulp in response to Ang II.
Figure S15 (A to C): CD11b<sup>+</sup> cells emigrate the subcapsular red pulp in response to Ang II.

(A) Representative immunofluorescence sections of the spleen stained with anti-CD11b (red) and DAPI (blue) depict the subcapsular red pulp from control mice (left) and mice 1 day after Ang II osmotic pump infusion (right). (B) Enumeration of CD11b+ cells per high power field in the subcapsular red pulp of the mice mentioned above. Mean and SEM are shown, n=10 high power fields. (C) Intravital microscopy of GFP+ cells (green) in the spleen subcapsular red pulp of live Cx3cr1gfp/+ mice. Images show clusters of monocytes in control mice (left) and their disappearance 1 day after Ang II osmotic pump infusion (right). *P < 0.05. All data are representative of at least two independent experiments.
Figure S16 (A to E): Identification of GFP+ Mø/DC, patrolling monocytes, and splenic monocytes by intravital microscopy in Cx3cr1gfp/+ mice.
Figure S16 (A to E): Identification of GFP<sup>+</sup> Mø/DC, patrolling monocytes, and splenic monocytes by intravital microscopy in <em>Cx<sub>3</sub>cr1<sup>gfp/+</sup></em> mice.

(A) Representative immunohistology of a large F4/80high GFP+ Mø/DC (top) and a smaller F4/80 lo/- GFP+ monocyte (bottom). (B) GFP+ cells recorded in the subcasular red pulp were divided based on their size (Mø/DC: Area ≥ 150 µm2, ~5% of recorded GFP+ cells; monocytes: Area < 150 µm2, ~5% of GFP+ cells). (C) After using size to separate GFP+ cells into Mø/DC and monocytes, monocytes were further divided into patrolling monocytes (cells physically interacting with blood vessels, ~5% of GFP+ cells), and splenic monocytes (not physically interacting with vessels, ~90% of GFP+ cells). It is worth noting that splenic monocytes represent the vast majority of recorded GFP+ cells. Mø/DC were in minority because our recordings were performed in the subcapsular red pulp (marginal zones are typically located at depths greater than 100 µm). (D) Mø/DC show on average slower mean track velocities than splenic monocytes (Mø/DC: 0.20 ± 0.10 µm/min; splenic monocytes: 2.28 ± 2.95 µm/min; t test: P < 0.028). (E) Mø/DC show on average a lower circularity (or 'roundness') index than splenic monocytes (Mø/DC: 0.48 ± 0.12; splenic monocytes: 0.78 ± 0.09; t test: P < 0.0001). Circularity Index = 4πA/P2, where A is the cell cross-sectional area and P the cell perimeter). Data pool (B, D, E) or are from one experiment representative of three independent experiments (A, C).
Figure S17: Displacement over time for all parenchymal GFP+ cells in Cx3cr1gfp/+ mice.
Figure S17: Displacement over time for all parenchymal GFP<sup>+</sup> cells in <em>Cx<sub>3</sub>cr1<sup>gfp/+</sup></em> mice.

Data show displacement over time for all splenic GFP+ cells, i.e., both splenic monocytes and splenic Mø/DC, in resting mice (control) and 1 day after MI or infusion of Ang II. n=155 (control), 171 (MI), and 134 (Ang II) cells. This figure illustrates that the combination of so-called 'splenic monocytes' and 'splenic macrophages/DCs' (analyzed separately in Fig. 4I) preserves statistically different displacements over time between control and MI-treated or Ang II-treated groups. *P < 0.01, **P < 0.001. Data pool three independent experiments.
Figure S18 (A to B): Activation of splenic monocytes in vivo occurs without physical contraction of the subcasular red pulp.
Figure S18 (A to B): Activation of splenic monocytes in vivo occurs without physical contraction of the subcasular red pulp.

(A) Anesthetized Cx3cr1gfp/+ mice received a blood pool agent to visualize vessels (Angiosense-680, red) and were imaged by intravital micrcoscopy at relatively low magnification in monocyte-rich regions of the subcapsular red pulp. (B) Mice then received Ang II as described previously and the same region was imaged at different time points. Images on top show intravital recordings, whereas images on the bottom include the outline of vessels identified at time=0. The outline serves as a fiducial marker and is reproduced on the subsequent images. Physical contraction of the subcapsular red pulp should associate with decreased distances between the fiducial marker, a phenomenon that we did not observe during the duration of the experiment. Note leakage of the blood pool agent in the red pulp parenchyma over time, which is in line with previous findings that Ang II increases microvascular permeability.

Tables (expand all 4 tables)

Table T1: Comparison of blood and spleen Ly-6Chigh monocyte transcriptomes analyzed by Whole Mouse Genome Oligo Microarray.
AdjP < 0.05a FDR < 0.05b
Number of similarly expressed probe-complementary sequences 41173 41172
Number of differentially expressed probe-complementary sequences 1 2
% Identity 99.998 99.995

a AdjP < 0.05 refers to adjusted P value and controls for family-wise the probability of having more than one false discovery. An AdjP cutoff of 0.05 indcates that the declared differentially expressed set has 5% chance to have at least one false positive. b FDR < 0.05 refers to False Discovery Rate, which is the percentage of falsely declared differentially expressed genes among the set of declared differentially expressed genes. A FDR cutoff of 0.05 indicates that 5% of the declared differentially expressed genes are expected to be false positives.
Table T2: List of genes differentially expressed in blood and spleen Ly-6Chigh monocytes based on either the AdjP < 0.05 or FDR < 0.05 tests.
Test Gene symbol Gene description Accession number Fold difference
AdjP < 0.05a Lcn2 Mus musculus lipocalin 2 NM_008491 15.7
FDR < 0.05 Lcn2 Mus musculus lipocalin 2 NM_008491 15.7
FDR < 0.05b Slc40a1 Mus musculus solute carrier family 40 NM_016917 4.5

