Elastin Shapes Small Molecule Distribution in Lymph Node Conduits

Activation of T cells or B cells occurs in distinct anatomical locations inside the lymph nodes (LNs) (1–7). Lymph-borne Ags stimulate LN resident APCs before tissue-originated Ag-bearing dendritic cells (DCs) enter the LNs (8). To precisely deliver Ags to specific locations, it is critical to understand how LNs shape spatial and temporal Ag distribution (7–11). The highest concentration of Ag is found in LN sinuses, where Ags are sampled by sinus-associated macrophages (12–16) and DCs (9). Only small molecules (molecular mass <70 kDa) can reach deeper into the LN parenchyma, such as the B cell or T cell zones, through LN conduits (17–20). The LN conduit network connects LN sinuses with the LN vasculature, including lymphatic endothelial cells and high endothelial venules (HEVs) (10, 21). Lymph-derived regulatory factors, such as cytokines, can use LN conduits to rapidly reach HEVs and regulate the HEV gene expression profile for LN cell homeostasis (10, 21–23). Transmission electron microscopy (TEM) images show that the core of LN conduits is composed of solid collagen fibers (10, 23), whereas fibroblastic reticular cells (FRCs) wrap around the collagen core (21, 24–26). Recently, it has been shown that the blood vessel plasmalemma vesicle–associated protein (PLVAP) is expressed on LN sinus lymphatic endothelial cells and is able to exclude large molecules from entering LN conduits (27). However, the exact mechanisms controlling the flow of fluid and small molecules in the LN remain largely unclear.

Elastin is one of the components of LN conduits (28). It is a major hydrophobic protein that provides an elastic property, allowing tissues to restore their shape after stretching and distortion (29). The absolute content of elastin in LN conduits is relatively low compared with collagens I and III, and the function of elastin is thought to provide elasticity to LNs (28, 30). The elastic properties of elastin are critical for the function of initial lymphatic vessels (29, 31, 32). The initial lymphatic vessels connect to the surrounding tissues by anchoring filaments, which link to elastin fibers in the skin. The lymphatic endothelial leaflets between the discontinuous button-like intercellular junctions become the initial lymphatic valve, which opens when interstitial fluid pressure increases (33). Elastin is believed to provide the initial lymphatic valve anchoring and the resilience of initial lymphatic valve movements (29, 31, 32). Although it is clear that elastin provides the elasticity for lymphatic vessels or LN stretching, whether elastin plays any other role in the regulation of lymph flow remains unclear.

In this study, using fluorescent molecule FITC as a tracer, we showed that lymph preferentially flows along LN capsule–associated (LNC) conduits. Elastin fibers are the only component that colocalized with high-intensity FITC or tetramethylrhodamine isothiocyanate (TRITC) inside LN conduits. Although fluorescent Alexa-555-labeled OVA (Alexa555-OVA) alone does not enter LN conduits as effectively as FITC, coinjection of LPS with Alexa555-OVA substantially enhances OVA accumulation along elastin fibers in LNC conduits. Finally, FITC is also highly concentrated along elastin fibers in skin lymphatic vessels. Our studies suggest that elastin fibers have a novel function in concentrating (or mediating) small-molecule trafficking to lymphatic vessels in the skin and in LN conduits. Although the molecular mechanisms remain unclear, our results suggest that Ags with high affinity to elastin fibers may be more effectively delivered to lymphatic vessels and LNs.

C57BL/6 mice were purchased from Jackson Laboratory; Prox-mOrange mice were a kind gift from Dr. F. Kiefer (34) and were bred at the Health Sciences Animal Resource Center at the University of Calgary. All the experiments were performed using 6- to 8-wk-old female mice. All animal protocols were reviewed and approved by the University of Calgary Animal Care and Ethics Committee and conformed to the guidelines established by the Canadian Council on Animal Care.

FITC (Sigma) was prepared in a 1:1 (v/v) acetone/dibutyl phthalate mixture (2%). To determine FITC distribution in inguinal LNs, 200 μl of FITC was applied on shaved skin on the flanks. To determine FITC distribution in the popliteal LN (PLN), 50 μl of FITC was applied on the dorsal side of the footpad and shaved leg. To determine FITC distribution in the ear skin, 25 μl of FITC was applied on the ear.

