Collecting lymphatic vessels (CLVs), surrounded by fat and endowed with contractile muscle and valves, transport lymph from tissues after it is absorbed into lymphatic capillaries. CLVs are not known to participate in immune responses. In this study, we observed that the inherent permeability of CLVs allowed broad distribution of lymph components within surrounding fat for uptake by adjacent macrophages and dendritic cells (DCs) that actively interacted with CLVs. Endocytosis of lymph-derived Ags by these cells supported recall T cell responses in the fat and also generated Ag-bearing DCs for emigration into adjacent lymph nodes (LNs). Enhanced recruitment of DCs to inflammation-reactive LNs significantly relied on adipose tissue DCs to maintain sufficient numbers of Ag-bearing DCs as the LN expanded. Thus, CLVs coordinate inflammation and immunity within adipose depots and foster the generation of an unexpected pool of APCs for Ag transport into the adjacent LN.

Absorptive lymphatic capillaries with blind-ended termini are positioned in the parenchyma of most organs (1) and consist of a single layer of lymphatic endothelial cells with elegantly organized intercellular junctions (2). Lymphatic capillaries take up fluid, macromolecules, and immune cells including dendritic cells (DCs) and T cells that traverse afferent lymphatic vessels en route to lymph nodes (LNs) (16). In the intestine, lymphatic capillaries, called lacteals, are crucial for absorption of chylomicrons. Before reaching the LN, lymphatic capillaries converge successively into afferent collecting lymphatic vessels (CLVs) that no longer serve an absorptive function for either molecules or cells. Instead, collecting vessels, distinguished by luminal valves and an organized wall containing contractile cells that promote lymph propulsion (3), are specialized for efficient transport of lymph and its contents to the draining LN and ultimately beyond the node in efferent lymphatic vessels (1).

As collecting vessels leave the parenchyma of organs and extend to the LN, they are encased in white adipose tissue (1, 7). In contrast with lymphatic capillaries, cells of the immune system have not been found to enter CLVs (6). Hence collecting vessels have received little consideration as players in innate or adaptive immunity, but instead have been viewed simply as conduits for immune cell passage to and from LNs. Furthermore, the historical view has been that collecting lymphatics are relatively impermeable to solutes (8), in addition to cells, reinforcing the general idea that these vessels solely function in lymph transport. However, recently, the notion of the impermeability of collecting lymphatics to macromolecules was refuted by the demonstration that muscular collecting lymphatics of the rat mesentery are as permeable to macromolecules, such as albumin (65 kDa), as the adjacent venules (4). Transport of macromolecules across the collecting lymphatic wall is coupled to water flux and sensitive to lymph pressure (4). It remains unknown whether and how the unexpected physiological permeability of lymphatic collecting vessels affects the surrounding adipose tissue. In conditions of reduced lymphatic integrity due to haploinsufficiency of the key lymphatic transcription factor Prox-1, mesenteric lymphatics appear especially leaky, and this leakiness may drive adipocyte expansion and obesity (9).

In this study, we characterized CLVs in adipose tissue with respect to their relationship with MHC class II+ (MHC II+) cells of the immune system. Then, we tracked the fate of soluble Ags from the point of tissue delivery to the draining LN and focused on the typically discarded white adipose tissue (perinodal adipose tissue [PAT]) rich in CLVs that is upstream of the LN. We show that the inherent permeability of CLVs can lead to several related consequences, including the onset of inflammation in PAT in response to inflammatory stimulants flowing in lymph, local presentation of lymph-derived Ags to these fat depots, and arming PAT DCs with Ag. We had earlier reported that adjuvant-reactive LNs remodel as part of a coordinated inflammatory program to allow increased numbers of Ag-transporting DCs to enter the inflamed LNs (10). A major source for these cells appears to be the PAT DCs that have acquired lymph-derived Ags.

Seven- to 9-wk-old male mice were studied, including standard CD45.2+ (Ly5.2) WT (Jackson Laboratories) mice, CD45.1+ (Ly5.1) congenic mice (NCI), plt/plt mice (11) (maintained at Mount Sinai), TCR-transgenic TEa mice (12) (shared with us by J.S. Bromberg), CD11c-EYFP mice (13) (maintained at The Rockefeller University), or CCR7-deficient mice (stock #005794; Jackson Laboratories) all bred onto the C57BL/6 background. K14-VEGFR-3-Ig mice and control littermates on a mixed background were previously described (14). Mice were housed in a specific pathogen-free environment at Mount Sinai School of Medicine, The Rockefeller University, or Swiss Federal Institute of Technology and were used in accordance with institutional and federal policies. Male Sprague–Dawley and Crl:CD (SD) rats (150–300 g) were purchased from Harlan Laboratories or Charles River, respectively, housed at Texas A&M American Association for the Accreditation of Laboratory Animal Care–accredited animal facilities, and were used in accordance with institutional and federal policies.

With approval of The Committee for the Protection of Human Subjects at National Jewish Health, deidentified human lungs and associated LNs not suitable for transplantation and donated for medical research were obtained from the National Disease Research Interchange (Philadelphia, PA) and the International Institute for the Advancement of Medicine (Edison, NJ).

FITC painting.

For contact sensitization (FITC painting assay), FITC (Sigma-Aldrich) was dissolved in acetone and dibutylphthalate (1:1) at 8 mg/ml (15). Aliquots of this solution (25 μl) were applied onto the shaved interscapular mouse skin that is drained by the brachial LN. Mice were euthanized 18 h later.

To track T cells after FITC painting, we isolated total CD4+ T cells from draining LNs 4 d after FITC skin painting using anti-CD4 magnetic beads (clone L3T4; Miltenyi Biotec). Then 3.5 × 106 of these purified CD4+ T cells or naive CD4+ T cells were transferred to C57BL/6 recipient mice i.v. Five hours later, some recipient mice received FITC solution applied to the back skin and ears, whereas others were injected intradermally (i.d.) with EαGFP. Approximately 36 h later, CD4+ T cells in draining LNs and PATs were analyzed by flow cytometry, and ear thickness was measured using fine digital calipers. Calipers were carefully placed on the ears to measure the thickness of the same area of the ear in different mice with the same individual performing all analysis in a semiblinded manner in which mice in different experimental groups were identified, through their toe clip pattern, only after ear thickness was measured.

EαGFP.

Plasmids containing EαGFP or EαCherry constructs were kindly provided by M.K. Jenkins (University of Minnesota Medical School). EαGFP and EαCherry protein induction and purification were carried out as described previously (16). Endotoxin was removed by ToxinEraser Endotoxin Removal Kit (GenScript) and measured by ToxinSensor Chromogenic LAL Endotoxin Assay Kit (GenScript). The residual endotoxin in EαGFP and EαCherry was ∼8.5 × 10−5 endotoxin units/μg EαGFP or EαCherry. For i.d. immunization, 40 μg EαGFP or EαCherry was injected into the dermis of the back skin. For direct delivery into PAT, the interscapular skin of anesthetized mice was shaved and a narrow opening was cut into the skin for access to the PAT around the brachial node. A total of 4 μl EαGFP at 5 mg/ml was injected into the PAT; then the skin opening was sealed using Vetbond (3M). To track T cell proliferation and activation in mice treated with EαCherry, we sorted naive CD4+ T cells from the spleen of TEa transgenic mice (12) by positive staining for CD4 and negative staining for CD44. These cells were labeled with CFSE (Invitrogen) at 10 mM to follow their proliferation after injection in vivo. One million of CFSE-labeled TEa naive CD4+ T cells were then i.v. injected into each C57BL/6 recipient. After 24 h, 20 or 40 μg EαCherry was then i.d. or intra-PAT delivered to one PAT around brachial LN or one side of scapular back skin, respectively. Proliferation and activation of transferred TEa CD4+ T cells were then evaluated in brachial LNs 2 d after EαCherry delivery.

Dextrans.

To label skin lymphatic vessels, we injected 10 μl of 1% 500 kDa FITC-conjugated or 70 kDa tetramethylrhodamine isothiocyanate (TRITC)-conjugated, lysine-fixable dextran (Molecular Probes) into mouse skin 7–10 min before mice were euthanized.

For preparation of mouse tissue sections, brachial LNs and associated PAT were carefully dissected and fixed in 4% paraformaldehyde, dehydrated in alcohol, embedded in glycol methacrylate (JB-4; Polysciences), sectioned (2 μm in thickness), and visualized using epifluorescence microscopy. For immunostaining analysis in JB-4 sections, whole-mount staining of the LN and associated adipose depot was performed before embedding in JB-4 resin. Tissues were fixed in 0.5% paraformaldehyde overnight at 4°C, permeabilized in 1× PBS containing 0.1% Tween 20 (Sigma-Aldrich) for a day at 4°C, and blocked using mouse Fc block (BD Pharmingen) for an additional day at 4°C. Then goat anti-CCL21 (R&D) was added for 1 wk, washed, and followed by Cy3- or Cy2-conjugated anti-goat (Jackson Immunoresearch) for 3 d.