a AdjP < 0.05 refers to adjusted P value and controls for family-wise the probability of having more than one false discovery. A cutoff of 0.05 indicates that the declared differentially expressed set has 5% chance to have at least one false positive. b FDR < 0.05 refers to False Discovery Rate, which is the percentage of falsely declared differentially expressed genes among the set of declared DE genes. A FDR cutoff of 0.05 indicates that 5% of the declared differentially expressed genes are expected to be false positives.
Table T3: Fold difference expression between monocyte subsets analyzed by real-time PCR.
Gene symbol Gene description Accession number Fold difference (Blood/Spleen) Ly-6Chigh Fold difference (Blood/Spleen) Ly-6Clow Fold difference (Ly-6Chigh/Ly-6Clow) Blood Fold difference (Ly-6Chigh/Ly-6Clow) Spleen
Arg1 Arginase-1 NM_007482 nsa ns ns ns
Ccl2 Monocyte chemoattractant protein 1 NM_011333 2.8b ns 25.4 50.8
Ccr2 C-C chemokine receptor type 2 NM_009915 ns ns 13.9 13.3
Cd209a DC-SIGN1 NM_133238 ns ns ns 2.3
Cd274 B7-H1/Programmed death ligand 1 NM_021893 ns ns -9.3 -15.2
Cd276 B7-H3/Costimulatory molecule NM_133983 ns ns ns ns
Cd68 gp110 NM_009853 ns ns ns ns
Cd86 Activation B7-1 antigen NM_019388 ns ns ns ns
Csf1r Macrophage colony-stimulating factor 1 receptor NM_007779 ns ns ns ns
Ctsb Cathespin B1 NM_007798 ns ns ns ns
Cx3cl1 Chemokine (C-X3-C motif) ligand 1/ Fractalkine NM_009142 ns ns ns ns
Cx3cr1 CX3C chemokine receptor 1/ Fractalkine receptor NM_009987 ns ns nsc nsc
Egf Epidermal growth factor NM_010113 ns ns ns ns
Emr1 Cell surface glycoprotein F4/80 NM_010130 ns ns ns ns
F13a1 Coagulation factor XIIIa NM_028784 ns ns 229.6 770.2
Fgf2 Basic fibroblast growth factor NM_008006 ns ns ns ns
Ifng Interferon gamma NM_008337 ns ns ns ns
Il10 Interleukin 10 NM_010548 ns ns -44.9 -17.9
Il12a Interleukin 12a NM_008351 ns ns -50.4 -7.4
Il13 Interleukin 13 NM_008355 ns ns ns ns
Il18 Interleukin 18 NM_008360 ns ns ns 2.6
Il1b Interleukin 1 beta NM_008361 ns ns ns -3.7
Il1rap Interleukin 1 receptor accessory protein NM_008364 ns -2.0 ns ns
Il23 Interleukin 23 NM_031252 ns ns ns ns
Il4 Interleukin 4 NM_021283 ns ns -82.8 -6.5
Il4ra Interleukin 4 receptor, alpha NM_001008700 ns ns ns ns
Il6 Interleukin 6 NM_031168 ns ns -31.0 -15.5
Itgam Integrin alpha M/ CD11b antigen NM_008401 ns ns ns ns
Itgax Integrin alpha X/ CD11c NM_021334 ns ns -8.6 -11.1
Ly6c Lymphocyte antigen 6 complex, locus C NM_010741 ns ns 63.8 58.4
Mmp9 Matrix metallopeptidase 9 NM_0135991 ns ns ns ns
Mpo Myeloperoxidase NM_010824 ns ns 9.8 3.4
Mrc1 Mannose receptor, C type 1 NM_008625 ns ns 9.1 16.7
Mrc2 Mannose receptor, C type 2 NM_008626 ns ns -250.1 -827.4
Nos2 Inducible NO synthase NM_010827 ns ns ns ns
Pdcd1 Programmed cell death 1 NM_008798 ns ns ns ns
Plau Plasminogen activator, urokinase/ u-PA NM_008873 ns ns 48.9 50.2
Retnla Resistin like alpha/ Fizz1 NM_020509 ns ns ns ns
Sfpi1 PU.1 NM_011355 ns ns ns ns
Stat3 Signal transducer and activator of transcription 3 NM_213659 ns ns ns ns
Tek Tyrosine-protein kinase receptor TIE-2 NM_013690 ns ns ns ns
Tgfb1 Transforming growth factor, beta 1 NM_011577 ns ns ns ns
Tnf Tumor necrosis factor NM_013693 ns ns -4.6 -2.4
Tnfrsf1a Tumor necrosis factor receptor superfamily, member 1a NM_011609 ns ns ns ns
Vegfa Vascular endothelial growth factor A NM_001025250 ns ns 3.5 5.2

a ns refers to nondifferentially expressed genes measured in low density array analysis and as described in the Methods section. b Positive values indicate higher expression in the first group of pairs compared (i.e., blood compared to spleen, or Ly-6Chigh compared to Ly-6Clow), whereas negative values indicate the opposite. c Separate qPCR analysis identified ~2.5-fold higher CX3CR1 expression in Ly-6Clow cells both in blood and spleen (n=4).
Table T4: Role of CCR2 on monocyte efflux from spleen and influx to infarct 1 day after MI.
Wild-type Ccr2-/-
Efflux from spleen (cells x 105) Yes (2.9 ± 0.4)a Yes (3.0 ± 0.5)
Influx to infarct (cells x 105) Yes (4.4 ± 2.1)a No (0.1 ± 0.1)

a Mean ± SEM are shown.

Movies (expand all 10 movies)

Movie M1: Ring of GFP+ macrophages/DCs in the marginal zone of the spleen.

The animation shows GFP+ macrophages/DCs (green) at different depths throughout the spleen from Cx3cr1gfp/+ mice, starting at 50 µm and ending at 110 µm below the capsule. The cells are arranged in a ring in the marginal zone surrounding the white pulp. Scale bar=50 µm.
Movie M2: GFP+ monocytes in the subcapsular red pulp of the spleen.

