Identification of Splenic Reservoir Monocytes and Their Deployment to Inflammatory Sites
Science31 July 2009Vol. 325 no. 5940 pp. 612-616
DOI: 10.1126/science.1175202
, Matthias Nahrendorf,1 Martin Etzrodt,1, 2 Moritz Wildgruber,1 Virna Cortez-Retamozo,1 Peter Panizzi,1 Jose-Luiz Figueiredo,1 Rainer H. Kohler,1 Aleksey Chudnovskiy,1 Peter Waterman,1 Elena Aikawa,1 Thorsten R. Mempel,1, 3 Peter Libby,4, 5 Ralph Weissleder,1, 6, Mikael J. Pittet,1
- 1 - Center for Systems Biology, Massachusetts General Hospital and Harvard Medical School, Boston, MA 02114, USA.
- 2 - Faculty of Biology and Medicine, University of Lausanne, CH-1015 Lausanne, Switzerland.
- 3 - Center for Immunology & Inflammatory Diseases, Massachusetts General Hospital & Harvard Medical School, Charlestown, MA 02129, USA.
- 4 - Cardiovascular Division, Department of Medicine, Brigham and Women’s Hospital, Boston, MA 02115, USA.
- 5 - Center for Excellence in Vascular Biology, Brigham and Women’s Hospital, Boston, MA 02115, USA.
- 6 - Department of Systems Biology, Harvard Medical School, Boston, MA 02115.
Overview
Abstract
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.
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
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
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
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
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.
Recent Publications
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Received for publication 20 April 2009. Accepted for publication 10 June 2009.
Full Text
Abstract
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.
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 (11–13). 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)
Supplementary Figures (expand all 18 supplementary figures)
Tables (expand all 4 tables)
| 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 |
| 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 |
| 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 |
Movies (expand all 10 movies)
Materials and Methods
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/).
References and Notes
| Ref. | Author / Title | Source | Year |
|---|---|---|---|
| 1 |
Luster, A. D., R. Alon, U. H. von Andrian
Immune cell migration in inflammation: present and future therapeutic targets
|
Nat. Immunol. 6, 1182 |
2005 |
| 2 |
Ley, K., C. Laudanna, M. I. Cybulsky, S. Nourshargh
Immune cell migration in inflammation: present and future therapeutic targets
|
Nat. Rev. Immunol. 7, 678 |
2007 |
| 3 |
Singer, A. J., R. A. Clark
Cutaneous Wound Healing
|
N. Engl. J. Med. 341, 738 |
1999 |
| 4 |
Auffray, C., M. H. Sieweke, F. Geissmann
Blood Monocytes: Development, Heterogeneity, and Relationship with Dendritic Cells
|
Annu. Rev. Immunol. 27, 669 |
2009 |
| 5 |
Gordon, S., P. R. Taylor
Monocyte and macrophage heterogeneity
|
Nat. Rev. Immunol. 5, 953 |
2005 |
| 6 |
Arnold, L., A. Henry, F. Poron, Y. Baba-Amer, N. van Rooijen, A. Plonquet, R. K. Gherardi, B. Chazaud
Inflammatory monocytes recruited after skeletal muscle injury switch into antiinflammatory macrophages to support myogenesis
|
J. Exp. Med. 204, 1057 |
2007 |
| 7 |
Nahrendorf, M., F. K. Swirski, E. Aikawa, L. Stangenberg, T. Wurdinger, J. Figueiredo, P. Libby, R. Weissleder, M. Pittet
The healing myocardium sequentially mobilizes two monocyte subsets with divergent and complementary functions
|
J. Exp. Med. 204, 3037 |
2007 |
| 8 |
Dunay, I. R., R. A. DaMatta, B. Fux, R. Presti, S. Greco, M. Colonna, L. D. Sibley
Gr1+ Inflammatory Monocytes Are Required for Mucosal Resistance to the Pathogen Toxoplasma gondii
|
Immunity 29, 306 |
2008 |
| 9 |
Serbina, N. V., E. G. Pamer
Monocyte emigration from bone marrow during bacterial infection requires signals mediated by chemokine receptor CCR2
|
Nat. Immunol. 7, 311 |
2006 |
| 10 |
Sunderkštter, C., T. Nikolic, M. J. Dillon, N. van Rooijen, M. Stehling, D. A. Drevets, P. J. M. Leenen
Subpopulations of Mouse Blood Monocytes Differ in Maturation Stage and Inflammatory Response
|
J. Immunol. 172, 4410 |
2004 |
| 11 |
Swirski, F. K., P. Libby, E. Aikawa, P. Alcaide, F. W. Luscinskas, R. Weissleder, M. J. Pittet
Ly-6Chi monocytes dominate hypercholesterolemia-associated monocytosis and give rise to macrophages in atheromata
