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Molecular and Cellular Biology, January 2007, p. 595-604, Vol. 27, No. 2
0270-7306/07/$08.00+0     doi:10.1128/MCB.01503-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Normal Lymphatic Development and Function in Mice Deficient for the Lymphatic Hyaluronan Receptor LYVE-1{triangledown} ,{dagger}

Nicholas W. Gale,1,{ddagger} Remko Prevo,2,{ddagger} Jorge Espinosa,2 David J. Ferguson,3 Melissa G. Dominguez,1 George D. Yancopoulos,1 Gavin Thurston,1 and David G. Jackson2*

Regeneron Pharmaceuticals, Inc., 777 Old Saw Mill River Road, Tarrytown, New York 10591,1 MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, Headington, Oxford OX3 9DS, United Kingdom,2 Nuffield Department of Pathology, John Radcliffe Hospital, Headington, Oxford OX3 9DS, United Kingdom3

Received 12 August 2006/ Returned for modification 19 September 2006/ Accepted 28 October 2006


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ABSTRACT
 
The hyaluronan receptor LYVE-1 is expressed abundantly on the surfaces of lymphatic vessels and lymph node sinus endothelial cells from early development, where it has been suggested to function both in cell adhesion/transmigration and as a scavenger for hyaluronan turnover. To investigate the physiological role(s) of LYVE-1, we generated mice in which the gene for the receptor was inactivated by replacement with a ß-galactosidase reporter. LYVE-1–/– mice displayed an apparently normal phenotype, with no obvious alteration in lymphatic vessel ultrastructure or function and no apparent change in secondary lymphoid tissue structure or cellularity. In addition, the levels of hyaluronan in tissue and blood were unchanged. LYVE-1–/– mice also displayed normal trafficking of cutaneous CD11c+ dendritic cells to draining lymph nodes via afferent lymphatics and normal resolution of oxazolone-induced skin inflammation. Finally, LYVE-1–/– mice supported normal growth of transplanted B16F10 melanomas and Lewis lung carcinomas. These results indicate that LYVE-1 is not obligatory for normal lymphatic development and function and suggest either the existence of compensatory receptors or a role more specific than that previously envisaged.


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INTRODUCTION
 
The lymphatic vasculature represents a second circulatory system that plays an important role in the maintenance of tissue fluid homeostasis (16). In addition, the network of vessels and lymph nodes that constitute the lymphatic system represents the main core of the immune system. For example, it is the afferent lymphatic vessels that act as vital conduits for antigen-presenting dendritic cells migrating from the tissues to the draining lymph nodes, where they stimulate naïve T cells to mount the primary immune response (30). These same vessels also provide routes for tumor metastasis; early colonization of the lymph nodes is a common finding for many common human cancers (26). However, unlike the blood vasculature, the mechanisms that regulate cellular recognition and trafficking within the lymphatic vasculature, including the complex sinus structures of the lymph nodes, are poorly understood.

Many details about the development of the lymphatics have become known only within the past decade, largely as a result of studies of gene-targeted mice (reviewed in reference 28). For mammals, it has been demonstrated that the lymphatics develop centrifugally by budding from the thoracic vessels (32), confirming predictions made over a century ago that they have a venous origin (24; reviewed in reference 19). During mouse embryogenesis, the first lymph sacs bud from the cardinal vein in response to the lymphangiogenic growth factor vascular endothelial growth factor C (VEGF-C) and its lymphatic endothelium-specific receptor VEGFR-3 around embryonic day 10.5 (E10.5), and these subsequently give rise to the developing vessel network in response to continuing cues from the homeobox domain-related transcriptional factor PROX-1 (31, 32). Patterning and interconnection of lymphatic vessels later in development and during postnatal life are regulated by the growth factor Ang-2 and its TIE-2 receptor on lymphatic endothelium (5) and by Eph-EphrinB2 interactions (15), while arterialization of larger lymphatic collecting vessels and valve formation is regulated by the transcription factor FoxC2 in combination with platelet-derived growth factor/platelet-derived growth factor receptor signaling (12, 21). Other proteins of lymphatic endothelial cells that contribute to lymphatic vessel function include the class III semaphorin receptor neuropilin 2 (33), the integrin {alpha}9ß1 (6), and the 38-kDa integral membrane mucoprotein podoplanin/T1alpha, deletion of which results variously in embryonic lethality and/or fluid leakage syndromes of chylothorax and chylous ascites (25). Finally, the lymphatic endothelial chemokine receptor D6 appears to be important for clearance of chemoattractants during the resolution phase of inflammation (11).