The animation shows GFP+ monocytes (green) in the subcapsular red pulp of the spleen from Cx3cr1gfp/+ mice. The dense collagen network (blue) of the splenic capsule indicates the boundary of the organ. The recording covers 120 µm at 2 µm increments. Scale bar=100 µm.
Movie M3: GFP+ monocytes in the subcapsular red pulp of the spleen.

The animation shows GFP+ monocytes (green) in the subcapsular red pulp of the spleen from Cx3cr1gfp/+ mice, starting at 20 and ending at 50 µm below the capsule, respectively. Collagen fibers are shown in blue. Scale bar=25 µm.
Movie M4: A splenic monocyte in the subcapsular red pulp.

The movie shows a GFP+ splenic monocyte (green) in the subcapsularred pulp of a live Cx3cr1gfp/+ mouse. A fluorescent blood pool agent was injected immediately before imaging and is shown in red. Tracks indicate the position of the cell centroid at 15 second intervals. Time is shown in minutes and seconds. Scale bar=20 µm.
Movie M5: A splenic DC or macrophage in the subcapsular red pulp.

The movie shows a GFP+ DC or macrophage (green) in the subcapsular red pulp of a live Cx3cr1gfp/+ mouse. A fluorescent blood pool agent was injected immediately before imaging and is shown in red. Tracks indicate the position of the cell centroid at 15 second intervals. Time is shown in minutes and seconds. Scale bar=20 µm.
Movie M6: A patrolling monocyte in the red pulp of the spleen.

The movie shows a GFP+ monocyte (green) patrolling a blood vessel (red) in the subcapsular red pulp of a live Cx3cr1gfp/+ mouse. Blood vessels were visualized by injecting a fluorescent blood pool agent immediately before imaging. Tracks indicate the position of the cell centroid at 15 second intervals. Time is shown in minutes and seconds. Scale bar=20 µm.
Movie M7: Migratory activity of GFP+ cells in the subcapsular red pulp of the resting spleen.

The movie on the left shows GFP+ cells (green) in the subcapsular red pulp of a live Cx3cr1gfp/+ mouse. A fluorescent blood pool agent was injected immediately before imaging and is shown in red. Tracks indicate the position of the centroid of all GFP+ cells at 15 second intervals. Movies on the right separately show tracks (top) and cells (bottom). Three types of GFP+ cells can be visualized: i) motile monocytes patrolling the blood vessel are located in the upper left region of the recording; ii) a sessile cell (labeled with a purple dot at the center of the recording), that is larger in size and has several cytoplasmic protrusions/dendrites, and thus likely represents a macrophage or DC; iii) monocytes located in the spleen parenchyma and not associated with blood vessels. Time is shown in minutes and seconds. Scale bar=20 µm.
Movie M8: Migratory activity of GFP+ cells in the subcapsular spleen red pulp 2 h after MI.

The movie on the left shows GFP+ cells (green) in the subcapsular red pulp of a live Cx3cr1gfp/+ mouse. A fluorescent blood pool agent was injected immediately before imaging and is shown in red. Tracks indicate the position of the centroid of each GFP+ cell at 15 second intervals. Movies on the right separately show tracks (top) and cells (bottom). Time is shown in minutes and seconds. Scale bar=20 µm.
Movie M9: Migratory activity of GFP+ cells in the subcapsular spleen red pulp 2 h after infusion of Ang II.

The movie on the left shows GFP+ cells (green) in the subcapsular red pulp of a live Cx3cr1gfp/+ mouse. A fluorescent blood pool agent was injected immediately before imaging and is shown in red. Tracks indicate the position of the centroid of each GFP+ cell at 15 second intervals. Movies on the right separately show tracks (top) and cells (bottom). Time is shown in minutes and seconds. Scale bar=20 µm.
Movie M10: Departing splenic monocyte.

The movie shows a splenic GFP+ monocyte migrating from the spleen parenchyma to the lumen of a vessel. The cell likely had entered the vessel at time point 52:30 min:sec, and then migrated ~220 µm during the next 6 min. This represents an average instantaneous velocity of ~16 µm/min. This is in range with the average instantaneous velocities observed for patrolling monocytes. However, starting at time point 58:45 and during the next 30 sec, the cell migrated over > 125 µm and disappeared from the field of view. This represents an average instantaneous velocity of > 250 µm/min, which is one order of magnitude higher than the one reported for patrolling monocytes. Conversely, the cell did not enter free flow immediately after intravasation since it could be detected relatively close to the site of entry as late as 6 min after intravasation. Thus, departing monocytes may adhere for some time to the endothelium before they enter free flow. Tracks indicate the position of the cell centroid. Time is shown in minutes and seconds. Scale bar=20 µm.

1. Animals

C57BL/6J, B6.129P-Cx3cr1tm1Litt/J (Cx3cr1gfp/gfp), B6.129S4-Ccr2tm1Ifc/J (Ccr2–/–), B6.SJL-PtprcaPep3b/BoyJ (CD45.1+) and B6.129P2-Agtr1atm1Unc (At-1–/–) female mice (all from Jackson Laboratories) were used in this study. Cx3cr1gfp/+ mice were obtained by breeding Cx3cr1gfp/gfp mice with C57BL/6J mice. Cx3cr1gfp/+ mice have one Cx3cr1 allele replaced with cDNA encoding Egfp, and can be used to track monocytes (29). Mice were 8-12 weeks old, except Ccr2–/– which were 1 year old (spleens of young Ccr2–/– mice are very small). Female Wistar rats were ~230 g (from Jackson Laboratories).

2. Cells

Peripheral blood was drawn via cardiac puncture with citrate solution (100 mM Na-citrate, 130 mM glucose, pH 6.5) as anti- coagulant and mononuclear cells were purified by density centrifugation. Total blood leukocyte numbers were determined using acetic acid lysis solution (3% HEMA 3 Solution II, 94% ddH2O, 3% glacial acetic acid). After organ harvest, single cell suspensions were obtained from brain, gut, heart, kidney, liver, lung, muscle and pancreas by digestion with a cocktail of 450 U/ml collagenase I, 125 U/ml collagenase XI, 60 U/ml DNase I and 60 U/ml hyaluronidase (Sigma-Aldrich, St. Louis, MO) for 1 h at 37°C while shaking. Some spleens were also prepared with the digestion cocktail. Total viable cell numbers were determined using Trypan Blue (Cellgro, Mediatech, Inc, VA).