TRITC in DMSO (50 μl) at 1 mg/ml (both Sigma) was injected in the flanks. Inguinal LNs were collected 2 h later for analysis. Alexa-555–conjugated OVA (Alexa555-OVA) (Thermo Fisher) was diluted in saline and injected intradermally in the right side of flank at 20 μg in 50 μl saline or at the footpad at 20 μg in 20 μl saline. When OVA was coinjected with LPS, 20 μg of Alexa555-OVA and 12 ng of LPS (Sigma) were mixed in 20 μl saline and injected at the footpad.

Tissues (LNs and ears) of interest were harvested from euthanized mice. Tissues were incubated in 4% formaldehyde for whole-amount staining or were frozen directly in OCT for frozen sections.

LNs were incubated twice in a 50% 1:2 mixture of benzyl alcohol and benzyl benzoate (BABB) in methanol and were incubated two to three times in 100% BABB. LNs were then kept in a microwell with 100% BABB on the slide, and coverslips were mounted on the microwell for imaging.

The immunofluorescent staining used standard protocol. Briefly, 10–20-μm frozen sections were blocked with 5% mouse serum for 1 h. Samples were incubated overnight with the primary Abs. After two to three washings in PBS, samples were incubated with fluorescent probe–conjugated secondary Abs (Jackson ImmunoResearch) for 60–90 min. The primary Abs were rabbit anti–Lyve-1 (Cell Sciences, aa 24–228), ER-TR7 (antireticular fibroblasts and reticular fibers Ab, clone ER-TR7, ab51824; Abcam), rabbit anti–collagen I (ab34710; Abcam), rabbit anti–collagen IV (AB8201; Millipore), rabbit anti-fibronectin (ab2413; Abcam), rat anti-B220 (RA3-6B2; BD Biosciences), rat anti-CD31 (MEC13.3; BD Biosciences), and MECA 32 (anti-mouse panendothelial cell Ag Ab, clone MECA 32; BioLegend). H&E staining was performed following the manufacturer’s standard protocol (VWR) using 5-μm paraffin sections.

Mice were anesthetized with Ketamine/Xylazine. The footpad was then injected with 10 μl FITC. A small cut was made at the skin to expose the PLN by removing the adipose tissue surrounding it. Each mouse was then fixed on the heating stage and time-lapse imaging was performed using an SP8 multiphoton microscope with 25× water objective.

Fluorescent samples were imaged with an SP8 multiphoton microscope (Leica) for whole-mount imaging and three-dimensional reconstruction or an SP8 confocal microscope (Leica). The objectives used were 20× air, 25× water, or 63× oil, depending on the study. For the high-resolution images, the slice thickness was 0.2 μm with a pinhole at 0.5 when using the 63× oil objective. The resolution was ∼300 nm with these settings.

For normal LNs, the second harmonic generation (SHG) signal was imaged using a 1050-nm laser with multiphoton microscopy (MP) because the longer wavelength permits imaging deeper into the LN. The FITC signal was detected using a 900-nm MP laser. The three-dimensional reconstruction and the diameter measurement were performed with the Leica LAX software. Some of the analysis and image brightness adjustment were performed with ImageJ.

Structured illumination superresolution microscopy was performed with a Zeiss ELYRA PS.1 microscope with 0.1-μm steps in the z-axis. A 28-μm grid was used for the 488 laser, whereas a 34-μm grid was used for the 561 and 642 lasers. Each channel was acquired using five rotations using a Plan-Apochromat 63×/1.4 M27 objective. Raw images were reconstructed using Zeiss ZEN 2012 black microscope (Zeiss, Germany). The resolution was 130 (lateral) and 300 nm (axial).

The tissue samples were prefixed with 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) for at least 2 h at 4°C. After washing three times with the same buffer, the samples were postfixed with cacodylate-buffered 1% osmium tetroxide for 2 h at room temperature and then dehydrated through a graded series of acetone and embedded in EPON resin. Ultrathin sections were cut on a Leica EM UC7 ultramicrotome using a diamond knife and were collected on single-hole grids with Formvar supporting film. The sections were stained with 2% aqueous uranyl acetate and Reynolds’s lead citrate and observed in a Hitachi H7650 TEM operated at 80 kV. The images were taken through an AMT16000 digital camera mounted on the microscope.

To better understand how lymph flows inside the LN conduits, we used FITC to trace lymph distribution in the inguinal LN. FITC was sensitized on the skin of flanks, and inguinal LNs were collected 2 h later. As expected, cryosections with anti-B220 and anti–collagen I staining showed that FITC is concentrated in the LN subcapsular sinus (SCS), in the medullary sinus (MS), and along LNC conduits (Fig. 1A). LNC conduits are positioned in the interfollicular zone (area between B cell zones) and in the area where the SCS is close to the MS (SCS–MS transition area) (Fig. 1A, boxes).