For multiphoton imaging of fixed tissue, the entire brachial LN and its PAT from mice treated with i.d. EαGFP for different time points and i.d. TRITC-dextran for 7 min before sacrifice were collected and fixed in 4% paraformaldehyde overnight at 4°C. Samples then were washed in 1× PBS and fixed onto a petri dish with instant glue (Krazy) and then imaged using a Bio-Rad Radiance 2000 multiphoton system mounted to a fixed-stage upright microscope with a Ti:sapphire laser pumped by a 10-W Verdi tunable at 720–920 nm with a two-channel external detection system and fitted with a 20×/0.95 NA water immersion objective. Image stacks were combined and analyzed using ImageJ or Volocity 5 (Perkin-Elmer) software.

For intravital imaging, mice were anesthetized with 100 mg ketamine, 15 mg xylazine, and 2.5 mg acepromazine per kilogram body weight. The hind legs of mice were shaved using a double-edged razor blade. Mice were restrained on a stage warmer at 37°C (BioTherm Micro S37; Biogenics), and an incision was made on the posterior side of one hind leg immediately below the knee joint. The leg was then held in position using a metal strap fixed to the stage warmer, with a small hole through which the lymphatic and adjacent fat pad could be visualized. Mice were imaged using an Olympus Fluoview FV 1000MPE multiphoton laser-scanning microscope connected to a Coherent Chameleon laser (tunable from 690 to 1040 nm) and an Olympus 25×/1.05 objective, controlled by Fluoview (FV-10) software (Olympus) at The Rockefeller Bio-Imaging Facility. To image EGFP, EYFP, and Evans blue, we set the excitation wavelength between 910 and 940 nm; emission light was split by 2 dichroic mirrors at 505 and 570 nm into 3 channels and band-pass filters optimized for detecting EGFP without bleed-through from EYFP. To create time-lapse sequences, we typically scanned 72 × 425 × 425-μm volumes of tissue at 6-μm Z-steps and 40-s intervals. Images were processed and analyzed using Volocity 5 (Perkin-Elmer) and Imaris 6 (Bitplane) software.

Brachial or inguinal or popliteal LNs were excised, teased with needles, and digested in collagenase D (Roche) for 30 min at 37°C. Cells were then lightly pressed through a 70-μm cell strainer and then washed, counted, and stained for flow cytometry.

For obtaining single-cell suspension from PATs, mice were perfused with 10 ml 1× Dulbecco’s PBS (DPBS), and PATs were collected and minced into small pieces and then digested in RPMI-1640 (cellgro) media containing 0.56 U/ml Liberase Blendzyme 3 (Roche) and 0.1 mg/ml DNase I (Sigma-Aldrich) for 1 h. Tissue was then homogenized using a 20-G needle. Debris was filtered through a 70-μm cell strainer, and erythrocytes were lysed by 1× BD Pharm Lyse (BD). Cells then were washed, counted, and stained for flow cytometry.

For flow cytometry, combinations of the following mAbs were used in staining. Acquisition of samples was performed using an LSR II (BD Bioscience) instrument, and data were analyzed with FlowJo software (Tree Star). The mAbs to the following molecules were obtained from BD Pharmingen or eBioscience except where otherwise indicated: CD11b, Gr-1 (recognizes Ly6G and Ly6C), MerTK (polyclonal goat; R&D Systems), goat IgG (R&D Systems), CD45, B220, CD8, IAb, CD25, CD3, Vα2, CD64, CD103, YAe, CD11c, CD44, CD4, Siglec F, Armenian hamster IgG isotype control, I-A;I-E, F4/80, rat IgG2a (Serotec), anti-FITC (Jackson Immunoresearch Laboratories), and mouse IgG1 isotype control. DAPI was used to gate our DAPI+ dead cells.

For immunostaining of single-cell suspensions, 100 μl of a suspension from LN or PATs was applied to Alcian blue–coated coverslips and then incubated in 37°C for 30 min, followed by fixation with 4% paraformaldehyde and washing with 1× PBS. Cells were permeabilized for 15 min at room temperature (RT) with medium containing 0.05% saponin (Sigma-Aldrich), 10% goat serum, and 10 mM glycine. Primary Abs, including Armenian hamster anti-CD11c (eBioscience) and mouse anti-IAb (BD Biosciences), were used and followed by secondary Abs, Cy3-conjugated anti-Armenian hamster, Cy5-conjugated anti-mouse, or Cy2-conjugated anti-FITC for detection. DAPI (100 ng/ml) solution was used for nuclei staining.

Optical projection tomography.

TRITC-dextran (70 kDa) was i.d. injected into WT mouse upper scapular skin for lymphatic vessel labeling. Ten minutes later, Alexa Fluor 488-dextran (3 kDa) was injected i.v. for blood vessel labeling. Mice were sacrificed 5 min later, and brachial LN and adipose tissue around it were collected together as a whole tissue and then was fixed in 4% paraformaldehyde. Fixed tissue was sent to Bioptonics (Edinburgh, U.K.) for further processing and scanning.

Quantitative real-time PCR.

Total RNA was extracted from whole frozen PATs first homogenized with Dounce homogenizer. TRIzol-LS (Invitrogen) was added and the RNeasy mini kit (Qiagen) was followed. First-strand cDNA was synthesized using SuperScript II Reverse Transcriptase (Invitrogen). To amplify cDNA, we used 2× SYBR Green JumpStart Taq ReadyMix for Quantitative PCR (Sigma); then real-time quantitative PCR was carried out in the Quantitative PCR Shared Resource Facility in Mount Sinai School of Medicine using the ABI PRISM 7900HT instrument. PCR cycling conditions were 95°C for 2 min for polymerase activation, then amplification for 40 cycles: 95°C for 15 s, 55°C for 15 s, and 72 for 30 s. Relative expression levels were calculated using Gapdh as endogenous control.

Primer sequences were as follows: Tnfa: forward, 5′-GACCCTCACACTCAGATCATCTTCT-3′, reverse, 5′-CCACTTGGTGGTTTGCTACGA-3′; Il6: forward, 5′-CGGCAAACCTAGTGCGTTAT-3′, reverse, 5′-TCTGACCACAGTGAGGAATGTC-3′; iNos: forward, 5′-ACTGGGGCAGTGGAGAGATT-3′, reverse, 5′-GGTCAAACTCTTGGGGTTGA-3′; Tlr4: forward, 5′-GCAGAAAATGCCAGGATGATG-3′, reverse, 5′-AACTACCTCTATGCAGGGATTCAAG-3′; Ccr2: forward, 5′-GGGAGACAGCAGATCGAGT-3′, reverse, 5′-TCCCTCCTTCCCTGCTTAAA-3′; Mcp-1: forward, 5′-GGCTCAGCCAGATGCAGTTA-3′, reverse, 5′-CCTACTCATTGGGATCATCT-3′; Cd36: forward, 5′-TTGTACCTATACTGTGGCTA-3′, reverse, 5′-CTTGTGTTTTGAACATTTCT-3′.

For isolated vessel immunostaining, fixed lymphatics were incubated in blocking solution (1% BSA, 5% normal goat serum in PBS) for 1 h at RT and then divided into two pieces. The two sections were incubated in blocking solution overnight at 4°C in the presence of intralumenal and extralumenal primary Abs or the corresponding normal Igs (negative control), respectively. The vessel sections were washed intralumenally and extralumenally in PBS three times for 5 min each and then intralumenally and extralumenally incubated with secondary Abs for 1 h at RT. The vessel sections were again intralumenally and extralumenally washed with PBS three times for 5 min each and then cannulated and tied onto two glass pipettes, pressurized to 2 cm H2O, and secured to the stage of the confocal microscope for immediate observation.

For whole-mount immunostaining, the intestines and associated mesenteries were exteriorized and dissected distal to the duodenojejunal flexure, proximal to the cecum, and dorsal to the mesenteric LNs, respectively, and placed in DPBS with 10 g/l BSA at RT. The intestine/mesentery was pinned into a silicone-coated petri dish, rinsed three times with DPBS, incubated in acetone for 20 min at 4°C, and washed in PBS four times for 10 min each. Mesenteric panels (∼20 × 50 mm) containing blood and lymphatic vessels were isolated and incubated 1 h at RT in blocking solution. The samples were then incubated 18 h at 4°C in blocking solution containing primary Abs. Negative controls were prepared as described earlier. The samples were washed three times for 20 min each in PBS before incubation for 2 h at RT in PBS containing 1% BSA and secondary Abs. The samples were washed three times for 20 min each followed by 30 min in PBS before being spread onto glass slides, dried, and mounted under coverslips with ProLong Gold Antifade Reagent (Invitrogen).

Primary Abs used include mouse anti-eNOS (IgG1, clone3; BD Transduction Laboratories), mouse anti-rat PECAM-1 (IgG1, clone TLD-3A12; BD Transduction Laboratories), mouse anti-MHC II (IgG2a, clone 10.3.6; Santa Cruz Biotechnology), and rabbit anti–α smooth muscle actin (ab5694; Abcam, Cambridge, MA). Secondary Abs include goat anti-mouse IgG1-Alexa Fluor 488, goat anti-mouse IgG2a-Alexa Fluor 647, and donkey anti-rabbit Alexa Fluor 594 (Invitrogen). All Ab concentrations were ∼10 μg/ml.