3. Flow Cytometry

Anti-CD90-PE, anti-CD90-FITC, 53-2.1 (BD Biosciences); anti-B220-PE, anti B220-FITC, RA3-6B2 (BD Biosciences); anti-CD49b-PE, anti CD49b-FITC, DX5 (BD Biosciences); anti-NK1.1-PE, anti-NK1.1-FITC, PK136 (BD Biosciences); anti-Ly-6G-PE, anti-Ly-6G-FITC, 1A8 (BD Biosciences); anti-CD11b-APC, M1/70 (BD Biosciences); anti-CD11b-PE (ED8) (Abcam); anti-CD11b-APC-Cy7 M1/70 (BD Biosciences); anti-F4/80- biotin, anti-F4/80-FITC, C1:A3-1 (BioLegend); anti-CD11c-biotin, anti-CD11c-FITC, anti-CD11c-APC, HL3 (BD Biosciences); anti-I-Ab- biotin, anti-I-Ab-FITC, AF6-120.1 (BD Biosciences); anti-Ly-6C-FITC, anti-Ly-6C-biotin, AL-21 (BD Biosciences); anti-CD43-FITC, S7 (BD Biosciences); anti-CD62L-FITC, MEL-14 (BD Biosciences); anti-CD68- FITC, FA-11 (AbD Serotec); anti-CD86-biotin, GL1 (BD Biosciences); anti-CD115-PE, 604B5-2E11 (AbD Serotec); anti-Mac-3-FITC, M3/84 (BD Biosciences); anti-Gr-1-PeCy7, RB6-8C5 (BD Biosciences); anti- CD45.2-FITC 104 (BD Biosciences); anti-CD45.1-biotin A20 (BD Biosciences); anti-CD45.1-APC A20 (BD Biosciences) were used for flow cytometric analyses in this study. Strep-PerCP (BD Biosciences) was used to label biotinylated antibodies. Monocytes were identified as CD11bhigh (CD90/B220/CD49b/NK1.1/Ly-6G)low (F4/80/I-Ab/CD11c)low Ly-6Chigh/low. Macrophages/DCs were identified as CD11bhigh (CD90/ B220/CD49b/NK1.1/Ly-6G)low (F4/80/I-Ab/CD11c)high Ly-6Clow or on the basis of F4/80 or CD11c expression only. Neutrophils were identified as CD11bhigh (CD90/B220/CD49b/NK1.1/Ly-6G)high (F4/80/I-Ab/ CD11c)low Ly-6Cint. Monocyte and macrophage/DC numbers were calculated as total cells multiplied by percent cells within the monocyte/ macrophage gate. Within this population, monocyte subsets were identified as (F4/80/I-Ab/CD11c)low and either Ly-6Chigh or Ly-6Clow. For calculation of total cell numbers in tissue, normalization to weight of tissue was performed. Data were acquired on an LSRII (BD Biosciences) and analyzed with FlowJo v.8.5.2 (Tree Star, Inc.). Cells were sorted on a BD FACSAria (BD Biosciences). For morphologic characterizations, sorted cells were prepared on slides by cytocentrifugation (Shandon, Inc.) at 10 g for 2 min, and stained with HEMA-3 (Fischer Scientific). For gene profiling studies, blood and spleens of 5 to 10 mice were pooled for each replicate. Splenic monocytes were enriched by lineage depletion using MACS LD columns (Miltenyi) and PE–conjugated antibodies against B220, CD49b, NK1.1, Ly-6G, CD90 and Ter-119 followed by anti-PE magnetic beads (Miltenyi). Lineage-depleted cells were further stained with specific antibodies to allow for phenotypic identification of monocyte subsets. Monocytes from blood were stained and sorted without prior enrichment.

4. Microarray gene expression profiling

Monocyte subsets from blood and spleen of a group of four mice were isolated by fluorescence activated cell sorting (FACS) as CD11bhigh (CD90/B220/CD49b/NK1.1/ Ly-6G)low (F4/80/I-Ab/CD11c)low Ly-6Chigh cells. To avoid effects of lengthy staining protocols on the transcriptome of the cells of interest we developed a protocol that allowed sorting of cell subsets into RNA lysis buffer within ~30 min after the animals were sacrificed. Briefly, heparin- blood was drawn from anesthetized mice by cardiac puncture and spleens were immediately homogenized through a nylon mesh into 3 ml of PBS. The volume of the heparinized blood was adjusted with PBS to 1 ml of blood and the antibody mix was added. Similarly the antibodies were added to 1 ml of the splenocyte suspension. Staining was performed for 10 min, samples were diluted by addition of 1 ml PBS, immediately loaded onto 2 ml histopaque, and spun for 10 min at 18 g, 22°C. The interphase was collected and diluted into one volume of sorting buffer containing PBS, 2% FCS and 2 mM EDTA. Cells were FACS-sorted without delay. The protocol neither imposed cell pelleting, extend dwelling on ice, nor induced major osmotic stress. Both preparations of blood and splenic monocytes were performed for each animal simultaneously and under the same conditions. Samples of 1,000 Ly-6Chigh blood and Ly-6Chigh splenic monocytes were collected directly into 20 µl lysis buffer of the PicoPure RNA isolation kit (Arcturus). Sorting times varied between 2 and 10 min. RNA extraction was subsequently performed according to the manufacturer’s instructions (Arcturus). RNA quality was assessed using RNA pico lab chips on the Agilent Bioanalyzer. For all samples a RIN above 8 could be achieved. On average 1,000 cells yielded 200 pg total RNA. All further steps were performed at the UCSF Shared Microarray Core Facilities according to standard protocols (http://www.arrays.ucsf.edu and http://www.agilent.com). RNA was amplified using the NuGen WT-Ovation Pico System, and the amplified cDNA was labeled using the FL-Ovation cDNA Fluorescent Module (NuGen Technologies, San Carlos, Ca). Briefly, input total RNA was reverse–transcribed into cDNA and then amplified using a linear isothermal amplification process (SPIA). The amplified products were CY-3 labeled and fragmented according to manufacturer’s guidelines. Labeled cDNA was assessed using the Nanodrop ND-100 (Nanodrop Technologies, Inc., Wilmington DE) and the Agilent 2100 Bioanalyzer; equal amounts of Cy3-labeled target were hybridized to Agilent whole mouse genome 4x44K Ink-jet arrays. Hybridizations were performed for 14 h according to the manufacturers protocol. Arrays were scanned using the Agilent microarray scanner and raw signal intensities were extracted with their Feature Extraction v9.1 software. The data set was normalized using the quantile normalization method (30). No background subtraction was performed, and the median feature pixel intensity was used as the raw signal before normalization. The moderated t-statistic and false discovery rate for each gene of comparison between blood and spleen were calculated. Adjusted p-values were produced according to the Holm-Bonferoni method (31). All procedures were carried out using functions in the LIMMA software package of the Bioconductor Project (www.bioconductor.org). MIAME compliant expression data have been deposited under the accession GSE14850.