Another interesting and particularly abundant component of lymphatic endothelium is the transmembrane protein LYVE-1, a member of the Link protein superfamily (1, 22) that binds the large extracellular matrix glycosaminoglycan hyaluronan [HA; (GlcNAcß1-4GlcUA)n] via a conserved C-type-lectin-like Link domain (3). Homologous to the leukocyte homing receptor CD44, which supports HA-mediated rolling on inflamed hemovascular endothelium (23), LYVE-1 begins to express almost simultaneously with PROX-1 during development, and this expression thereafter remains almost exclusively confined to lymphatic vessel and lymph node sinus endothelium (19, 20). These latter properties have made LYVE-1 a powerful and widely exploited molecular marker in studies of normal and pathological lymphangiogenesis (9). Nevertheless, the physiological function of LYVE-1 has remained unresolved (7). The known involvement of the lymphatics and lymph nodes in particular in the clearance and degradation of hyaluronan (4, 13), allied with the observed capacity of LYVE-1-transfected cells to internalize hyaluronan (22), would tend to suggest a role in glycosaminoglycan homeostasis (8). Furthermore, its structural similarity to CD44 and the observed capacity to form trimolecular complexes with CD44 and hyaluronan in vitro suggest a role for LYVE-1 in mediating the transmigration of CD44-expressing leukocytes across lymphatic vessels (7). Definitive evidence in favor of either role, however, is lacking.

In a more direct approach to elucidating the true physiological function of LYVE-1, we have generated mice that lack the LYVE-1 gene by targeted replacement with a ß-galactosidase (ß-Gal) reporter. Here we report that LYVE-1 gene-targeted mice develop normally and establish a functional network of lymphatic vessels and lymph nodes that is indistinguishable from that of wild-type animals. We have found no evidence for any disruption in the metabolism of hyaluronan or in the development or compartmentalization of leukocyte subsets and no defect in dendritic cell trafficking or tumor growth. This lack of an obvious phenotype argues against a major structural, developmental, or regulatory role for LYVE-1 and suggests either compensation by an as-yet-unidentified component or a role much more specialized than that previously envisaged.


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MATERIALS AND METHODS
 
Generation of LYVE-1 gene-targeted mice. Gene targeting of the LYVE-1 locus was accomplished using VelociGene technology and was initially reported in a summary table (29). Briefly, a bacterial artificial chromosome (BAC)-based targeting vector was derived using short regions of homology flanking exons 2 and 5 (termed 5' and 3' homology boxes, respectively) (Fig. 1A). These homology boxes were ligated to a reporter/selection cassette encoding transmembrane ß-Gal (TM-LacZ) and a loxP-flanked neomycin resistance selection gene. Homologous recombination in bacteria through these homology boxes was used to replace the intervening sequences in a BAC containing the complete LYVE-1 gene. The replacement leads to the deletion of exons 2 to 5 and intervening introns and is designed, after splicing, to result in the fusion of the first 32 amino acids of LYVE-1 (containing the signal peptide) to TM-LacZ. The BAC employed in the construction was clone 343p22 (Incyte Genomics), which is approximately 150 kb in size. The cassette replacement results in a targeting vector with homology arms of 95 kb and 7 kb and a deletion of approximately 7 kb (Fig. 1A). This BAC-based targeting vector was used as described previously (29) to target the LYVE-1 locus in F1H4 hybrid embryonic stem (ES) cells. Several TaqMan probes, termed "loss-of-allele" probes, were designed to the deleted region and were used to establish that proper targeting had occurred in ES cells and subsequently in mice as described previously (29) (Fig. 1A). Two independent ES cell clones were derived and tested, and both clones behaved identically in their abilities to generate chimeric F1 and F2 mice. Knockout mice were backcrossed one additional generation onto the C57BL/6 background, resulting in an approximate 75% contribution of the C57BL/6 and 25% of the 129 backgrounds in the resulting N2F2 mice for studies reported here. Subsequent backcrossing onto the C57BL/6 background for more than five generations has produced no apparent change in phenotype. All procedures were carried out according to relevant U.S. federal or United Kingdom Home Office guidelines and institutional procedures as appropriate.