The samples were imaged using a Leica TCS SP2 AOBS confocal microscope (Leica Microsystems) with either a Leica HC PL APO 20× (dry, 0.7 NA) or a Leica U APO 340/cc 40× (water immersion, 1.15 NA) objective. At multiple sites within each sample, 0.5-μm z-axis steps were taken with a 1-Airy disk pinhole. The samples were excited with an argon ion laser at 488 nm and a He-Ne laser at 594 or 633 nm. Emission wavelengths were specifically selected for each fluorochrome. Z-stacks were taken at numerous sites along each isolated lymphatic and throughout the lymphatic network in whole-mount mesenteries from the first collecting lymphatics outside the intestinal wall all the way to the largest downstream collecting lymphatics near the nodal/antimesenteric border. Image reconstruction and orthogonal viewing on the image stacks were performed using the Leica Confocal Software and ImageJ64. The negative controls for all experiments were produced and analyzed via the same instrumental and image processing procedures.

All data are presented as mean ± SEM. The statistical significance of differences in means was analyzed with the unpaired, two-tailed Student t test. The p values <0.05 were considered significant. All statistical analyses were performed using Prism version 5.0 for Mac OS X (GraphPad).

Fat pads typically surround LNs. When we mapped the lymphatic drainage system in mouse anterior s.c. adipose tissue by using optical projection tomography (Fig. 1A–D), we identified a cluster of lymphatic vessels at the origin of the fat pad near skin that drained into a lymph sinus (Fig. 1A–D, arrowhead). From this sinus, a lymphatic vessel corresponding to the afferent collecting vessel drained into the brachial LN and then departed the LN as an efferent lymphatic vessel (Fig. 1A–D, arrows). This collecting vessel was surrounded by smooth muscle actin+ cells (Fig. 1E). Further associated with such lymphatic vessels were CD11c+ cells that were highly fluorescent cells in the CD11c-EYFP mouse strain (Fig. 1F). Cells with the highest levels of EYFP were DCs, not macrophages, because few adipose macrophages, identified as CD64+ MERTK+ cells (17), were fluorescent, in contrast with CD64MERTKCD11c+ MHC II+ DCs (Fig. 1G, FACS plots). Intravital imaging analyses revealed that PAT CD11c-EYFP+ DCs that surrounded the CLVs outside of the popliteal LN were highly motile and actively interacted with the exterior of these vessels, mobilizing along the lymphatic vessel wall, away from it, and within the adipose tissue (Fig. 1H, Supplemental Video 1).

FIGURE 1.

CLVs in PAT and their interactions with DCs. (AD) Optical projection tomography of the brachial LN removed from scapular skin depicts the gross relationship between this LN and the surrounding lymphatic vessels and PAT. Green represents lymphatic tracer; red represents blood tracer. White in (A) shows the contour of PAT around LNs, with (B)–(D) showing internal fluorescence in orientations identical to (A) or partially rotated (B and C). Convergent lymphatics (Conv LVs) near the skin pool at a lymphatic sinus (B, arrowhead); then a single major afferent lymphatic (Aff LV, arrows) moves along the truck of the PAT to the LN, with the efferent vessel following the blood supply emerging from the LN hilum. (E) Cross section of the afferent lymphatic in the PAT trunk showing smooth muscle actin and podoplanin costaining. (F) CD11c-YFP+ cells visualized outside of podoplanin+ lymphatic vessels in PAT. (G) Gating on macrophages, DCs, or eosinophils in PAT shows that only DCs are YFP+ in CD11c-YFP mice. (H) Time-lapse images of CD11c-YFP DCs around popliteal PAT after injection of Evans blue dye (fluoresces red) in the rear footpad. All images shown are representative of two independent experiments performed in triplicate.

FIGURE 1.

CLVs in PAT and their interactions with DCs. (AD) Optical projection tomography of the brachial LN removed from scapular skin depicts the gross relationship between this LN and the surrounding lymphatic vessels and PAT. Green represents lymphatic tracer; red represents blood tracer. White in (A) shows the contour of PAT around LNs, with (B)–(D) showing internal fluorescence in orientations identical to (A) or partially rotated (B and C). Convergent lymphatics (Conv LVs) near the skin pool at a lymphatic sinus (B, arrowhead); then a single major afferent lymphatic (Aff LV, arrows) moves along the truck of the PAT to the LN, with the efferent vessel following the blood supply emerging from the LN hilum. (E) Cross section of the afferent lymphatic in the PAT trunk showing smooth muscle actin and podoplanin costaining. (F) CD11c-YFP+ cells visualized outside of podoplanin+ lymphatic vessels in PAT. (G) Gating on macrophages, DCs, or eosinophils in PAT shows that only DCs are YFP+ in CD11c-YFP mice. (H) Time-lapse images of CD11c-YFP DCs around popliteal PAT after injection of Evans blue dye (fluoresces red) in the rear footpad. All images shown are representative of two independent experiments performed in triplicate.

Close modal

We also examined rat mesenteric lymphatic vessels; these were surrounded by MHC II+ cells that retained association with the vessels even when they were isolated from the mesentery (Fig. 2). These rat prenodal mesenteric collecting lymphatics (average diameter, 125 ± 21 mm) were invested with numerous MHC II+ cells that covered 38 ± 6% of the lymphatic wall surface area, scattered throughout all layers of the lymphatic wall (Fig. 2A–E). MHC II+ cells were also distributed throughout the nearby adipose and associated blood vessels (Fig. 2F, 2G).

FIGURE 2.

MHC II+ cells within the wall of rat mesenteric CLVs. (AE) Shown are z-axis projections of isolated, rat mesenteric collecting lymphatics stained for MHC II (green), α-smooth muscle actin (red), and eNOS (blue). Higher (A) and lower magnification (B) show the localization of MHC II+ cells within the α-smooth muscle actin+ lymphatic wall. (C–E) Panels reveal MHC II+ cells within the lymphatic wall, here costained for eNOS. Boxed insets in (C) show the downstream edge of valve leaflets; (D and E) higher magnification images from (C) (white squares) depicting the orthogonal cross-sectional views at the locations marked by the red and orange lines revealing that the macrophage bodies are primarily ablumenal to the lymphatic endothelial cells. (F and G) Shown are z-axis projections of rat mesenteric whole-mount preparations stained for MHC II+ cells (green) and endothelium by CD31 (blue). The collecting lymphatic wall is indicated by thin white lines. (F) Tissue section with an upstream lymphangion within the adipose near the gut; (G) tissue section with a lymphangion within the adipose downstream of (F), in the same lymphatic network near the LN. Scale bar, 100 μm unless otherwise indicated. Data in these panels are derived from two to eight experiments, with n ≥ 2 replicates per condition.

FIGURE 2.

MHC II+ cells within the wall of rat mesenteric CLVs. (AE) Shown are z-axis projections of isolated, rat mesenteric collecting lymphatics stained for MHC II (green), α-smooth muscle actin (red), and eNOS (blue). Higher (A) and lower magnification (B) show the localization of MHC II+ cells within the α-smooth muscle actin+ lymphatic wall. (C–E) Panels reveal MHC II+ cells within the lymphatic wall, here costained for eNOS. Boxed insets in (C) show the downstream edge of valve leaflets; (D and E) higher magnification images from (C) (white squares) depicting the orthogonal cross-sectional views at the locations marked by the red and orange lines revealing that the macrophage bodies are primarily ablumenal to the lymphatic endothelial cells. (F and G) Shown are z-axis projections of rat mesenteric whole-mount preparations stained for MHC II+ cells (green) and endothelium by CD31 (blue). The collecting lymphatic wall is indicated by thin white lines. (F) Tissue section with an upstream lymphangion within the adipose near the gut; (G) tissue section with a lymphangion within the adipose downstream of (F), in the same lymphatic network near the LN. Scale bar, 100 μm unless otherwise indicated. Data in these panels are derived from two to eight experiments, with n ≥ 2 replicates per condition.

Close modal

Continuing with the mouse as an experimental model, we investigated whether adipose tissue DCs positioned near CLVs would become charged with lymph-derived tracers that passed through CLVs, given their proximity to these permeable vessels. Accordingly, we applied a contact sensitizer mixed with chemically reactive FITC to the skin surface of WT mice to assess whether DCs in PAT became FITC+. In this so-called FITC painting assay, DCs transport FITC-conjugated macromolecules from skin to LNs over an 18-h period, while additional FITC-conjugated molecules are carried in the lymph in soluble form and are readily visualized in the LN sinus and intranodal conduits (5, 15). Eighteen hours after applying FITC sensitizer to the skin surface, brightly fluorescent cells were found not only in draining LNs but were also nested between adipocytes throughout the PAT (Fig. 3A–C). FITC+ cells recovered from the PAT were not limited to DCs. Instead, FITC was taken up by all types of cells in the adipose tissue (Fig. 3D). However, macrophages and DCs acquired significantly more FITC than other cells, such as eosinophils (Fig. 3D), likely because of their robust endocytic capacity.

FIGURE 3.