5. Real-time PCR reactions of preselected genes

For phenotypic differentiation of monocyte subsets by expression analysis we designed a TaqMan custom low–density array (Applied Biosystems) comprising 45 genes of interest and three endogenous control genes for quality controls purposes (Table T3). The technical details of the procedure can be found here: http://www3.appliedbiosystems.com/cms/groups/mcb_marketing/documents/generaldocuments/cms_040595.pdf. RNA was extracted from FACS-sorted monocyte subsets using the RNAeasy mini Kit (Qiagen). Typically, 250,000 monocytes of each subset were obtained from pooled blood and spleen samples of 10-15 mice. 250,000 cells yielded 100-250 1ng RNA in a volume of 35 µl. RNA yield and integrity were assessed with a NanoDrop spectralphotometer (Thermo-Scientific) and an Agilent Bioanalyzer (Agilent) using the eukaryotic RNApico lab chip. Only samples with a RNA integrity number of above 7 were used for further processing. For low density array profiling, real time PCR cDNA was generated from 50 ng or 100 ng of RNA per sample by reverse transcription (RT) using multiplex RT pools (Applied Biosystems) according to the manufacturer’s protocol. The cDNA was then applied to the micro-fluidic card and real time PCR was performed on a 7900HT real time PCR machine (Applied Biosystems). Cycle threshold (Ct) values (auto thresholding) of the real time PCR readouts were compared among subsets derived from blood and spleen. Four independent replicates of each subset and from each organ were used for analysis. We used the Global Pattern Recognition (GPR) and geNorm algorithms to compare gene expression between groups. GPR detects significant changes in gene expression by multiple gene normalization, which does not require or assume constant level of expression of a single normalizer gene (i.e., 'housekeeping gene'). By comparing the expression of each gene to every other gene in the array, a global pattern is established, and significant changes are identified and ranked (32). The geNorm algorithm implemented in the GPR program was used to calculate fold changes of gene expression based on the geometrical mean of a group of 10 best normalizers identified by GPR. Taken together, gene expression could be compared by two means: (i) ranking scores determining the significance of a difference in gene expression among the groups compared and (ii) by a reliable determination method of fold-changes in gene expression. Based on these readouts a gene was considered to be differentially expressed when it had been scored by GPR and showed a fold change >2.

6. In vitro phagocytosis

FACS-sorted monocytes from blood and spleen were incubated for 4 h with latex beads at a 1/10 cell/bead ratio (yellow- green latex beads, 2.5 µm, Sigma) in 200 µl RPMI 1640 (Mediatech, Inc, VA) in a 96-well plate (100K/well) (Costar, Corning Inc, NY).

7. In vitro differentiation

FACS-sorted Ly-6Chigh and Ly-6Clow monocytes from blood and spleen were treated either with M-CSF (0.02 µg/ml) or GM-CSF (0.5 µg/ml) and IL-4 (0.2 µg/ml) in RPMI 1640 medium (Cellgro, Mediatech, Inc, VA) supplemented with 10% FCS (Valley Biomedical, Inc.), 50 µM 2-Mercaptoethanol (Cellgro, Mediatech, Inc, VA) and 100 U/ml Penicillin-Streptomycin (Cellgro, Mediatech, Inc, VA). Cells (105) were plated in triplicate in 96-well round-bottom plates (Costar, Corning Inc, NY) and cultured in a humidified incubator at 37°C, 5% CO2. The medium was replaced every second day to keep the growth factors fresh. Cells were harvested at day 7, and expression of the cell surface markers F4/80 and CD11c was determined by staining for 30 min with anti-F4/80-biotin, C1:A3-1 (AbD Serotec) and anti-CD11c-APC, HL3 (BD Biosciences); Strep-PerCP (BD Biosciences) labeled the biotinylated antibody.

8. Histology

Histology of spleens was assessed for the following groups: wild type C57BL/6 mice, wild type mice 1 day after MI, wild type mice 1 day after Ang II, Cx3cr1gfp/+ mice, Cx3cr1gfp/+ mice 1 day after MI, and Cx3cr1gfp/+ mice 1 day after Ang II. Spleens were excised, rinsed in PBS and embedded in OCT (Sakura Finetek). Fresh-frozen serial 6 µm thick sections were used for overall histological analysis and immunofluorescence staining. Hematoxylin and eosin staining was used to identify red and white pulps. Sections were incubated with anti-CD11b, M1/70 (BD Pharmingen); anti-F4/80, A3-1 (Abcam); anti-CD11c, 3.9 (Abcam); anti-neutrophil, NIMP-R14 (Santa Cruz Biotechnology, Inc); anti-CD163, G-17 (Santa Cruz Biotechnology, Inc); or anti-CD49b, Hal/29 (BD Pharmingen) antibodies followed by an appropriate biotinylated secondary antibody, and texas red-conjugated streptavidin (GE Healthcare). DAPI (Vector Laboratories) was used to identify cell nuclei. Negative controls were obtained by incubating tissue sections with the corresponding secondary antibodies only. Cell numbers were quantified using IPLab (version 3.9.3; Scanalytics, Inc., Fairfax, VA) and signal intensities were calculated using ImageJ (version 1.38x).

