Two physically, functionally, and developmentally distinct peritoneal macrophage subsets

Mouse PerC Contains a Variety of Immune Cells from the Lymphoid and Myeloid Lineages.

To identify the immune cell subsets present in the mouse PerC, we used high-dimensional flow cytometry (i.e., FACS) with stain sets capable of distinguishing as many as 11 markers per cell. We collected data for single live PerC cells from untreated BALB/c adult mice and gated to reveal the following subsets (Fig. 1): B cells, MØ, dendritic cells (DCs), eosinophils, mast cells, neutrophils, T cells, natural killer cells, and invariant natural killer T cells.

Approximately 40% of the PerC cells are B-lymphocytes (CD19⁺; Fig. 1, second row). Among the remaining “non-B” PerC cells, the majority coexpress CD11b and F4/80 and hence are appropriately characterized as MØ. Among these, we find the two unique MØ subsets that are the principal focus of this manuscript.

Peritoneal MØ Are Composed of Two Distinct Subsets: SPMs and LPMs.

Within the CD11b^hi population (gated as in Fig. 1), we find two subsets that differ strikingly in size, surface marker phenotype, and function. Based on size differences determined by FACS scatter profiles and microscopy images (Fig. 2 A and B), we provisionally refer to these subsets as SPMs and LPMs. LPM is the most abundant MØ subset (approximately 90% of PerC MØ) in unstimulated mice; SPM accounts for the remaining 10% of the PerC MØ (Fig. 1, last panel).

The expression of a series of surface markers clearly distinguishes SPM from LPM (Table 1). Both subsets express the prototypical MØ markers CD11b and F4/80. However, LPM expresses approximately three times as much CD11b and approximately 80 times as much F4/80 as SPM, setting the two subsets apart from each other and from other PerC cells (Fig. 1 and Table 1).

Importantly, the expression of three additional surface markers qualitatively distinguishes SPMs from LPMs. Thus, LPMs express Gr-1 and AA4.1 whereas SPMs do not (Table 1). In contrast, SPMs express MHC- II (I-A) whereas LPM strikingly do not express this classical marker, measured by FACS analyses (Table 1).

The quantitative expression of a series of other well known surface markers also distinguishes SPMs from LPMs (Table 1). Notably, CD40, CD80, CD86, CD11c (N418 MAb), and TLR4 are all expressed at higher levels on LPMs than SPMs.

Eosinophils, which are commonly found in PerC, will contaminate the SPM and LPM compartments unless explicitly excluded by the gating strategy. The eosinophils express F4/80 at a level intermediate between SPM and LPM (Fig. 1, last panel). However, they can be excluded because they characteristically express low levels of CD11b and low to intermediate levels of Gr-1, and have a unique scatter profile (CD11b^low, Gr-1^int, and SSc^hi; Figs. 1 and 3 and Fig. S1). Therefore, we standardly use a gating strategy that excludes these cells from the SPM and LPM subsets. Importantly, these data strongly argue against the common practice of using either F4/80 or CD11b as the sole surface marker for identifying or isolating MØ.

SPMs and LPMs Are Distinct from DCs and Monocytes.

DCs in PerC and spleen are smaller than either SPMs or LPMs and do not express CD11b and F4/80 at the levels found on SPMs and LPMs (Fig. 1, third row). In addition, PerC and splenic DCs both express high levels of CD11c, although PerC DCs have only approximately half as much CD11c as splenic DCs (Table 1). SPMs and LPMs, in contrast, express much less CD11c. This difference is detectable when CD11c is revealed by either HL3 or N418 monoclonal anti CD11c. Interestingly, staining with the monoclonal HL3 detects very little (if any) CD11c on LPMs whereas staining with the monoclonal N418 reveals somewhat more CD11c on LPMs than SPMs (Table 1).

Monocytes differ from SPMs and LPMs with respect to phenotype and principal location. Monocytes are present in blood (Fig. 3) and migrate to tissues to become resident MØ (8, 9). In contrast, neither SPMs nor LPMs are detectable in blood (Fig. 3), and these cells are virtually absent in spleen and lymph nodes (Fig. S2). We note, however, that the L-selectin CD62L is expressed on approximately half of SPMs in PerC (Fig. S3), suggesting that, with appropriate stimulation, SPMs may be able to migrate to lymphoid organs.

Phenotypically, SPMs and LPMs are clearly distinct from monocytes. CD11b expression is markedly higher on both SPM and LPM than it is on monocytes, and indeed on all blood leukocytes (Fig. 3). In contrast, Ly-6C is not detectable on SPMs or LPMs but is very high on monocytes (8, 9) (confirmed in Fig. 3). F4/80 expression on SPMs and LPMs also distinguishes these cells from blood leukocytes (Fig. 3), even though the F4/80 levels on SPMs are only a little above background.