Passage of soluble Ags from CLVs to adipose tissue phagocytes. (A and B) Low-power cross sections depicting PAT outside of the brachial LN 18 h after application to the skin surface of a contact sensitizer containing FITC (green) as a hapten (B) compared with the same PAT without FITC painting in (A). (C) Higher-power view of FITC+ cells (green) in PAT as in (B). (D) Flow-cytometric analysis to quantify FITC uptake 18 h after FITC skin painting in DCs (CD45+CD11c+MHCII+MERTKCD64), macrophages (CD45+MERTK+CD64+), and eosinophils (CD45+MERTKSiglecF+). (E) Low-power tile-reconstructed view of a cross section of PAT and brachial LN 20 h after EαGFP (green) was administered i.d. and 10 min after 70 kDa TRITC-dextran (red) was injected i.d. (F) Higher-power view of PAT cross section 10 min after FITC-dextran (green) i.d. shows a collecting vessel wall with dextran-enriched cells nearby (arrow). (G) Mice with defective lymphatics resulting from expression of VEGFR3-Ig from the K14 promoter and their littermate WT controls were subjected to FITC skin painting for 18 h. FITC+ cells in PAT cross sections were counted (n = 3/group). Arrows point to the subcapsular sinus in each strain. Numbers above scale bars correspond to number of micrometers. All images shown are representative of at least three animals per group. *p < 0.05.

FIGURE 3.

Passage of soluble Ags from CLVs to adipose tissue phagocytes. (A and B) Low-power cross sections depicting PAT outside of the brachial LN 18 h after application to the skin surface of a contact sensitizer containing FITC (green) as a hapten (B) compared with the same PAT without FITC painting in (A). (C) Higher-power view of FITC+ cells (green) in PAT as in (B). (D) Flow-cytometric analysis to quantify FITC uptake 18 h after FITC skin painting in DCs (CD45+CD11c+MHCII+MERTKCD64), macrophages (CD45+MERTK+CD64+), and eosinophils (CD45+MERTKSiglecF+). (E) Low-power tile-reconstructed view of a cross section of PAT and brachial LN 20 h after EαGFP (green) was administered i.d. and 10 min after 70 kDa TRITC-dextran (red) was injected i.d. (F) Higher-power view of PAT cross section 10 min after FITC-dextran (green) i.d. shows a collecting vessel wall with dextran-enriched cells nearby (arrow). (G) Mice with defective lymphatics resulting from expression of VEGFR3-Ig from the K14 promoter and their littermate WT controls were subjected to FITC skin painting for 18 h. FITC+ cells in PAT cross sections were counted (n = 3/group). Arrows point to the subcapsular sinus in each strain. Numbers above scale bars correspond to number of micrometers. All images shown are representative of at least three animals per group. *p < 0.05.

Close modal

Intradermal injection of recombinant EαGFP (16) also distributed broadly within PAT. Administration of fluorescent dextran as a tracer that readily drains through and therefore identifies lymphatic vessels (18) led to fluorescent cells in PAT and within the wall of CLVs in only 7–10 min (Fig. 3E, 3F, Supplemental Fig. 1). This rapid rate of labeling, far faster than DCs emigrate from the dermis to LNs (16), suggested that dextran was acquired locally by cells resident within PAT. Indeed, after FITC skin painting, more than half of the FITC+ cells appeared in PAT within 20 min, whereas migration of FITC+ DCs from skin was much slower (15, 19), thus eliminating the possibility that FITC+ cells in PAT emigrated to LNs first and then mobilized into PAT. Injection of labeled lymph-migrating DCs into the skin did not result in their appearance within PAT (E.L. Kuan and G.J. Randolph, unpublished observations). Taken together, these data indicate that acquisition of lymph-derived tracer by PAT cells was due to local sampling and uptake of material within lymph.

Accumulation of FITC+ cells in PAT after FITC skin painting depended on a functional lymphatic network, because the number of FITC+ cells in PAT was >75% reduced in K14 VEGFR3-Ig mice, which lack skin lymphatic capillaries and cannot transport lymph to skin-draining LNs (14, 20) (Fig. 3G). Considering the importance of lymphatic vessels in promoting the distribution of Ag to PAT, we conducted further imaging studies to visualize the fate of lymph-borne tracers as they passed through the collecting lymphatics within PAT. Multiphoton microscopy of fixed specimens of PAT, prepared 12 h after EαGFP and 10 min after TRITC-dextran was injected into the mouse dermis, confirmed that Ag-acquiring cells were present within the wall of CLVs. Cells that acquired TRITC-dextran within minutes after its administration were all EαGFP+, indicating that after uptake of EαGFP, these phagocytes remain local and were not stimulated to emigrate to LNs (Fig. 4A, arrows, 4B). Lymphocytes (tracer-negative) within the lymph were clearly observed (Fig. 4B, arrows). Although most of the phagocytic cells remained partially anchored within the collecting vessel wall or just outside of it, occasional GFP+ cytoplasmic projections appeared to extend into the lymphatic lumen (Fig. 4A, 4B).

FIGURE 4.

Endocytic acquisition of tracers by phagocytes in PAT. (A) Z-stacks acquired from multiphoton microscopy on fixed PAT obtained from mice injected with EαGFP i.d. 12 h previously and then injected with TRITC-dextran (red) 10 min before euthanasia. Red color identifies lymphatic lumen filled with TRITC dextran and cells acquiring TRITC dextran. Left panel shows maximum projection of z-stacks compiled through 200 μm PAT. Right panels show three individual z stack images; upper panels demonstrate both green (FITC) and red (TRITC) channels, and lower panels depict only the TRITC channel (in white contrast). Arrows indicate cells that are both FITC+ and TRITC. (B) Higher-power view of Z-stack image. Dark circles (arrows) in the lumen are poorly endocytic T cells. Arrowhead indicates cells that took up EαGFP and extend pseudopods into the lumen. (C and D) FITC-BSA was injected in the exteriorized rat mesentery. Live confocal imaging of the fluorescent tracer as it passed through mesenteric collecting vessels began immediately. Red line delineates the border of the lymphatic vessel with surrounding adipose tissue and the lymphatic lumen as marked. Arrows depict cell bodies that appear to concentrate the tracer. Live imaging precluded immunostaining to identify these cells. Scale bars, 30 μm. Data in each panel are representative of two to three independent experiments.

FIGURE 4.

Endocytic acquisition of tracers by phagocytes in PAT. (A) Z-stacks acquired from multiphoton microscopy on fixed PAT obtained from mice injected with EαGFP i.d. 12 h previously and then injected with TRITC-dextran (red) 10 min before euthanasia. Red color identifies lymphatic lumen filled with TRITC dextran and cells acquiring TRITC dextran. Left panel shows maximum projection of z-stacks compiled through 200 μm PAT. Right panels show three individual z stack images; upper panels demonstrate both green (FITC) and red (TRITC) channels, and lower panels depict only the TRITC channel (in white contrast). Arrows indicate cells that are both FITC+ and TRITC. (B) Higher-power view of Z-stack image. Dark circles (arrows) in the lumen are poorly endocytic T cells. Arrowhead indicates cells that took up EαGFP and extend pseudopods into the lumen. (C and D) FITC-BSA was injected in the exteriorized rat mesentery. Live confocal imaging of the fluorescent tracer as it passed through mesenteric collecting vessels began immediately. Red line delineates the border of the lymphatic vessel with surrounding adipose tissue and the lymphatic lumen as marked. Arrows depict cell bodies that appear to concentrate the tracer. Live imaging precluded immunostaining to identify these cells. Scale bars, 30 μm. Data in each panel are representative of two to three independent experiments.

Close modal

Collectively, these data indicate that soluble components in lymph are shared with other cells within adipose tissue stromal vascular fraction in mice.

To investigate whether this conclusion could be extended to other species, we injected FITC-conjugated bovine albumin directly into the rat lamina propria during intravital imaging. We observed cell bodies concentrating FITC-albumin along the borders of the mesenteric collecting vessel within 40 min (Fig. 4C, 4D), suggesting the presence of endocytic cells in the lymphatic wall able to sample lymph contents as observed in mice. Furthermore, we observed that PAT around peribronchial human LNs obtained from a cigarette smoker contained phagocytes filled with dark particulate matter that were likely derived from cigarette smoke (Supplemental Fig. 2). Considering that such particulate matter is not found in circulating cells, the most probable mechanism behind its localization within PAT phagocytes is through emigration of phagocytes from lymph into PAT or, particularly, from lymph sampling. Collectively, the data from studies conducted in rodents suggest that lymph sampling leads to Ag deposition in PAT, a mechanism that could also be responsible for depositing foreign material in PAT in humans.

After FITC skin painting (Fig. 5A), leukocytes, especially CD11c+ cells, were recruited to PAT. A similar response was evident after EαGFP was injected i.d. (Fig. 5A). Gene expression profiles in PAT showed increases in TNF-α, IL-6, iNOS, TLR4, CCR2, MCP-1, and CD36 (Fig. 5B). To additionally assess whether lymph-sampled Ag might influence T cell accumulation in PAT in an Ag-dependent manner, we harvested CD45.1+CD4+ T cells in these draining LNs 4 d after FITC skin painting. These T cells, enriched in primed T cells specific for contact sensitizer Ags (21), or naive T cells were transferred i.v. into nonimmunized recipient CD45.2+ WT mice. Some of these WT recipients received no further manipulations, whereas other cohorts were immunized i.d. with EαGFP 1 d later, serving as an irrelevant Ag in this case, or by FITC skin painting, which set the stage for a recall response (21). Ear-swelling tests indicated that transferred T cells supported the expected recall response (21) after FITC painting challenge (data not shown). Transferred reactive T cells and endogenous CD4+ T cells accumulated more in the PAT in recipient mice that were challenged with FITC painting compared with no Ag or the irrelevant Ag-EαGFP treatment (Fig. 5C, 5D).