9. Parabiosis

Surgical gloves and autoclaved sterilized instruments were used. Animals were kept warm with a heating pad. Mice were weight- matched. We administered analgesia (buprenorphine 0.05-0.2 mg/kg) 30 minutes before surgery. Mice were anesthetized with isoflurane (2%/2L) and joined by a technique adapted from Bunster and Meyer (33). After shaving the corresponding lateral aspects of each mouse, matching skin incisions were made from behind the ear to the tail of each mouse, and the subcutaneous fascia was bluntly dissected to create about ½ cm of free skin. The olecranon and knee joints were attached by a mono-nylon 5.0 (Ethicon, Albuquerque, NM), and the dorsal and ventral skins were approximated by continuous suture. In some experiments, after an interval of several weeks, parabiosed mice were surgically separated by a reversal of the procedure. Percent chimerism was defined for gated monocytes as %CD45.1 / (%CD45.1 + %CD45.2) in CD45.2 mice, and as %CD45.2 / (%CD45.2 + %CD45.1) in CD45.1 mice.

10. Myocardial infarction

Mice or rats were anesthetized with gas anesthesia (isoflurane 2% / 2L O2), and intubated and ventilated with an Inspira Advanced Safety Single Animal Pressure/Volume Controlled Ventilator (Harvard Apparatus, Holliston, MA). The chest wall was shaved and left thoracotomy was performed in the 4th left intercostal space. The left ventricle was visualized and the left coronary artery was permanently ligated with monofilament nylon 8-0 sutures (Ethicon, Somerville, NJ) at the site of its emergence from under the left atrium. The chest wall was closed with 7-0 nylon sutures and the skin was sealed with superglue. Notably, for mice used in this study, the infarcts were of small to moderate size (~15% in delayed enhancement MRI) and therefore did not alter blood pressure or cardiac output. We measured cardiac index in infarcted and non-infarcted mice using gated high field cardiac MRI volumetry as described previously (34). Cardiac index was not changed on day 1 after coronary ligation: MI 780 ± 53 ml/min*kg, no MI 792 ± 94 ml/min*kg, n=6 per group.

11. Splenectomy

During isoflurane anesthesia, the abdominal cavity of mice was opened and the spleen vessels were cauterized.The spleen was carefully removed and placed in cold PBS solution. For control experiments, the abdomen was opened, but the spleen was not removed. In rats, splenic vessels were ligated with 7-0 sutures.

12. Spleen transplant

A pictorial representation of the procedure is shown in Fig. S7. Spleen donor mice (CD45.2) were anesthetized with a subcutaneous injection of ketamine (90 mg/kg) and xylazine (10 mg/kg), followed by an intravenous injection of 200 units of heparin (American Pharmaceutical, Schaumburg, Il). The complete inhibition of clotting ensures that no vascular or intrasplenic thrombosis occurs. In deep anesthesia, the thorax was then opened and the right atrium incised to allow blood to exit during perfusion. Over a period of 5 minutes, the entire mouse was then perfused with a total of 20 ml of normal saline through a cannula inserted into the apex of the left ventricle. At the end of this procedure, fluid exiting the right atrium was clear which indicated thorough removal of the donor blood. The abdomen of the donor was then opened with a longitudinal incision. The pancreas, the spleen and the abdominal vasculature in the epigastric region were visualized. Small vessels between the pancreas and the intestine were ligated with 6.0 cotton (Ethicon). The celiac artery was then isolated, and the hepatic and gastric artery ligated with 10.0 suture (Ethicon). The abdominal aorta was ligated and cut just below the celiac artery with micro-dissection scissors (ROBOZ, Rockville, MD), and dissected above the celiac artery. This approach resulted in an aortic cuff connected to the splenic artery, which allowed vascular anastomosis of the spleen to the recipient. Following ligation of the bile duct, the portal vein was isolated, and the superior and inferior mesenteric and gastric veins were ligated. The portal vein was intersected closely to the liver. The entire organ package containing the vascular connections, spleen and the pancreas was then removed and stored in ice cold saline for 15 minutes while the recipient was prepared. The recipient (CD45.1) was anesthetized with isoflurane supplemented with oxygen (2-3 Vol%). An abdominal midline incision was made and the inferior vena cava and the descending aorta were isolated below the renal arteries. The recipient vessels were clamped with an atraumatic bulldog clamp (ASSI, Westbury, NY) and opened with micro-scissors. The portal vein was anastomosed to the inferior vena cava and the donor aortic cuff was connected with an end-to side anastomosis to the recipient aorta using 10.0 suture. The clamp was then removed to restore blood flow. The time of ischemia of the donor spleen, which ended after completion of both vascular anastomoses by unclamping the recipient aorta and vein, was ~60 min. Flow cytometric analysis of transplanted spleens (in mice without MI) indicated that the procedure reduced on average the reservoir of donor splenic monocytes by ~6-fold (donor monocytes in donor spleens 12 h after operation: 0.23 ± 0.01 X 106 cells, n=2; control monocytes in control spleens: 1.4 ± 0.2 X 106 cells, n=8). The ‘missing’ monocytes in transplanted spleens likely matured into Mø/ DC because the number of these cells increased locally (donor CD45.2+ CD11b+ Mø/DC in donor spleens: 0.99 ± 0.6 X 106 cells; control CD11b+ Mø/DC in control spleens: 0.36 ± 0.5 X 106 cells). Some monocytes may also have died locally, however they virtually did not enter circulation (donor monocytes in blood: 0.0012 ± 0.0004 X 106 cells). The reduced availability of donor monocytes in transplantation experiments was taken into account when quantifying their relative contribution in infarcts (see below). In an additional cohort of mice, 1 h after this procedure, the mouse was re-anesthetized and myocardial infarction was induced as described above (n=2). 24 h later, flow cytometry analysis of cells from MI revealed 10.1 ± 2% monocytes of splenic origin. Taking into account the reduced reservoir of splenic monocytes in transplanted animals, we calculated that a normal spleen should contribute ~40% of the recruited monocytes (6 X 10.1% splenic monocytes versus 89.9% other monocytes ˜40% splenic monocytes versus 60% other monocytes). We also performed the two following control experiments: The first experiment involved mice (n=2) transplanted with a spleen as mentioned above, but the spleens were excised just before unclamping the host vasculature, leaving only the donor vasculature, connective tissue and parts of the pancreas in the recipient mouse. The mice were then subjected to MI and the infarcts investigated 24 h later by flow cytometry. We detected no CD45.2+ cells in these mice. The second experiment involved mice (n=2) that received a spleen graft but were not subjected to MI. In some experiments (Fig. S11), donor spleens were transplanted into the great omentum of recipients. Specifically, awhile submerged in ice-cold PBS solution, donor spleens were sectioned into 5 pieces of approximately 3 mm thickness, transplanted, and fixed in place with 8-0 nylon sutures. To facilitate microvascular connections, the hilar aspect of the splenic segments was brought in proximity to the great omental vessels. The institutional Subcommittee on Research Animal Care at Massachusetts General Hospital (MGH) approved all animal studies.