Figure 1
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  		FIG. 1. Generation of LYVE-1 gene-targeted mice. (A) The native LYVE-1 locus (top) showing the intron-exon structure, location of loss-of-allele probes (red circles), and 3' and 5' homology boxes (purple boxes) (see Materials and Methods). The targeting cassette (middle) shows the reporter/selection replacement cassette comprised of a TM-LacZ gene and a neomycin selection gene. Synthetic 3' and 5' homology boxes (purple) ligated to the reporter/selection cassette direct its targeting to the appropriate location within the LYVE-1 gene by homologous recombination in bacteria. The final targeted allele, achieved by homologous recombination in ES cells (bottom), in which exons 2 and 5 and the intervening sequences are precisely replaced with the targeting cassette, is shown. The normal LYVE-1 gene is ablated in the targeted allele, and splicing of the targeted allele leads to the expression of mRNA containing a transmembrane ß-Gal reporter gene. (B to E) X-Gal staining of whole-mount (B and D) sections of intestine whole mounts (viewed from the serosal surface) from heterozygous mice reveals ß-Gal expression in cells in a pattern identical to that for whole mounts stained with anti-LYVE-1 (C and E). Panels B and C show ß-Gal versus immunohistochemical staining in similar regions of normal intestinal serosa; panels D and E show regions of intestine harboring Peyer's patches. (F) Compared to that with wild-type controls, whole-mount immunohistochemistry of intestine (viewed from the mucosal surface) with anti-LYVE-1 antisera reveals a complete absence of LYVE-1 expression in knockout (KO) mice.

Whole-mount histochemistry and immunohistology. Whole-mount staining for ß-Gal in LYVE-1+/– and LYVE-1–/– mice and tumor cryostat sections was performed as described previously (2). For whole-mount preparations of dermal lymphatics, ear tissues were fixed overnight in 4% paraformaldehyde in phosphate-buffered saline (PBS), and the dorsal segment was blocked for 3 h in 30 mg/ml milk powder-10% fetal bovine serum (FBS) in PBS-0.1% (wt/vol) Triton X-100 prior to double staining overnight with rat anti-mouse CD31 (MEC 13.3)/Alexa594-conjugated goat anti-rat and hamster anti-mouse podoplanin (8.1.1)/Alexa488-conjugated goat anti-hamster antibodies. Preparation of intestinal whole mounts and immunostaining for lymphatics was performed essentially as described in reference 5 by use of antisera generated against mouse LYVE-1-human immunoglobulin G1 (IgG1)-Fc (1/10,000 dilution) or a directly biotinylated goat anti-mouse VEGFR-3 antiserum (R&D Systems). Fluorescent antibody-stained sections were viewed with a Bio-Rad Radiance 2000 laser scanning confocal microscope equipped with argon and green helium/neon lasers and analyzed using LaserSharp2000 software. Images were recorded in simultaneous scanning mode with lambda strobing at 4-µm intervals over a depth of 24 µm using a focusing motor and subsequently merged.

Immunohistology of cryostat sections. Tissues were embedded in OCT mounting medium (Tissue-Tek) and frozen in liquid N2 or in dry ice-ethanol and cut into 10-µm sections for fixation in acetone. Sections were stained with hamster anti-mouse podoplanin (clone 8.1.1; Developmental Studies Hybridoma Bank, IA), goat anti-mouse VEGFR-3 (R&D Systems), rat anti-peripheral lymph node addressin (PNAd), clone MECA79 (Becton Dickinson), or biotinylated hyaluronan binding protein (bHABP) (Seikagaku). After being washed, slides were incubated with the appropriate secondary antibodies: Alexa594-conjugated goat anti-hamster, Alexa594-conjugated donkey anti-goat, Alexa488-conjugated streptavidin (all from Molecular Probes), or fluorescein isothiocyanate (FITC)-conjugated goat anti-rat IgM (Southern Biotech). Then, viewing was performed with a Zeiss Axiophot microscope with epifluorescent illumination.