Anatomically, LNC conduits [previously described by less defined terms such as “sinus strands” (27) or “the trabeculae”] provide a physical connection between the LN capsule and LN parenchyma. Using anti–collagen I staining on inguinal LNs collected from untreated mice, the average diameter of the LNC conduits in the SCS and interfollicular area was 2–3 μm, with some even reaching 3–4 μm (Fig. 1B). The diameters of LNC conduits were larger than the average diameter of the LN conduits in the T and B cell zones (1–2 μm). In the SCS, the distance between the LNC conduits varied from several micrometers to 30–40 μm, which remained unchanged within 2 h after FITC sensitization (Fig. 1C). To exclude potential bias in image quantification from two-dimensional images, we also imaged untreated whole-mount inguinal LNs with MP microscopy using SHG. Three-dimensionally reconstructed SHG images showed two types of LN conduits: the large LNC conduits initiated from the LN capsule (Fig. 1D) and the interconnected conduit network that required much stronger MP laser power to be detected (Fig. 1E). To determine if FITC only colocalized with the LNC conduits or actually flowed along the LNC conduits, we performed time-lapse live imaging to trace FITC flow direction in the PLN. Time-lapse video also showed FITC flowing along LNC conduits (Supplemental Video 1). The morphology, size, and density of LNC conduits suggest that they contribute to the flow of large volumes in the LN.

To determine how FITC flows inside LNC conduits, we used TEM to define the ultrastructure of LNC conduits in the inguinal LN. TEM images showed that LNC conduits also had a core of collagen fibers surrounded by fibroblasts (Fig. 1F, 1G). This ultrastructure of LNC is similar to the previously reported LN conduits (23). Despite LNC conduits being larger in diameter, the actual pore size of the LNC conduits and the surrounding fibroblasts did not appear to permit large molecules into the LNC conduits. Consistent with previous reports, larger m.w. tracers (2 M FITC-dextran) cannot enter LNC conduits (data not shown).

To understand how FITC distributes inside LNC conduits, we used anti–collagen I staining to mark LN conduits. Superresolution microscopy showed that FITC did not evenly distribute inside LNC conduits. Instead, it appears that within the low intensity FITC-filled lumen of the LNC conduits (Fig. 1H, 1I stars), FITC was preferentially concentrated along several cords (FITC^intense cords) in the LNC conduits (Fig. 1H, 1I, arrows). However, FITC^intense cords did not colocalize with anti–collagen I staining. FITC^intense cords in the lumen of LNC conduits were also observed using other conduit markers, such as laminin (Fig. 1J–M). Together, our fluorescent images identified previously unknown FITC^intense cords in LNC conduits.

To determine the nature of the components associated with the FITC^intense cords, we examined different LN conduit markers, including collagen I (Fig. 1H, 1I), laminin (Fig. 1J–M), collagen IV, fibronectin, antireticular fibroblasts and reticular fibers Ab (ER-TR7), podoplanin, and elastin (Supplemental Fig. 1A–G). We also looked at blood endothelial cell markers (CD31 and MECA32) and lymphatic endothelial cell markers (Lyve-1 and Prox-1) (Supplemental Fig. 1H–N). Among all markers, only elastin was colocalized with FITC^intense cords (Fig. 2A–D). Elastin fibers were not limited to LNC conduits (Fig. 2E–H) but instead appeared to be larger (∼500 nm, Fig. 2B, 2D, arrows) compared with the other LN conduits (<200 nm) (Fig. 2B, 2D). However, our image resolution limits accurate measurements of the elastin fiber diameter. Consistent with the size of elastin fibers, the diameters of the FITC^intense cords were also larger inside LNC conduits compared with the rest of the conduits. Three-dimensionally reconstructed whole-mount LNs further corroborated this observation and showed that FITC preferentially located in LNC conduits (Fig. 2I). FITC distribution along LNC conduits was also seen when imaging PLNs 2 h after FITC sensitization on leg skin (Supplemental Video 2). To determine if the colocalization of FITC^intense cords with elastin fibers is unique to FITC, we next used TRITC as alternative fluorescecent tracer. Results showed that TRITC was also well colocalized along elastin fibers inside the LNC conduits in inguinal LNs (Fig. 2J–M). Thus, both FITC and TRITC can effectively enter the conduits and colocalize with elastin fibers. The distribution pattern is consistent in LNs positioned in different anatomical locations, such as in the inguinal LNs or PLNs.