SPM and LPM Are Present in All Mouse Strains Tested.

SPMs and LPMs are detectable in PerC from all of the strains analyzed in this study, including BALB/c, C57BL/6, 129/S6, FVB/N, SJL/J, and RAG^−/− (Fig. S4). The relatively constant frequencies of these subsets in PerC, and their similar proportionality among non-B PerC leukocytes suggests that there may be an evolutionary advantage to maintenance of the composition of the apparently free-living peritoneal leukocytes.

SPM and LPM Are Functionally Distinct.

Origins of the SPM Subset.

Studies described in the previous section were conducted at 20 h after i.p. LPS injection because, at this time, PerC SPM and LPM frequencies are still comparable to the frequencies in unstimulated mice. However, by 2 d after stimulation, SPM becomes the dominant PerC MØ subset and LPM frequencies dwindle to less than 10% (Fig. 6). The most dramatic shifts occur in mice stimulated with higher doses of LPS (≥10 μg).

To determine whether this shift occurs because LPMs differentiate to SPMs, we sorted LPMs, labeled the sorted cells with CFDA-SE, and transferred them into PerC of unstimulated mice. One hour later, we stimulated the recipient mice with LPS and, at various times thereafter, we harvested PerC and determined the phenotype of the CFDA-SE⁺ cells.

Results from these studies showed that, whether tested at 9 h or 2 d after stimulation, the sorted LPM population retains its LPM phenotype (Fig. 7). No CFDA-SE⁺ cells appear in the SPM compartment, despite the virtual loss of the resident and the injected LPM cells by 2 d after stimulation. Thus, we conclude that LPS stimulation does not induce LPMs to differentiate to SPMs, and hence that LPM differentiation does not account for the increase in SPM frequency.

In contrast, examination of host PerC cells shows clearly that LPS stimulation induces a massive influx of cells that initially express the phenotype of the circulating monocytes in blood (Ly-6C^hiMHC-II^–). With time, the phenotype expressed by this large population of “monocytes” shifts toward the typical Ly-6C^–MHC-II^hi SPM phenotype (Fig. 6). Thus, our findings indicate that a wave of blood monocytes enters the PerC shortly after LPS stimulation and, over time, differentiates to SPMs.

Thioglycolate-Elicited MØ.

Intraperitoneal injection of thioglycolate is widely used to enrich the yield of so-called activated PerC MØ, which are commonly collected 4 d after thioglycolate stimulation for use in functional MØ assays (12 –14). Examination of PerC harvested from thioglycolate-stimulated animals at this time shows a shift in SPM and LPM frequencies similar to that induced by LPS (Fig. 8). LPM, normally the dominant PerC MØ subset, is barely detectable (<5% of total MØ). Thus, 4 d after thioglycolate stimulation, the myeloid population in PerC mainly contains SPM (FSc^low, CD11b^int, F4/80^low, Ly-6C^–, Gr-1^–). However, it also contains a small subset of monocytes (Ly-6C^hi) that most likely have infiltrated from blood (Fig. 8).

In addition, as in the response to LPS stimulation, we find that the monocyte population in thioglycolate-stimulated PerC shifts with time toward expression of the SPM phenotype. From d 1 onward, the “monocytes” in the thioglycolate-stimulated PerC show decreasing levels of Ly-6C and, somewhat later, increasing levels of MHC-II. By d 4, SPMs predominate with only a small percentage of cells expressing a typical monocyte phenotype (Fig. 8).

Overall, these findings indicate that results from studies with thioglycolate-elicited MØ may mainly reflect the behavior of SPMs and of monocytes that have recently developed into SPMs. However, it is important to recognize that, in addition to MØ, and regardless of harvest day, thioglycolate-stimulated PerC commonly contains additional subsets of immune cells that may participate in the responses being measured. Notably, on d 1 after stimulation, neutrophils can outnumber MØ and lymphocytes. Further, even on d 4, lymphocytes and granulocytes can be present in relatively large numbers (Fig. S1) and can complicate interpretation of findings putatively as a result of the activity of thioglycolate-elicited PerC “macrophages.”

Mice and Tissue Preparation.

BALB/c, C57BL/6, 129/S6, FVB/N, SJL/J, and RAG1-KO mice, 6 to 8 weeks old, were purchased from Jackson Labs or bred locally. All mice were maintained at the Stanford Medical School Animal Care Facility. All experiments were conducted with institutional animal care and use committee approval. Peritoneal cells were harvested by injecting 7 mL of staining medium (deficient RPMI plus 3% newborn calf serum) into PerC. Spleen and lymph nodes were disrupted and resuspended to obtain single cell suspensions. Mouse blood was drawn into heparin-containing PBS solution. All cell samples were resuspended at 25 × 10⁶ cells/mL using custom RPMI 1640 medium deficient in biotin, L-glutamine, phenol red, riboflavin, and sodium bicarbonate (Invitrogen).