Electron microscopy. Samples of formaldehyde-fixed skin and intestine were embedded in Spurr's epoxy resin, and thin sections were coated on Formvar-coated gold grids prior to staining with uranyl acetate and lead citrate. Preparations were examined with a JEOL 1200EX electron microscope.

FITC-dextran and Evans blue dye lymphangiography. Fluorescence microlymphangiography of the tail was carried out essentially as described in reference 27. Mice were anesthetized with ketamine (80 µg/g body weight) and medetomidine (Domitol; 1 µg/g), placed on a heating pad, and injected with a solution of 10 mg/ml FITC-labeled dextran (2,000 kDa; Sigma) in PBS administered over a period of 20 min through a 29-gauge 1/2-in. needle inserted intradermally in the tail tip. Dye uptake was monitored continuously using a Zeiss inverted epifluorescence microscope.

For Evans blue dye lymphangiography, 40 µl of 1% (wt/vol) Evans blue was injected into hind-limb footpads of mice that were first anesthetized with isoflurane. After 20 min, mice were perfused with PBS and then photographed.

Measurement of tissue hyaluronan levels. Levels of hyaluronan extracted from mouse tissues were estimated using a competitive enzyme-linked immunosorbent assay (ELISA). For the extraction step, tissue samples were digested with papain (250 µg/ml) in 5 mM cysteine-5 mM EDTA, pH 7.5, at 60°C for 24 h, followed by inactivation of the enzyme (100°C, 10 min) and centrifugation (900 x g, 5 min) to remove particulate matter. For the ELISA, 96-well Nunc Maxisorp plates were coated overnight (4°C) with 25 µg/ml rooster comb hyaluronan (Sigma) in coating buffer (15 mM sodium carbonate and 34 mM sodium bicarbonate, pH 9.4) followed by blocking in bovine serum albumin (10 mg/ml in PBS, pH 7.5) for 1 h at 37°C. Tissue samples (50 µl) or hyaluronan standards (12 to 1,600 ng/ml) were then mixed with biotinylated bHABP (50 µl, 1 mg/ml; Seikagaku) and incubated for 3 h at 37°C prior to washing and detection with peroxidase-conjugated streptavidin (1:500 DAKO) followed by O-phenylenediamine substrate (Sigma) and measurement of absorbance at 490 nm in a Bio-Rad plate reader.

Determination of serum Ig levels. Levels of serum Ig classes were estimated by antigen capture ELISA. Briefly, ELISA plates (Maxisorp; Nunc) were coated overnight (2 µg/ml) with either rat anti-mouse IgA, rat anti-mouse IgG1, rat anti-mouse IgG2a, rat anti-mouse IgG2b, rat anti-mouse IgG3 (all from BD Pharmingen), rat anti-mouse IgM, or rat anti-mouse IgE (Southern Biotechnology) followed by rinsing and blocking (1 h) with PBS-0.1% Triton X-100 + 1% bovine serum albumin. Appropriately diluted sera were applied at room temperature for 1 h prior to detection with Ig-specific biotinylated secondary antibodies (0.25 µg/ml; Pharmingen) and poly-horseradish peroxidase-streptavidin (Endogen)/tetramethylbenzidine substrate (Pharmingen). Values were determined from measurements of absorption at 430 nm in a Bio-Rad ELISA plate reader.

Analysis of leukocyte populations by flow cytometry. For quantitative estimation of tissue leukocyte populations, lymphoid organs were dissected, mechanically disrupted, and put through a 70-µm nylon mesh prior to staining (30 min on ice) with FITC-conjugated CD4 (GK1.5 clone), FITC-conjugated CD3, allophycocyanin-conjugated CD19, phycoerythrin (PE)-conjugated CD45R/B220-PE (clone RA3-6B2), NK1.1 (PK136) (all from BD Pharmingen), or allophycocyanin-conjugated rat anti-mouse CD8{alpha} (clones 5H10 and 53-6.7) as appropriate.