Next, we used fluorescent dye Alexa555-OVA to study how OVA is distributed in LNC conduits. Compared with FITC or TRITC, Alexa555-OVA from intradermal injection at the flanks cannot effectively enter LNC conduits (Fig. 3A). To determine if an injection of Alexa555-OVA into a different anatomical location may enhance Alexa555-OVA entry into the LNC conduits, we injected OVA into the footpad where lymph is well known to be restricted to the PLN. In 2 out of 10 PLNs we analyzed, a small number of Alexa555-OVA⁺ LNC conduits were observed by using whole-mount LN imaging (Fig. 3B).

It is well known that LPS substantially increases immune response to OVA by activating APCs through TLR 4 (35). We hypothesize that LPS-induced immune stimulation may help OVA enter the LNC conduits and travel along elastin fibers. To test this idea, we injected a mixture of Alexa555-OVA with LPS at the footpad and collected the PLNs 2 h after the injection. LPS substantially increased the number of Alexa555-OVA⁺ LNC conduits (Fig. 3C, 3D). Anti-elastin staining showed that Alexa555-OVA was colocalized with elastin in LNC conduits (Fig. 3E–H). Thus, Alexa555-OVA alone does not effectively enter the conduits, but the addition of LPS enhances the distribution of Alexa555-OVA along elastin fibers in LNC conduits.

The observed colocalization of FITC with elastin fibers in LN conduits raises a new question: do elastin fibers in the skin also contribute to lymph flow? It has been previously suggested that elastin provides an elastic connection between tissue and the initial lymphatic vessels and helps initial lymphatic vessel valve motion (36). Using an untreated ear as a control, we adjusted imaging settings to avoid green autofluorescence (Fig. 4A). Lymphatic vessels and blood vessels were labeled with anti–Lyve-1 and MECA 32 Abs, respectively (Fig. 4A). To determine the relationship between elastin, lymphatic vessels, and blood vessels in ear skin, we used immunofluorescent staining with anti-elastin, anti–Lyve-1, and MECA32 Abs. Elastin preferentially surrounded Lyve-1⁺ lymphatic vessels rather than MECA 32⁺ blood vessels in untreated ear skin (Fig. 4B).

In skin collected 2 h after FITC sensitization, FITC had diffused into the skin (Fig. 4C, 4D). FITC^intense cords were observed to surround the Lyve-1⁺ lymphatic vessels rather than the MECA32⁺ blood vessels in ear skin (Fig. 4D, white arrows). This pattern was better visualized using higher magnification images: FITC^intense cords were colocalized with elastin, which was highly concentrated around Lyve-1⁺ lymphatic vessels but not MECA32⁺ blood vessels (Fig. 4E). Thus, FITC is concentrated along elastin fibers in the skin and preferentially connects to the lymphatic vessels. This pattern indicates the novel role of elastin in guiding interstitial fluid flow toward lymphatic vessels (Fig. 5).

Lymphatic vessels transport Ag-activated DCs from tissue to the draining LN to initiate a protective immunity. Additionally, Ags can travel with lymph flow to the LN and stimulate LN resident APCs hours before tissue-originated Ag-bearing DCs enter the LNs (8). In fact, the LN resident APCs are sufficient to generate a protective immune response to the invading pathogens (9, 37, 38). The microarchitecture of the LN is highly organized. It can be divided into the T cell zone, B cell zone, interfollicular zone (area between the B cell zone), cortical ridge (area between T cell and B cell zone), and areas proximal to the SCS or MS. Accordingly, different subsets of LN resident APCs locate at different locations in the LN. The importance of LN resident APC activation by the soluble Ags is demonstrated by the T cell and B cell activation, which occurs in various anatomic locations in the LN. The interfollicular area and the area close to the sinus support Th1 cell differentiation or memory T cell recall responses (1, 2). Rapid Ag delivery and high concentrations of Ag accumulation in the interfollicular and medullary zone are essential for B cell activation or cell egress from the LN (5, 6). Recently, it was reported that an Ag gradient in the LN causes different subsets of LN resident DCs to activate and prime CD4⁺ or CD8⁺ T cell responses at different locations in the LN (4). Thus, the spatial and temporal Ag distribution determines the types of immune responses at the different locations in the LN. However, the mechanism that controls Ag distribution remains largely unclear.