FIGURE 5.

Inflammatory responses in PAT after skin immunization. (A) Each line in the graphs represents an independent experiment in which the fold-increase in PAT CD11c+ inflammatory cells was measured 18 h after epicutaneous application of FITC sensitizer or 18 h after EαGFP was injected i.d. (B) Quantitative PCR analysis to determine fold increase in mRNA for inflammatory mediators and key adipose-derived cytokines, TNF-α, IL-6, iNOS, TLR4, CCR2, MCP-1, and CD36, 12 h after application of FITC sensitizer to skin. (C and D) CD4+ T cells were purified from either naive LNs (+Naive T) or reactive LNs (+T) of CD45.1+ WT mice 4 d after FITC skin painting and transferred into CD45.2+ WT mice. These recipients were challenged 5 h later with FITC painting or an irrelevant Ag to which they were naive (EαGFP) or not challenged (No Ag). Accumulation of transferred CD45.1+ CD4+ and endogenous CD45.2+ CD4+ T cells in the site of skin challenge or the associated PAT was analyzed ∼36 h after challenge. (C) Representative FACS plots of transferred T cells in skin or in PATs. (D) Graphs chart the number of T cells accumulated in PAT of Ag-challenged mice, expressed as fold increase over the number accumulated in the absence of Ag challenge. All data were obtained from at least two independent experiments with more than three replicates per group in each experiment. **p < 0.01.

FIGURE 5.

Inflammatory responses in PAT after skin immunization. (A) Each line in the graphs represents an independent experiment in which the fold-increase in PAT CD11c+ inflammatory cells was measured 18 h after epicutaneous application of FITC sensitizer or 18 h after EαGFP was injected i.d. (B) Quantitative PCR analysis to determine fold increase in mRNA for inflammatory mediators and key adipose-derived cytokines, TNF-α, IL-6, iNOS, TLR4, CCR2, MCP-1, and CD36, 12 h after application of FITC sensitizer to skin. (C and D) CD4+ T cells were purified from either naive LNs (+Naive T) or reactive LNs (+T) of CD45.1+ WT mice 4 d after FITC skin painting and transferred into CD45.2+ WT mice. These recipients were challenged 5 h later with FITC painting or an irrelevant Ag to which they were naive (EαGFP) or not challenged (No Ag). Accumulation of transferred CD45.1+ CD4+ and endogenous CD45.2+ CD4+ T cells in the site of skin challenge or the associated PAT was analyzed ∼36 h after challenge. (C) Representative FACS plots of transferred T cells in skin or in PATs. (D) Graphs chart the number of T cells accumulated in PAT of Ag-challenged mice, expressed as fold increase over the number accumulated in the absence of Ag challenge. All data were obtained from at least two independent experiments with more than three replicates per group in each experiment. **p < 0.01.

Close modal

Although DCs emigrating to skin-draining LNs from lymphatic vessels are thought to arise from one of several skin DC subsets (22), the present findings indicating that lymph-derived Ags could be acquired by PAT DCs raised the possibility that PAT DCs might enter skin-draining LNs. To test this idea, we bypassed the dermal route of Ag administration, to avoid labeling dermal DCs, and injected EαGFP into PAT directly. Over the course of ∼8 h, the focal injection of EαGFP within PAT led to many individual EαGFP+ cells scattered throughout the surrounding adipose (Fig. 6A, 6B). Some of the labeled GFP+ cells migrated to the adjacent LN (Fig. 6A, right micrograph), peaking there between 12 and 36 h (Fig. 6B). These cells displayed Eα peptide on MHC II (I-Ab), as detected using the YAe mAb (23) and coexpression of GFP (Fig. 6C). However, in concert with earlier studies, a small subset of cells with the highest levels of GFP and YAe reactivity originated solely from the skin (16). Activation of TEa transgenic T cells, whose TCR specifically engages I-Ab complex with the Eα recognized by YAe mAb (12), occurred as efficiently for PAT-delivered EαGFP as it did for skin-delivered EαGFP (Fig. 6D). The rather slow time frame (hours rather than minutes) of YAe+ DCs appearing in the LN (Fig. 6B) and the requirement for CCR7 or its ligands (Fig. 6E) revealed that the appearance of YAe+ DCs in the LN after EαGFP was administered to PAT was not the result of passive movement of EαGFP into the LN from the adipose tissue. Adoptive transfer of green fluorescent ex vivo–generated DCs into PAT led to a small fraction entering the LN, as is true with DCs injected into skin. Sections through the PAT where the DCs were injected revealed a large collection of DCs at the site of injection with some DCs radiating away from the central site in the direction of the PAT collecting lymphatics (Fig. 6F). This pattern was not observed when CCR7−/− DCs were injected (Fig. 6G), suggesting that cells progressing toward the lymphatics from the injection site is CCR7 dependent. Indeed, we detected high levels of the CCR7 ligand CCL21 were displayed by lymphatic collecting vessels (Supplemental Fig. 3).

FIGURE 6.

DCs from PAT use CCR7 to home to adjacent LNs. (A) EαGFP or EαCherry (20 μg in 2 μl) was injected (intra-PAT) into two sites of mouse PAT around brachial LN. The distribution of EαGFP+ cells was analyzed 18 h later. Arrowhead indicates injection site; arrow indicates EαGFP+ cells in LN. Dashed line delineates border of PAT and brachial LN on left, and the border of the LN B cell and T cell zone on right. (B) Total number of EαGFP+YAe+CD11c+CD45+ cells in PAT and LN over time after intra-PAT EαGFP delivery. At least three mice per time point were used. (C) EαGFP and YAe expression in CD11c+ cells from brachial LNs 18 h after intra-PAT (20 μg) or i.d. (40 μg in scapular skin) delivery of EαGFP. (D) Sorted, CFSE-labeled naive TEa transgenic T cells were transferred i.v. into naive recipient mice that 24 h later received intra-PAT (20 μg/PAT, black bars) or i.d. (40 μg in upper scapular skin, white bars) injection of EαCherry. Total proliferated TEa+CD4+CFSElo T cells and CD69+ or CD44+ proportions in draining LNs were quantified. (E) WT, CCR7 KO, or plt/plt mice received intra-PAT EαGFP. After 18 h, percent EαGFP+YAe+ cells among CD11c+ LN cells was analyzed. (F and G) Fluorescent bead-labeled bone marrow–derived DCs (green) from WT (F) or CCR7 KO (G) mice were injected directly into WT mice PAT for 24 h; 10 min before euthanasia, TRITC-dextran (red) was delivered i.d. to label lymphatics (white arrows). Numbers above scale bars correspond to number of micrometers. (A and C–G) Data were obtained from three independent experiments with at least three replicates per group in each experiment. Data are mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. NS, not statistically different.

FIGURE 6.

DCs from PAT use CCR7 to home to adjacent LNs. (A) EαGFP or EαCherry (20 μg in 2 μl) was injected (intra-PAT) into two sites of mouse PAT around brachial LN. The distribution of EαGFP+ cells was analyzed 18 h later. Arrowhead indicates injection site; arrow indicates EαGFP+ cells in LN. Dashed line delineates border of PAT and brachial LN on left, and the border of the LN B cell and T cell zone on right. (B) Total number of EαGFP+YAe+CD11c+CD45+ cells in PAT and LN over time after intra-PAT EαGFP delivery. At least three mice per time point were used. (C) EαGFP and YAe expression in CD11c+ cells from brachial LNs 18 h after intra-PAT (20 μg) or i.d. (40 μg in scapular skin) delivery of EαGFP. (D) Sorted, CFSE-labeled naive TEa transgenic T cells were transferred i.v. into naive recipient mice that 24 h later received intra-PAT (20 μg/PAT, black bars) or i.d. (40 μg in upper scapular skin, white bars) injection of EαCherry. Total proliferated TEa+CD4+CFSElo T cells and CD69+ or CD44+ proportions in draining LNs were quantified. (E) WT, CCR7 KO, or plt/plt mice received intra-PAT EαGFP. After 18 h, percent EαGFP+YAe+ cells among CD11c+ LN cells was analyzed. (F and G) Fluorescent bead-labeled bone marrow–derived DCs (green) from WT (F) or CCR7 KO (G) mice were injected directly into WT mice PAT for 24 h; 10 min before euthanasia, TRITC-dextran (red) was delivered i.d. to label lymphatics (white arrows). Numbers above scale bars correspond to number of micrometers. (A and C–G) Data were obtained from three independent experiments with at least three replicates per group in each experiment. Data are mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. NS, not statistically different.