13. Fluorescence Molecular Tomography (FMT) - Magnetic Resonance Imaging (MRI)

Animal preparation: Mice were shaved and depilated to remove all hair that otherwise absorb light and interfere with optical imaging. Mice were anesthetized (Isoflurane 1.5%, O2 2 L/min) and rendered immobile in a multimodal imaging cartridge, which lightly compresses the anesthetized mouse between optically translucent windows. The latter do not interfere with fluorescence imaging. The cartridge provides fiducial landmarks on the frame that enable exact, robust and observer-independent fusion of images. The cartridge also prevents motion of the mouse during transfer between modalities. CLIO-680 and ProSense-750 (VisEn Medical) were injected intravenously on day 1 after coronary ligation at a dose of 15 mg iron oxide/kg body weight and 5 nmol respectively, and FMT-MRI was carried out 24 hours later. Twelve mice with MI were imaged; 6 received splenectomy at the time of coronary ligation. A commercially available FMT system was used for in vivo imaging studies (FMT2500, VisEn Medical). The system allows for dual-channel imaging without the need for an index matching fluid, i.e., is a noninvasive free-space imaging system. It is equipped with MRI-safe mouse holders, allowing for imaging in the FMT and MRI systems without the need to reposition the mouse. Typical FMT scan times were less than 5 min per mouse. Data were post-processed using a normalized Born forward equation to calculate a 3D map of fluorochrome concentration (35). To reliably identify the region of interest within the heart, FMT was hybridized with MRI for anatomic reference. After completion of FMT, the imaging cartridge holding the anesthetized mouse was inserted into a custom- made plexiglass holder that supplies isoflurane anesthesia and optimal positioning inside a transmit-receive MR body coil. We used a 7 Tesla horizontal bore scanner (Pharmascan, Bruker, Billerica) and a RARE sequence (TE 26.9 ms, TR 2500 ms, slice thickness 1 mm, 24 slices, matrix 256x256, FOV 6x6 cm) to image the entire mouse and the fiducial markers on the imaging cartridge. The fiducial wells were filled with fluorochrome solution, and therefore were readily identified by FMT (fluorescence) and MRI (proton signal). FMT and MRI DICOM images were fused with OsiriX (The Osirix Foundation, Geneva, Switzerland). Within both data sets, fiducial points were tagged to define their X-Y-Z-coordinates. Using these coordinates, FMT data were then resampled, rotated and translated to match the MR image matrix, and finally fused. After identification of infarcts on MRI, regions of interest were defined in both FMT channels. FMT-derived fluorescence is given in pmol and represents the absolute quantity of the excited fluorochrome within the infarct.

14. Fluorescence reflectance imaging (FRI)

Myocardial short-axis sections were produced after harvest of rat hearts and then exposed on a custom- built fluorescence reflectance system 24 h after i.v. injection of 5 mg/kg CLIO-VT680.

15. Delivery of Angiotensin (Ang) II

C57BL/6 mice were implanted subcutaneously in the dorsum of the neck with osmotic mini-pumps (Alzet) for 2 to 24 hours. Mini-pumps were pre-incubated in PBS for 4 h to assure immediate delivery of the agent after implantation. Ang II (Bachem) was delivered at a rate of 1 µl/h at 1.5 mg/kg/day. Control mice were implanted with mini-pumps delivering saline (0.9% NaCl) (n=3-6).

16. ELISA for determination of serum Ang II levels

Blood was obtained from mice under anesthesia by cardiac puncture with a syringe pre-loaded with 80 µl of 100 mM EDTA anticoagulant. The blood was supplemented with 50mM p-hydroxymercuribenzoid acid (Sigma), centrifuged, and supernatant loaded onto Amprep Phenyl PH mini-columns (GE Biosciences) to isolate peptides from the sera. Methanol- eluted peptides were dried by vacuum centrifugation. Ang II concentration was determined by using an Ang II ELISA (Cayman Chemical), and normalized to the volume of blood isolated.

17. Western for Ang II Type 1 (AT-1) receptor on splenic monocytes

Monocytes were isolated by FACS as described above. Sample pellets of ~175,000 monocytes were resuspended in Laemmli buffer (BioRad), and sonicated to lyse cells and shear genomic DNA. Samples were developed by electrophoresis on a 4-15% polyacrilamide gel (BioRad). The proteins were transferred to a polyvinylidene difluoride membrane (Fischer) by semidry transfer. Membranes were blocked with carnation milk and PBS supplemented with 0.05% Tween 20 overnight. Membranes were washed, stained initially with anti-AT-1 receptor antibody (Abcam), stripped with Restore buffer (Pierce), and stained with anti- glyceraldehyde-3-phosphate (GAPDH) (Rockland Immunochemicals for Research). Blots were developed with Western Lightning Chemiluminescence reagent (PerkinElmer Instruments) and molecular weights were compared to bands for Precision Plus Protein Western C standards (BioRad).