|
J. Clin. Invest. 117, 195 |
2007 |
| 12 |
F. Tacke et al.
Monocyte subsets differentially employ CCR2, CCR5, and CX3CR1 to accumulate within atherosclerotic plaques
|
J. Clin. Invest. 117, 185 |
2007 |
| 13 |
Mantovani, A., P. Allavena, A. Sica, F. Balkwill
Cancer-related inflammation
|
Nature 454, 436 |
2008 |
| 14 |
Auffray, C., D. Fogg, M. Garfa, G. Elain, O. Join-Lambert, S. Kayal, S. Sarnacki, A. Cumano, G. Lauvau, F. Geissmann
Monitoring of Blood Vessels and Tissues by a Population of Monocytes with Patrolling Behavior
|
Science 317, 666 |
2007 |
| 15 |
Geissmann, F., S. Jung, D. R. Littman
Blood Monocytes Consist of Two Principal Subsets with Distinct Migratory Properties
|
Immunity 19, 71 |
2003 |
| 16 |
van Furth, R., Z. A. Cohn
The Origin and Kinetics of Mononuclear Phagocytes
|
J. Exp. Med. 128, 415 |
1968 |
| 17 |
Materials and methods are available as supporting material on Science Online
|
|
|
| 18 |
Mebius, R. E., G. Kraal
Structure and function of the spleen
|
Nat. Rev. Immunol. 5, 606 |
2005 |
| 19 |
Mosser, D. M., J. P. Edwards
Exploring the full spectrum of macrophage activation
|
Nat. Rev. Immunol. 8, 958 |
2008 |
| 20 |
Kristiansen, M., J. H. Graversen, C. Jacobsen, O. Sonne, H. Hoffman, S. K. A. Law, S. K. Moestrup
Identification of the haemoglobin scavenger receptor
|
Nature 409, 198 |
2001 |
| 21 |
Tsou, C., W. Peters, Y. Si, S. Slaymaker, A. M. Aslanian, S. P. Weisberg, M. Mack, I. F. Charo
Critical roles for CCR2 and MCP-3 in monocyte mobilization from bone marrow and recruitment to inflammatory sites
|
J. Clin. Invest. 117, 902 |
2007 |
| 22 |
Weissleder, R., M. J. Pittet
Imaging in the era of molecular oncology
|
Nature 452, 580 |
2008 |
| 23 |
Nahrendorf, M., D. E. Sosnovik, P. Waterman, F. K. Swirski, A. N. Pande, E. Aikawa, J. Figueiredo, M. J. Pittet, R. Weissleder
Dual Channel Optical Tomographic Imaging of Leukocyte Recruitment and Protease Activity in the Healing Myocardial Infarct
|
Circ. Res. 100, 1218 |
2007 |
| 24 |
Medzhitov, R.
Origin and physiological roles of inflammation
|
Nature 454, 428 |
2008 |
| 25 |
Kintscher, U., S. Wakino, S. Kim, E. Fleck, W. A. Hsueh, R. E. Law
Angiotensin II Induces Migration and Pyk2/Paxillin Phosphorylation of Human Monocytes
|
Hypertension 37, 587 |
2001 |
| 26 |
AbdAlla, S., H. Lother, A. Langer, Y. el Faramawy, U. Quitterer
Factor XIIIA Transglutaminase Crosslinks AT1 Receptor Dimers of Monocytes at the Onset of Atherosclerosis
|
Cell 119, 343 |
2004 |
| 27 |
McAlpine, H. M., S. M. Cobbe
Neuroendocrine changes in acute myocardial infarction
|
Am. J. Med. 84, 61 |
1988 |
| 28 |
Davies, B. N., P. G. Withrington
The Actions of Drugs on the Smooth Muscle of the Capsule and Blood Vessels of the Spleen
|
Pharmacol. Rev. 25, 373 |
1973 |
| 29 |
Jung, S., J. Aliberti, P. Graemmel, M. J. Sunshine, G. W. Kreutzberg, A. Sher, D. R. Littman
Analysis of Fractalkine Receptor CX3CR1 Function by Targeted Deletion and Green Fluorescent Protein Reporter Gene Insertion
|
Mol. Cell. Bio. 20, 4106 |
2000 |
| 30 |
Bolstad, B. M., R. A. Irizarry, M. Åstrand, T. P. Speed
A comparison of normalization methods for high density oligonucleotide array data based on variance and bias
|
Bioinformatics 19, 185 |
2003 |
| 31 |
Holm, S.
A Simple Sequentially Rejective Multiple Test Procedure
|
Scand. J. Stat. 6, 65 |
1979 |
| 32 |
Akilesh, S., D. J. Shaffer, D. Roopenian
Customized Molecular Phenotyping by Quantitative Gene Expression and Pattern Recognition Analysis
|
Genome Res. 13, 1719 |
2003 |
| 33 |
Bunster, E., R. K. Meyer
An Improved Method of Parabiosis
|
Anat. Rec. 57, 339 |
1933 |
| 34 |
Nahrendorf, M., et al.
Factor XIII Deficiency Causes Cardiac Rupture, Impairs Wound Healing, and Aggravates Cardiac Remodeling in Mice With Myocardial Infarction
|
Circulation 113, 1196 |
2006 |
| 35 |
Nahrendorf, M., D. E. Sosnovik, P. Waterman, F. K. Swirski, A. N. Pande, E. Aikawa, J. Figueiredo, M. J. Pittet, R. Weissleder
Dual Channel Optical Tomographic Imaging of Leukocyte Recruitment and Protease Activity in the Healing Myocardial Infarct
|
Circ. Res. 100, 1218 |
2007 |
| 36 |
Alencar, H., U. Mahmood, Y. Kawano, T. Hirata, R. Weissleder
Novel multi-wavelength microscopic scanner for mouse imaging
|
Neoplasia 7, 977 |
2005 |
| 37 |
Liu, K., C. Waskow, X. Liu, K. Yao, J. Hoh, M. Nussenzweig
Origin of dendritic cells in peripheral lymphoid organs of mice
|
Nat. Immunol. 8, 578 |
2007 |
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.