Close modal

Our past work revealed that adjuvant-reactive LNs recruited more DCs as they undergo hypertrophy. In the PAT of mice receiving EαGFP i.d., however, we visualized fewer GFP+ cells in the spaces between adipocytes in mice treated with CFA in one front forepaw 3 d earlier compared with control mice not receiving CFA (Fig. 7A, 7B). In the same samples, more GFP+ cells were observed in adjacent LNs, raising the possibility that inflamed, CFA-reactive LNs recruited more GFP+ cells from the PAT. Thus, we set out to address whether and to what extent reactive LNs may draw upon surrounding adipose tissue in recruiting Ag-bearing DCs. To aid in distinguishing lymph-migratory DCs that originated from skin versus PAT, we took advantage of an ancillary observation that skin painting with TRITC did not label PAT cells (negative data not shown) and did not lead to soluble TRITC+ molecules in LN sinus or conduits (Fig. 7B, 7C). Instead, it appeared that TRITC, more reactive and hydrophobic than FITC, was carried to LNs only by lymph-migratory DCs that become TRITC-labeled in skin (24), although it did not flow through lymph in soluble form that would enable access to LN sinus and conduits and sampling by PAT macrophages and DCs. By contrast, as noted earlier in this study and others (15, 16), FITC and EαGFP filled LN sinus and conduits and were sampled by PAT macrophages and DCs (Fig. 7C, 7D). We thus set up experiments in which we carried out FITC or TRITC skin painting in different cohorts of WT mice. Because we had earlier shown that adjuvant-reactive LNs could support increased DC migration during the ensuing LN hypertrophy (e.g., induction of DC migration induced from the scapular skin 3 d after CFA was injected in a front footpad) (10), we quantified the number of FITC+ or TRITC+ DCs in LNs of different cohorts of mice pretreated or not with CFA in one front footpad. The number of migratory FITC+ DCs in the CFA-reactive brachial LN doubled compared with the number in nonreactive LNs (Fig. 7E). However, no differences were observed in the number of TRITC+ DCs in nonreactive versus CFA-reactive LNs (Fig. 7E). This finding was consistent with the possibility that FITC+ DCs in PAT, lacking TRITC+ counterparts in PAT following TRITC painting, might be a key source of migratory DCs to enter CFA-reactive LNs.

FIGURE 7.

Inflammation increases mobilization of DCs from PAT to the adjacent LN. Mouse brachial LNs were inflamed using CFA (+ CFA) or not (control). Then mice intra-PAT injection of EαGFP (20 μg/ PAT) or FITC or TRITC painting on the skin. PAT and the adjacent LN were harvested 18 h later for imaging or flow-cytometric analysis. (A) Photomicrographs depicting distribution and density of EαGFP+ cells in PAT of control or CFA-treated mice. (B) Total number of CD11c+EαGFP+YAe+ cells in PAT (upper panel) or LNs (lower panel) of control and CFA-treated mice. (C) Images of PAT from mice painted with FITC or TRITC for 18 h. White arrows indicate the subcapsular sinus that feeds the conduit system in draining LNs. Scale bar, 50 μm. (D) FITC (green) and TRITC (red) were painted together on mouse scapular back skin; 18 h later, the draining LN was collected for imaging analysis. Conduits (threadlike structures) in the LN are labeled with FITC, but not TRITC. (E) The total number of FITC+CD11c+ or TRITC+CD11c+ DCs in brachial LNs of control or CFA-treated mice. (F) In control or CFA-treated mice, upper scapular skin was injected i.d. with EαGFP (40 μg/site) and the same skin site was painted with TRITC 9 h later. Draining LNs were collected 17 h after TRITC painting, and the number of EαGFP+TRITCCD11c+MHCIIhi (black bar), EαGFP+TRITC+CD11c+MHCIIhi (white bar), and EαGFPTRITC+CD11c+MHCIIhi (gray bar) cells were quantified. Data depict mean ± SEM. All data were obtained from three independent experiments with more than three replicates per group in each experiment. *p < 0.05, **p < 0.01.

FIGURE 7.

Inflammation increases mobilization of DCs from PAT to the adjacent LN. Mouse brachial LNs were inflamed using CFA (+ CFA) or not (control). Then mice intra-PAT injection of EαGFP (20 μg/ PAT) or FITC or TRITC painting on the skin. PAT and the adjacent LN were harvested 18 h later for imaging or flow-cytometric analysis. (A) Photomicrographs depicting distribution and density of EαGFP+ cells in PAT of control or CFA-treated mice. (B) Total number of CD11c+EαGFP+YAe+ cells in PAT (upper panel) or LNs (lower panel) of control and CFA-treated mice. (C) Images of PAT from mice painted with FITC or TRITC for 18 h. White arrows indicate the subcapsular sinus that feeds the conduit system in draining LNs. Scale bar, 50 μm. (D) FITC (green) and TRITC (red) were painted together on mouse scapular back skin; 18 h later, the draining LN was collected for imaging analysis. Conduits (threadlike structures) in the LN are labeled with FITC, but not TRITC. (E) The total number of FITC+CD11c+ or TRITC+CD11c+ DCs in brachial LNs of control or CFA-treated mice. (F) In control or CFA-treated mice, upper scapular skin was injected i.d. with EαGFP (40 μg/site) and the same skin site was painted with TRITC 9 h later. Draining LNs were collected 17 h after TRITC painting, and the number of EαGFP+TRITCCD11c+MHCIIhi (black bar), EαGFP+TRITC+CD11c+MHCIIhi (white bar), and EαGFPTRITC+CD11c+MHCIIhi (gray bar) cells were quantified. Data depict mean ± SEM. All data were obtained from three independent experiments with more than three replicates per group in each experiment. *p < 0.05, **p < 0.01.

Close modal

To pursue this possibility further and quantify PAT-derived and skin-derived DCs simultaneously, we injected EαGFP i.d. to permit labeling of PAT DCs 4 h before applying TRITC to the skin of the same mice in the area that drains to the same brachial LN. When we harvested the LNs 18 h later, we found three populations of migratory DCs (CD11chiMHCIIhi) in the draining LN: TRITC+EαGFP+, TRITC+EαGFP, and much more rarely TRITCEαGFP+, with only the latter potentially originating from PAT (Fig. 7F). By contrast, when this experiment was performed in mice with CFA-reactive LNs, TRITCEαGFP+ cells were the only population to substantially increase and they increased sufficiently greatly (>3-fold) to comprise approximately one third of the Ag-labeled DCs in such LNs (Fig. 7F). When we repeated this experiment but instead injected EαGFP on the contralateral side of the mouse to make sure that TRITCEαGFP+ DCs did not arise from systemic spread and uptake of soluble EαGFP in DCs or DC precursors that might enter the LN through the high endothelial venules, we found no EαGFP+ DCs in CFA-reactive contralateral LN (negative result not shown). Thus, these data collectively suggest that PAT DCs emigrate to LNs, especially under conditions of LN hypertrophy, where they contribute substantially to the overall pool of migratory DCs.

Perinodal adipose tissue colocalizes with LNs throughout the body and is a conserved feature of mammalian anatomy (25). The immune function of this fat depot and its relationship with local LNs is not known. The fat pads around LNs extend beyond the nodes along the CLVs that feed into (afferent) and run out of (efferent) LNs, such that CLVs are most closely associated with PAT. We show in this study that the inherent permeability of CLVs is sufficient to broadcast Ags, passing within lymph to LNs, throughout PAT. The delivery of soluble Ags, such as FITC-conjugated endogenous proteins and Eα-GFP, is likely a passive consequence of the permeability of CLVs. This process exposes a large community of endocytic and phagocytic cells, particularly DCs and macrophages, to Ags that would otherwise be unavailable to them.

It is also possible that DCs or macrophages can actively sample the contents of the lymphatic lumen. DCs, in particular, were observed by intravital imaging to intimately interact with CLVs along with images displaying pseudopod-like projections in the lumen. Moreover, our findings of particulate matter in PAT around human bronchial LNs are consistent with this possibility, although anecdotal. Active lymph sampling by adipose DCs/macrophages could explain the accumulation of micro-organisms, such as Mycobacterium tuberculosis, in adipose tissue (26).

Our findings suggest persistent inflammation within fat depots is caused, at least in part, by disease-promoting Ags that course through the lymph in CLVs. Our observation that fat around CLVs can accumulate T cells with specificity to Ags derived from lymph contents supports the notion that changes in the adipose tissue T cell compartment associated with disease might likewise occur in response to Ags that originate from lymph. In this context, it is interesting to consider connections to adipose tissue inflammation in the context of obesity, where macrophages, DCs, and T cells appear to play a critical role (2729). Furthermore, mesenteric adipose tissue would be expected to be exposed to and accumulate lymph-derived molecules from the intestine, whereas PAT would acquire Ags from a variety of organs drained by the relevant LNs.

At these sites, signals derived from lymph may impact the adipocyte environment, perhaps to promote adipose growth (9) through cross talk with preadipocytes themselves located in walls of nearby vessels (30). Many PAT in mice and humans correspond with depots containing beige adipocytes (3135), which are capable of assuming a fat-storing white adipocyte phenotype or a fuel-demanding thermogenic brown fat. That macrophages have been shown to impact the activity of brown fat raises the possibility that factors in lymph that condition macrophages may, in turn, affect adipocyte differentiation (36). Thus, the present findings support the need for future studies of immunological diseases in the intestine and beyond, including Crohn disease or HIV, known to be linked in still obscure ways to changes in adipose tissue, lymphatic vessels, or both (25, 3741).