FACS.

Cell suspensions were preincubated with anti–CD16/CD32 mAb to block FcγRII/III receptors and stained on ice for 15 min. with the following fluorochrome-conjugated mAb in an 11-color staining combination: FITC-labeled anti-Ly-6C (AL-21), F4/80 (BM8), CD80 (16-10A1), or CD49b (DX5); PE-labeled anti–I-A^d (AMS-32.1), H-2K^d (SF1-1.1), CD40 (3/23), TLR4-MD-2 (MTS510), or CD49b (DX5); PECy5-labeled anti-CD5 (53–7.3); PECy5.5-labeled anti-CD19 (1D3) or CD11c (N418); PECy7-labeled anti–Gr-1 (RB6-8C5) or IgM (331); APC-labeled anti-B220 (RA3–6B2), CD86 (GL-1), CD117 (c-kit 2B8) or AA4.1; Alexa700-labeled anti-IgD (11-26); APCCy7-labeled anti-CD11b (M1/70); Pacific Blue–labeled anti-F4/80 (BM8) or Gr-1 (RB6-8C5) and biotin-labeled anti-CD11c (HL3). Cells were then washed and stained again on ice for 15 min with streptavidin Qdot 605 (Invitrogen) to reveal biotin-coupled antibodies. Antibodies were either purchased (Invitrogen and BD Pharmingen) or conjugated in our laboratory. After washing, stained cells were resuspended in 10 μg/mL propidium iodide, to exclude dead (i.e., propidium iodide–negative) cells. Cells were analyzed and sorted on Stanford FACS facility instruments (Becton Dickinson LSRII or FACSAria). Data were collected for 0.2 to 1 × 10⁶ cells. Staining protocols were designed with CytoGenie software (WoodSideLogic); data were analyzed with FlowJo software (TreeStar). To distinguish autofluorescent cells from cells expressing low levels of individual surface markers, we established upper thresholds for autofluorescence by staining samples with fluorescence-minus-one control stain sets (18, 19) in which a reagent for a channel of interest is omitted.

In Vitro and in Vivo Stimulation.

In vitro, 10⁶ PerC cells were cultured with and without 1μg/mL of LPS (E. coli 055:B5; Sigma) in a final volume of 500 μL of RPMI-1640 plus 10% FCS. Cells were incubated at physiologic oxygen levels (5% O₂) for indicated times. For NO measurement, cells were resuspended in 1mL of medium containing 0.1% probenecid (pH 7.4) and maintained subsequently in this medium. Cells were incubated in the presence of 1 μM of 4-amino-5-methylamino-2′,7′-difluorescein (DAF-FM) diacetate (Molecular Probes) at room temperature for 30 min and stained and analyzed by flow cytometry. In vivo, 1 mL of thioglycolate broth or 5 μg or 10 μg of LPS (in 200 μL of PBS solution) were injected i.p. At indicated times, PerC was harvested, stained, and analyzed.

In Vivo Phagocytosis Assay.

E. coli (K-12) bacteria (5 × 10⁶) expressing GFP were taken from overnight culture and resuspended in 500 μL of PBS solution. The GFP⁺ bacteria were then injected into the PerC of BALB/c mice. After 2 h, total PerCs were harvested, stained, and analyzed for the presence of GFP⁺ bacteria.

Confocal Microscopy.

GFP⁺ SPMs and LPMs were FACS-sorted directly onto a coverslip, fixed in 4% formalin for 15 min, washed in PBS solution, and air-dried. Finally, the coverslip was mounted to the slide using ProLong Gold antifade reagent containing DAPI (Invitrogen; Molecular Probes). The images were collected on a microscope (Optiphot-2; Nikon) attached to a confocal laser scanning (MRC1024; Bio-Rad) by using LaserSharp software (Bio-Rad). To identify extracellular bacteria, we used primary rabbit polyclonal anti-LPS followed by Alexa594-labeled goat anti-rabbit antibodies. Thus, bacteria that have remained attached on the cell membrane (not internalized) appear red. Internalized bacteria were visualized as green. The blue staining shows phalloidin coupled to Alexa647, which stains actin and gives the outline of the cell cytoplasm. Volocity 3.0 software (Improvision) was used to format and then merge the green (GFP), red (Alexa594), and blue (Alexa647) images. To measure individual cell size (Fig. 2), differential interference contrast images were acquired by confocal microscopy (LSM 510 META; Zeiss) with a ×63 Plan Apochromat objective lens (NA 1.4). Cell diameter was determined using Zeiss software.