Granulocytes, NK cells, and monocytes/macrophages were assayed using rat anti-mouse Ly6G (Gr-1) or Ly6C-FITC (RB6-8C5 clone), FITC-conjugated rat anti-mouse CD49b or DX5-PE (DX5 clone), and rat anti-mouse CD11b-APC (M1/70 clone) or F4/80 (BM8), respectively. Cells were analyzed on a Becton Dickinson FACSCalibur instrument, and a total of 106 cells per sample was collected.

Analysis of dendritic cell trafficking by FITC skin painting. FITC skin painting was performed essentially as described previously (14). FITC (2.25 mg dissolved in 150 µl acetone dibutylphthalate) was applied to the shaved abdominal skin of wild-type, LYVE-1–/–, and LYVE-1+/– littermates. The following day, axillary and inguinal lymph nodes were dissected, the two lymph nodes on the left and right side were pooled, digested with collagenase D (0.5 mg/ml 30 min at 37°C), put through a 70-µm nylon mesh, stained with PE-labeled hamster anti-mouse CD11c (HL3) (BD Pharmingen), and analyzed by flow cytometry to quantitate dendritic cells dually labeled with FITC and CD11c.

Oxazolone-mediated contact hypersensitivity response. Contact hypersensitivity was induced in appropriate mice by initial sensitization with 100 µl 3% (wt/vol) oxazolone solution (Sigma) in 4:1 acetone-olive oil applied to the shaved abdominal skin followed by sequential elicitation 5 and 12 days later by application of 10 µl 1% (wt/vol) oxazolone in acetone-olive oil on both dorsal and ventral aspects of the left ear. The control right ear was treated with acetone:olive oil only. Ear swelling was measured with a Mitutoyo loop handle dial thickness gauge (McMaster-Carr) immediately before and at various times after elicitation.

Growth of subcutaneous tumors. B16F10 melanoma cells (kindly donated by Katja Simon, MRC Human Immunology Unit, Oxford) were resuspended in PBS, pH 7.5, and injected (0.5 x 106 in 200 µl) subcutaneously into wild-type (n = 9) or LYVE-1–/– (n = 11) mice. Tumor diameter was measured over a period of 5 weeks.

Lewis lung carcinoma cells (LL/2; American Type Culture Collection) were infected with retrovirus encoding VEGF-C in front of a green fluorescent protein-internal ribosome entry site construct or with green fluorescent protein-internal ribosome entry site constructs alone for controls. Infected cell populations were subjected to fluorescence-activated cell sorting twice on a DakoCytomation MoFlo high-speed cell sorter (Fort Collins, CO) prior to expansion and subcutaneous implantation in appropriate mice (11 to 12 weeks of age). After 14 days, tumors were harvested and processed for histological analysis. Tumor sizes were recorded after sacrifice by use of calipers, and tumor volumes were calculated using the formula length x width x height. Tumors were cut into 80-µm sections, stained with X-Gal (5-bromo-4-chloro-3-indolyl-ß-D-galactopyranoside) reaction, and photographed with a Zeiss Axiophot microscope.


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RESULTS
 
Generation of LYVE-1–/– mice. To generate mice lacking LYVE-1, we constructed a targeting vector that replaced the region from codon 33 (in exon 2) through exon 5 with a TM-LacZ reporter gene (Fig. 1A). The targeted gene thus encoded the N-terminal leader and the first eight amino acids from the mature LYVE-1 protein fused to a transmembrane version encoded by the TM-LacZ gene; when spliced, this gene encodes a ß-Gal protein which is targeted to the cell membrane. Mice carrying the mutated allele were screened by quantitative genomic PCR using primers and probes as described for the wild-type or mutant allele ES cell screening in Materials and Methods (Fig. 1A). Mice heterozygous or homozygous for the mutated LYVE-1 allele were born at normal Mendelian ratios and displayed no obvious physical or behavioral abnormalities.

To confirm that expression of the LacZ gene was driven by the LYVE-1 promoter, we stained heterozygous LYVE-1+/– mouse tissues for ß-Gal activity and compared this with the pattern obtained using LYVE-1 antibody. As shown in Fig. 1B to E, both the ß-Gal and the LYVE-1 antibody revealed the same extensive network of vessels characteristic of lymphatics on the surface of the intestine and associated Peyer's patches. Finally, the total absence of LYVE-1 antigen from LYVE-1–/– mice was confirmed for whole-mount tissue sections by immunohistochemistry with cognate antibody (for example, see results for intestine shown in Fig. 1F).