We have emphasized the uptake of Ags as a component of lymph capable of being processed by phagocytes. However, we recognize that the broadcast and sampling of endogenous macromolecules from lymph might also transmit to fat information about the physiological status of adjacent organs drained by a common lymphatic vasculature. An example might be the breakdown of hyaluronan within tissues in response to tissue injury (42). The receptor LYVE-1 on lymphatic capillaries in organs binds hyaluronan, but CLVs do not express it (43). In scenarios wherein hyaluronan breaks down to a sufficient degree to enter lymph and not be cleared in the lymphatic capillary before reaching the CLV, permeable CLVs will provide PAT DCs and macrophages with a signal via hyaluronan fragments. Because these promote DC migration to LNs (44), among other defense and healing reactions (42), the adipose tissue may become a critical source of signals that affect the response to injury, as well as a source of migratory DCs to enter the draining lymph.

It is clear that enhancing or dampening the number of lymph-migratory DCs bearing Ag impacts the quality and character of an ongoing immune response (19, 44, 45). We have had a long-standing interest in how the LN coordinates signals to undergo hypertrophy during a vigorous immune response and still maintain ideal ratios of lymphocytes, which enter often through HEV, to Ag-bearing DCs coming from lymph (10). We show in this study that one of the consequences of charging PAT DCs with Ags acquired through permeable CLVs is the supply to the LN of a reserve of Ag-bearing DCs for the hypertrophic LN. Thus, in addition to the wide array of different DC subtypes now described in the parenchyma of organs like skin (22), the DC compartment capable of acquiring and transporting Ags to LNs must be expanded to consider DC populations beyond the organ residing within the PAT.

We thank Marc Jenkins (University of Minnesota) for the gift of plasmids encoding EαGFP and EαCherry, Kari Alitalo (University of Helsinki) for the K14-VEGFR3 Ig transgenic mice, Jonathan Bromberg for sharing TCR-transgenic TEa mice, and Jens Stein (University of Bern) for protocols in whole-mount staining of the LN and adipose tissue. We are also grateful for the expert assistance we received at several Shared Resource Facilities (Imaging, Flow Cytometry, Quantitative PCR) at Mount Sinai School of Medicine, at the Integrated Microscopy Imaging Laboratory at the Texas A&M Health Science Center, and at The Rockefeller University Bio-Imaging Resource Center.

This work was supported by an Established Investigator Award from the American Heart Association, National Institutes of Health (NIH) Grants AI 049653 and R21 AG046743, a Nutrition Obesity Research Center Pilot and Feasibility subaward from the NIH (Grant P30 DK05341), and an Innovation Award from the Rainin Foundation (all to G.J.R.); NIH Grants HL084312 (to G.J.R. and M.A.S.), AI055037 (to M.L.D.), HL075199 (to D.C.Z.), HL70308 (to D.C.Z.), HL085659 (to D.C.Z.), AG030578 (to A.A.G.), and AI082982 (to R.J.M.); and an NIH supplement to C.V.J. linked to NIH Grant HL081151 to Peter Henson (National Jewish Health). Imaging studies conducted at The Rockefeller University were supported by the Empire State Stem Cell Fund through New York State Department of Health Contract C023046.

Opinions expressed in this article are solely those of the author and do not necessarily reflect those of the Empire State Stem Cell Fund, the New York State Department of Health, or the State of New York.

The online version of this article contains supplemental material.

Abbreviations used in this article:

CLV

collecting lymphatic vessel

DC

dendritic cell

DPBS

Dulbecco’s PBS

i.d.

intradermally

LN

lymph node

MHC II

MHC class II

PAT

perinodal adipose tissue

RT

room temperature

TRITC

tetramethylrhodamine isothiocyanate.