18. In vitro migration

Migration experiments using Ang II as a chemoattractant were performed in BD BioCoat invasion chambers (BD Biosciences). Sorted monocytes from spleen were suspended in RPMI 1640 media (Cellgro) supplemented with 0.2% FCS (Valley Biomedical, Inc.). 2x105 cells were placed on the matrigel-coated 8 µm pore size PET membrane and incubated in a humidified incubator at 37°C, 5% CO2 for 1 h, allowing the cells to attach to the matrigel. Migration was induced by addition of Angiotensin II (1 µM) (Bachem) to the lower compartment. After 2 h, non-migrating cells were removed with a cotton tip and the membranes were fixed and stained with HEMA 3 staining set (Fisher Scientific) to identify cells that had migrated to the lower surface of the membrane. The number of migrated cells was determined per 200 × high-power field. Cells that had migrated to the lower chamber were counted using Trypan Blue (Cellgro).

19. Intravital imaging

Animal preparation: During isoflurane anesthesia, the peritoneal cavity was opened with a transverse incision in the disinfected abdominal wall. The gastric-splenic ligament was dissected and the spleen carefully exteriorized. Robust blood flow was observed in the splenic artery during the duration of each experiment and splenic perfusion was confirmed by inspection through fluorescence microscopy upon tail vein injection of an intravascular imaging agent. The exteriorized spleen was completely submerged in temperature-controlled saline solution. Temperature near the spleen was carefully monitored using an Omega HH12A thermometer with fine wire thermocouples (Omega Engineering Inc., Stamford, CT) and kept at 37ºC. Confocal Microscopy: Images were collected with a prototypical intravital laser scanning microscope (IV100, Olympus Corporation, Tokyo, Japan) (36) using an Olympus 20x UplanFL (NA. 0.5) objective and the Olympus FluoView FV300 version 4.3 program. Samples were excited at 488 nm with an air-cooled argon laser (Melles Griot, Carlsbad, CA) for visualization of the GFP+ cells, and at 748 nm with a red diode laser (Model FV10-LD748, Olympus Corporation, Tokyo, Japan) for visualization of the blood pool agent (AngioSense-680, VisEn Medical, MA). Light was collected using custom-built dichroic mirrors SDM-570 and SDM-750, and emission filters BA 505-550 and BA 770 nm IF (Olympus Corporation, Tokyo, Japan). Both channels were collected simultaneously. A prototypical tissue stabilizer (Olympus Corporation, Tokyo, Japan) was used to reduce motion and stabilize the focal plane. The stabilizer was attached to the objective and its z-position was fine adjusted using a micrometer screw to apply soft pressure on the tissue. Time-lapse recordings were made by collecting images of 256x256 pixels at 15 s intervals over 1 h in a single after either MI or infusion of Ang II. Multiphoton Microscopy: Mice were anesthetized with ketamine (150 mg/kg BW) and xylazine (10 mg/kg BW) i.m., and the spleen was immobilized by placing a coverslip on its ventral surface. Images were collected with Praireview software on an Ultima IV upright multiphoton microscope (Prairie Technologies, Middleton, WI) equipped with an Olympus 20x/0.95 NA water immersion objective. For multiphoton excitation and second harmonic generation, a Ti:sapphire laser with 10-W MilleniaXs pump (Mai Tai HP, Spectra-Physics, Mountain View, CA) was tuned to 920 nm. Emitted light and second harmonic signals were detected through 525/50 and 460/50 nm bandpass filters using non-descanned detectors to generate two-color images and stacks, which were volume-rendered using the brightest-spot rendering mode within Volocity software (Improvision, Coventry, UK). Optical slides were acquired at 1 or 2 µm intervals. The number of stacks varied between 16 and 60 (please refer to Movies S1-3 for more information). Data Analysis: All GFP+ cells were identified manually in each recording. To determine the displacement over time of individual cells, the centroid position (x-y dimension) of these cells was recorded at the first and last time-point when they could be identified during a recording; then the distance between these two points was calculated, and divided by the elapsed time. Single cell tracks for GFP+ cells were generated based on the position of cell centroids from a series of images recorded at 15 s intervals, and ImageJ and the Manual Tracking plugin (http://rsbweb.nih.gov/ij/plugins/track/track.html) were used for display and quantification. Motion-artifacts in recordings were corrected using the auto-alignment plugin (stackreg) of ImageJ (http://rsb.info.nih.gov/ij/).

20. Statistics

Results were expressed as mean ± SEM. Statistical tests included unpaired, 2-tailed Student's t test using Welch's correction for unequal variances and 1-way ANOVA followed by Bonferroni’s multiple comparison test. P values of 0.05 or less were considered to denote significance.

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Acknowledgements and Declarations

This work was supported in part by NIH grants U01 HL080731, P50 CA86355, R24 CA69246, U54 CA126515, and P01 A154904 (to R.W.), MGH-Center for Systems Biology (to M.J.P.), AHA SDG 0835623D (to M.N.), and NIH grant 1R01HL095612 (to F.K.S). The authors thank U. von Andrian for critical assessment of the manuscript; A. Luster for providing Ccr2–/– mice; D. Erle, A. Barcak, and C. Eisley for microarray hybridization and data analysis; M. Waring for sorting cells; A. Moseman for helpful discussion and feedback with parabiosis experiments; and A. Newton, C. Siegel, N. Sergeyev, and Y. Iwamoto for technical assistance. MIAME (minimum information about a microarray experiment)–compliant expression data have been deposited under accession no. GSE14850 [NCBI GEO] . This work is dedicated to the memory of Marc-Henri Pittet.