LN conduits provide a highway to help fluid and small m.w. Ags travel faster and deeper into the LN to activate conduit-associated LN resident APCs (17–20). How Ags travel along the conduits remains unclear. In this study, by characterizing the fluid distribution pattern and analyzing most of the documented conduit components, we found that fluid and small molecules are preferentially colocalized with elastin fibers in the LN conduits (Fig. 5). The colocalization of FITC^intense cords or TRITC^intense cords with elastin fibers was consistently observed in almost all LNC conduits (Fig. 5, insert). However, when OVA (molecular mass ∼45 kDa) was injected alone, the number of LNC conduits with Alexa555-OVA^intense cords was far less. It could be that FITC and TRITC, being much smaller molecules (m.w. ∼400), better represent fluid flow. Another possibility is that the hydrophobic characteristic of elastin may have a higher affinity for FITC or TRITC. Importantly, injection of Alexa555-OVA with LPS substantially enhances OVA distribution along elastin in LNC conduits. Therefore, both the biochemical properties of Ags and immune stimulation are important for Ag distribution. The underlying mechanism remains unclear. It is possible that LPS has activated DCs or FRCs to enhance OVA entrance into the LNC conduits. Additionally, because LPS is composed of hydrophilic polysaccharides and a hydrophobic component lipid A, it is possible that the hydrophobic portion of LPS helps OVA enter the LNC conduits and move along elastin fibers. Future studies will be carried out to investigate the biochemistry of Ags and what type of adjuvant to combine to improve their trafficking along elastin, which may improve the efficiency of Ag delivery or help deliver Ags to designated locations in the LN. In contrast, a better understanding of the biochemical properties of elastin fibers will provide a new entry point to study the regulation of Ag distribution.

Elastin has been previously identified as a component of the LN conduits, but very little attention has been given to elastin compared with other conduit components (28, 30). It was previously suggested that elastin may act as a low-resistance path, which guides Langerhans cell migration from the epidermis to the lymphatic vessels (36). In this study, we found that FITC^intense cords are colocalized with elastin fibers in the skin, which preferentially surround lymphatic vessels rather than blood vessels (Fig. 5). This pattern suggests that elastin fibers also favor the entrance of FITC-containing interstitial fluid into lymphatic vessels. In LNs, the FITC signal is much more intense along LNC conduits compared with the diffusion signal. Morphological images show that FITC^intense cords are colocalized with elastin fibers inside LNC conduits (Fig. 5, box). Although our image resolution did not allow for a precise measurement, we can estimate the diameter of elastin fibers in LNC conduits to be ∼500 nm, which is larger than the other LN conduits in the LN (<200 nm). Although LNC conduits may contain several elastin fibers with different sizes, small conduits only contain one small-size elastin fiber. Consequently, more FITC accumulates along the LNC conduits.

Although the diameter of LNC conduits is larger than the rest of the conduits, LNC conduits are also composed of solid fibers with surrounding FRCs (10, 19, 23, 26). Therefore, the pore size of the LNC conduits remains very small. A previous report determined that PLVAP (recognized by MECA32 Ab) expressed by sinus lymphatic endothelial cells controlled the size of materials entering LN conduits (27). Similarly, we also found LNC conduits ensheathed by endothelial cells. Therefore, despite having a larger diameter, LNC conduits do not allow entry of larger materials (data not shown).

While characterizing the LNC conduit components, anti–collagen I staining showed that collagen I was surrounding LNC conduits rather than forming its core. The pattern of collagen I surrounding LNC conduits was consistent with the staining patterns of several extracellular matrix proteins (collagen IV, fibronectin, and laminin). Only elastin was found to be colocalized with FITC^intense cords inside the LNC conduits. Thus, the resolution of our images was sufficient to distinguish the signal in the lumen of the LNC conduit and those surrounding the LNC conduits. We cannot determine if this observation was because the Ab we used could only recognize the collagen I surrounding the conduits, or if it is because collagen I was not the major component of the core fibers shown in TEM. If the latter is correct, because the sizes and the number of elastin fibers do not make up the entire core component of LNC conduits as seen in TEM images, other components of the LN conduit core remain to be clarified. Whether other conduit components may be better associated with different types of Ags remains to be determined.

In summary, our studies show that elastin not only provides tissue elasticity but is also involved in fluid and Ag distribution in the skin and LNs. Although the mechanism remains to be explored, our studies suggest that future studies on deciphering the mechanism of Ag trafficking along elastin fibers, and on potentially improving it, may be a new strategy to direct Ag distribution. This could be especially useful to generate designated T or B cell responses in the LN during vaccine design.

We thank Dr. Pierre-Yves von der Weid for his critical inputs during this study and the manuscript preparation.

The authors have no financial conflicts of interest.