1
Schmid-Schönbein
G. W.
1990
.
Microlymphatics and lymph flow.
Physiol. Rev.
70
:
987
1028
.
2
Baluk
P.
,
Fuxe
J.
,
Hashizume
H.
,
Romano
T.
,
Lashnits
E.
,
Butz
S.
,
Vestweber
D.
,
Corada
M.
,
Molendini
C.
,
Dejana
E.
,
McDonald
D. M.
.
2007
.
Functionally specialized junctions between endothelial cells of lymphatic vessels.
J. Exp. Med.
204
:
2349
2362
.
3
Muthuchamy
M.
,
Zawieja
D.
.
2008
.
Molecular regulation of lymphatic contractility.
Ann. N. Y. Acad. Sci.
1131
:
89
99
.
4
Scallan
J. P.
,
Huxley
V. H.
.
2010
.
In vivo determination of collecting lymphatic vessel permeability to albumin: a role for lymphatics in exchange.
J. Physiol.
588
:
243
254
.
5
Randolph
G. J.
,
Angeli
V.
,
Swartz
M. A.
.
2005
.
Dendritic-cell trafficking to lymph nodes through lymphatic vessels.
Nat. Rev. Immunol.
5
:
617
628
.
6
Pflicke
H.
,
Sixt
M.
.
2009
.
Preformed portals facilitate dendritic cell entry into afferent lymphatic vessels.
J. Exp. Med.
206
:
2925
2935
.
7
Harvey
N. L.
2008
.
The link between lymphatic function and adipose biology.
Ann. N. Y. Acad. Sci.
1131
:
82
88
.
8
Mayerson
H. S.
1963
.
On lymph and lymphatics.
Circulation
28
:
839
842
.
9
Harvey
N. L.
,
Srinivasan
R. S.
,
Dillard
M. E.
,
Johnson
N. C.
,
Witte
M. H.
,
Boyd
K.
,
Sleeman
M. W.
,
Oliver
G.
.
2005
.
Lymphatic vascular defects promoted by Prox1 haploinsufficiency cause adult-onset obesity.
Nat. Genet.
37
:
1072
1081
.
10
Angeli
V.
,
Ginhoux
F.
,
Llodrà
J.
,
Quemeneur
L.
,
Frenette
P. S.
,
Skobe
M.
,
Jessberger
R.
,
Merad
M.
,
Randolph
G. J.
.
2006
.
B cell-driven lymphangiogenesis in inflamed lymph nodes enhances dendritic cell mobilization.
Immunity
24
:
203
215
.
11
Gunn
M. D.
,
Kyuwa
S.
,
Tam
C.
,
Kakiuchi
T.
,
Matsuzawa
A.
,
Williams
L. T.
,
Nakano
H.
.
1999
.
Mice lacking expression of secondary lymphoid organ chemokine have defects in lymphocyte homing and dendritic cell localization.
J. Exp. Med.
189
:
451
460
.
12
Grubin
C. E.
,
Kovats
S.
,
deRoos
P.
,
Rudensky
A. Y.
.
1997
.
Deficient positive selection of CD4 T cells in mice displaying altered repertoires of MHC class II-bound self-peptides.
Immunity
7
:
197
208
.
13
Lindquist
R. L.
,
Shakhar
G.
,
Dudziak
D.
,
Wardemann
H.
,
Eisenreich
T.
,
Dustin
M. L.
,
Nussenzweig
M. C.
.
2004
.
Visualizing dendritic cell networks in vivo.
Nat. Immunol.
5
:
1243
1250
.
14
Mäkinen
T.
,
Jussila
L.
,
Veikkola
T.
,
Karpanen
T.
,
Kettunen
M. I.
,
Pulkkanen
K. J.
,
Kauppinen
R.
,
Jackson
D. G.
,
Kubo
H.
,
Nishikawa
S.
, et al
.
2001
.
Inhibition of lymphangiogenesis with resulting lymphedema in transgenic mice expressing soluble VEGF receptor-3.
Nat. Med.
7
:
199
205
.
15
Robbiani
D. F.
,
Finch
R. A.
,
Jäger
D.
,
Muller
W. A.
,
Sartorelli
A. C.
,
Randolph
G. J.
.
2000
.
The leukotriene C(4) transporter MRP1 regulates CCL19 (MIP-3beta, ELC)-dependent mobilization of dendritic cells to lymph nodes.
Cell
103
:
757
768
.
16
Itano
A. A.
,
McSorley
S. J.
,
Reinhardt
R. L.
,
Ehst
B. D.
,
Ingulli
E.
,
Rudensky
A. Y.
,
Jenkins
M. K.
.
2003
.
Distinct dendritic cell populations sequentially present antigen to CD4 T cells and stimulate different aspects of cell-mediated immunity.
Immunity
19
:
47
57
.
17
Gautier
E. L.
,
Shay
T.
,
Miller
J.
,
Greter
M.
,
Jakubzick
C.
,
Ivanov
S.
,
Helft
J.
,
Chow
A.
,
Elpek
K. G.
,
Gordonov
S.
, et al
.
2012
.
Gene-expression profiles and transcriptional regulatory pathways that underlie the identity and diversity of mouse tissue macrophages.
Nat. Immunol.
13
:
1118
1128
.
18
Swartz
M. A.
,
Berk
D. A.
,
Jain
R. K.
.
1996
.
Transport in lymphatic capillaries. I. Macroscopic measurements using residence time distribution theory.
Am. J. Physiol.
270
:
H324
H329
.
19
Allan
R. S.
,
Waithman
J.
,
Bedoui
S.
,
Jones
C. M.
,
Villadangos
J. A.
,
Zhan
Y.
,
Lew
A. M.
,
Shortman
K.
,
Heath
W. R.
,
Carbone
F. R.
.
2006
.
Migratory dendritic cells transfer antigen to a lymph node-resident dendritic cell population for efficient CTL priming.
Immunity
25
:
153
162
.
20
Rutkowski
J. M.
,
Markhus
C. E.
,
Gyenge
C. C.
,
Alitalo
K.
,
Wiig
H.
,
Swartz
M. A.
.
2010
.
Dermal collagen and lipid deposition correlate with tissue swelling and hydraulic conductivity in murine primary lymphedema.
Am. J. Pathol.
176
:
1122
1129
.
21
Macatonia
S. E.
,
Knight
S. C.
.
1989
.
Dendritic cells and T cells transfer sensitization for delayed-type hypersensitivity after skin painting with contact sensitizer.
Immunology
66
:
96
99
.
22
Henri
S.
,
Guilliams
M.
,
Poulin
L. F.
,
Tamoutounour
S.
,
Ardouin
L.
,
Dalod
M.
,
Malissen
B.
.
2010
.
Disentangling the complexity of the skin dendritic cell network.
Immunol. Cell Biol.
88
:
366
375
.
23
Rudensky AYu
S.
,
Rath
P.
,
Preston-Hurlburt
D. B.
,
Murphy
,
Janeway
C. A.
 Jr.
1991
.
On the complexity of self.
Nature
353
:
660
662
.
24
Jakubzick
C.
,
Bogunovic
M.
,
Bonito
A. J.
,
Kuan
E. L.
,
Merad
M.
,
Randolph
G. J.
.
2008
.
Lymph-migrating, tissue-derived dendritic cells are minor constituents within steady-state lymph nodes.
J. Exp. Med.
205
:
2839
2850
.
25
Knight
S. C.
2008
.
Specialized perinodal fat fuels and fashions immunity.
Immunity
28
:
135
138
.
26
Neyrolles
O.
,
Hernández-Pando
R.
,
Pietri-Rouxel
F.
,
Fornès
P.
,
Tailleux
L.
,
Barrios Payán
J. A.
,
Pivert
E.
,
Bordat
Y.
,
Aguilar
D.
,
Prévost
M. C.
, et al
.
2006
.
Is adipose tissue a place for Mycobacterium tuberculosis persistence?
PLoS ONE
1
:
e43
27
Nishimura
S.
,
Manabe
I.
,
Nagasaki
M.
,
Eto
K.
,
Yamashita
H.
,
Ohsugi
M.
,
Otsu
M.
,
Hara
K.
,
Ueki
K.
,
Sugiura
S.
, et al
.
2009
.
CD8+ effector T cells contribute to macrophage recruitment and adipose tissue inflammation in obesity.
Nat. Med.
15
:
914
920
.
28
Feuerer
M.
,
Herrero
L.
,
Cipolletta
D.
,
Naaz
A.
,
Wong
J.
,
Nayer
A.
,
Lee
J.
,
Goldfine
A. B.
,
Benoist
C.
,
Shoelson
S.
,
Mathis
D.
.
2009
.
Lean, but not obese, fat is enriched for a unique population of regulatory T cells that affect metabolic parameters.
Nat. Med.
15
:
930
939
.
29
Winer
S.
,
Chan
Y.
,
Paltser
G.
,
Truong
D.
,
Tsui
H.
,
Bahrami
J.
,
Dorfman
R.
,
Wang
Y.
,
Zielenski
J.
,
Mastronardi
F.
, et al
.
2009
.
Normalization of obesity-associated insulin resistance through immunotherapy.
Nat. Med.
15
:
921
929
.
30
Tang
W.
,
Zeve
D.
,
Suh
J. M.
,
Bosnakovski
D.
,
Kyba
M.
,
Hammer
R. E.
,
Tallquist
M. D.
,
Graff
J. M.
.
2008
.
White fat progenitor cells reside in the adipose vasculature.
Science
322
:
583
586
.
31
Vitali
A.
,
Murano
I.
,
Zingaretti
M. C.
,
Frontini
A.
,
Ricquier
D.
,
Cinti
S.
.
2012
.
The adipose organ of obesity-prone C57BL/6J mice is composed of mixed white and brown adipocytes.
J. Lipid Res.
53
:
619
629
.
32
Cypess
A. M.
,
Lehman
S.
,
Williams
G.
,
Tal
I.
,
Rodman
D.
,
Goldfine
A. B.
,
Kuo
F. C.
,
Palmer
E. L.
,
Tseng
Y. H.
,
Doria
A.
, et al
.
2009
.
Identification and importance of brown adipose tissue in adult humans.
N. Engl. J. Med.
360
:
1509
1517
.
33
van Marken Lichtenbelt
W. D.
,
Vanhommerig
J. W.
,
Smulders
N. M.
,
Drossaerts
J. M.
,
Kemerink
G. J.
,
Bouvy
N. D.
,
Schrauwen
P.
,
Teule
G. J.
.
2009
.
Cold-activated brown adipose tissue in healthy men.
N. Engl. J. Med.
360
:
1500
1508
.
34
Saito
M.
,
Okamatsu-Ogura
Y.
,
Matsushita
M.
,
Watanabe
K.
,
Yoneshiro
T.
,
Nio-Kobayashi
J.
,
Iwanaga
T.
,
Miyagawa
M.
,
Kameya
T.
,
Nakada
K.
, et al
.
2009
.
High incidence of metabolically active brown adipose tissue in healthy adult humans: effects of cold exposure and adiposity.
Diabetes
58
:
1526
1531
.
35
Jespersen
N. Z.
,
Larsen
T. J.
,
Peijs
L.
,
Daugaard
S.
,
Homøe
P.
,
Loft
A.
,
de Jong
J.
,
Mathur
N.
,
Cannon
B.
,
Nedergaard
J.
, et al
.
2013
.
A classical brown adipose tissue mRNA signature partly overlaps with brite in the supraclavicular region of adult humans.
Cell Metab.
17
:
798
805
.
36
Nguyen
K. D.
,
Qiu
Y.
,
Cui
X.
,
Goh
Y. P.
,
Mwangi
J.
,
David
T.
,
Mukundan
L.
,
Brombacher
F.
,
Locksley
R. M.
,
Chawla
A.
.
2011
.
Alternatively activated macrophages produce catecholamines to sustain adaptive thermogenesis.
Nature
480
:
104
108
.
37
Van Kruiningen
H. J.
,
Colombel
J. F.
.
2008
.
The forgotten role of lymphangitis in Crohn’s disease.
Gut
57
:
1
4
.
38
Lake
J. E.
,
Currier
J. S.
.
2013
.
Metabolic disease in HIV infection.
Lancet Infect. Dis.
13
:
964
975
.
39
Agarwal, N., D. Iyer, S. G. Patel, R. V. Sekhar, T. M. Phillips, U. Schubert, T. Oplt, E. D. Buras, S. L. Samson, J. Couturier, et al. 2013. HIV-1 Vpr induces adipose dysfunction in vivo through reciprocal effects on PPAR/GR co-regulation. Sci. Transl. Med. 5: 213ra164
.
40
Guaraldi
G.
,
Luzi
K.
,
Bellistrì
G. M.
,
Zona
S.
,
Domingues da Silva
A. R.
,
Bai
F.
,
Garlassi
E.
,
Marchetti
G.
,
Capeau
J.
,
Monforte
Ad.
.
2013
.
CD8 T-cell activation is associated with lipodystrophy and visceral fat accumulation in antiretroviral therapy-treated virologically suppressed HIV-infected patients.
J. Acquir. Immune Defic. Syndr.
64
:
360
366
.
41
Shikuma
C. M.
,
Gangcuangco
L. M.
,
Killebrew
D. A.
,
Libutti
D. E.
,
Chow
D. C.
,
Nakamoto
B. K.
,
Liang
C. Y.
,
Milne
C. I.
,
Ndhlovu
L. C.
,
Barbour
J. D.
, et al
.
2014
.
The role of HIV and monocytes/macrophages in adipose tissue biology.
J. Acquir. Immune Defic. Syndr.
65
:
151
159
.
42
Jackson
D. G.
2009
.
Immunological functions of hyaluronan and its receptors in the lymphatics.
Immunol. Rev.
230
:
216
231
.
43
Mäkinen
T.
,
Adams
R. H.
,
Bailey
J.
,
Lu
Q.
,
Ziemiecki
A.
,
Alitalo
K.
,
Klein
R.
,
Wilkinson
G. A.
.
2005
.
PDZ interaction site in ephrinB2 is required for the remodeling of lymphatic vasculature.
Genes Dev.
19
:
397
410
.
44
Muto
J.
,
Morioka
Y.
,
Yamasaki
K.
,
Kim
M.
,
Garcia
A.
,
Carlin
A. F.
,
Varki
A.
,
Gallo
R. L.
.
2014
.
Hyaluronan digestion controls DC migration from the skin.
J. Clin. Invest.
124
:
1309
1319
.
45
MartIn-Fontecha
A.
,
Sebastiani
S.
,
Höpken
U. E.
,
Uguccioni
M.
,
Lipp
M.
,
Lanzavecchia
A.
,
Sallusto
F.
.
2003
.
Regulation of dendritic cell migration to the draining lymph node: impact on T lymphocyte traffic and priming.
J. Exp. Med.
198
:
615
621
.

The authors have no financial conflicts of interest.

Supplementary data