Normal development, ultrastructure, and fluid drainage function of lymphatics in LYVE-1–/– mice. To investigate whether LYVE-1 deficiency affects the growth or patterning of the lymphatic vessels, we stained whole-mount sections of intestine with the lymphatic marker VEGFR-3. As shown in Fig. 2A and B, the lymphatic vessels formed similar networks of mostly small and often blind-ended capillaries in both wild-type and LYVE-1 gene-targeted mice. Similar results were found for dermal lymphatics within whole-mount sections of the ear stained with the lymphatic marker podoplanin and the panendothelial marker CD31, in which the morphology of the mostly small and often blind-ended capillaries (diameter, 50 to 100 µm) could be clearly seen (Fig. 2C and D). Moreover, the complex architecture of lymph node cortical and medullary sinuses—conduits for circulating lymphocytes and antigen-presenting cells—was also found to be conserved in LYVE-1 gene-targeted mice, as visualized by immunofluorescence staining for the lymphatic marker VEGFR-3 (Fig. 2F and G). Finally, transmission electron microscopy of initial lymphatic vessels in intestinal mucosa and skin (see Fig. S1 in the supplemental material) indicated a formation of overlapping interendothelial junctions in LYVE-1–/– mice similar to that in LYVE-1+/+ mice. Consistent with this apparently normal patterning of the lymphatic vasculature in LYVE-1 gene-targeted mice, such animals showed no obvious signs of lymphedema.


Figure 2
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  		FIG. 2. Lymphatic patterning and lymph node architecture in LYVE-1 gene-targeted mice. (A and B) Whole-mount VEGFR-3 staining of intestine (viewed from the mucosal surface) from wild-type and LYVE-1 gene-targeted mice reveal that lymphatic vessels appear in the normal density and in a normal morphological pattern in control and knockout (ko) mice. (C and D) Whole-mount dorsal ear skin sections of wild-type and LYVE-1 gene-targeted mice were stained with hamster anti-mouse podoplanin (in green) and rat anti-mouse CD31 (in red). Frozen sections from wild-type (F) and LYVE-1 gene-targeted (G) mice were stained with goat anti-mouse VEGFR-3 and rabbit anti-LYVE-1.

In order to uncover more-subtle defects induced by gene disruption, we visualized the uptake and drainage of the lymphatic tracking dye Evans blue after direct injection into the footpad. As shown in Fig. 3A and B, the dye was distributed equally well between lymphatic vessels and draining lymph nodes in wild-type and LYVE-1 gene-targeted mice. To corroborate these findings using a second, independent method, we monitored the uptake and clearance of the fluorescent tracker dye FITC-dextran by the dense lymphatic network present in the tail skin. To allow continuous monitoring of uptake, the dye solution was injected at a constant pressure of 45 cm H2O for 20 min, and images were recorded using a fluorescence microscope. As shown in Fig. 3C and D, the FITC-dextran complex permeated the characteristic honeycomb network of dermal lymph vessels to similar depths (up to 20 µm) and at similar rates in both wild-type and LYVE-1–/– mice. Overall, these results indicate that LYVE-1 integrity is not required for normal lymphatic fluid drainage function.


Figure 3
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  		FIG. 3. Direct visualization of fluid drainage in LYVE-1 gene-targeted mice. (A and B) Distribution of Evans blue dye after injection into the hind footpads of wild-type (A) and LYVE-1 gene-targeted (B) mice. The dye labels large lymphatic vessels in the leg (insets) and draining lymph nodes within the abdomen (main images in panels A and B; iliac and renal nodes are shown by arrowheads) to similar extents in both control and knockout (KO) mice. (C and D) Levels of FITC-dextran uptake after injection into the dermis are similar in lymphatic vessels of the tails of wild-type (C) and LYVE-1 gene-targeted (D) mice as observed by fluorescence microscopy. Images represent tail lymphatics at successive distances (as shown) from the original injection site.

Hyaluronan homeostasis. Aside from its role in tissue fluid homeostasis, the lymphatic system plays a significant role in the uptake and turnover of extracellular matrix components. Indeed, hyaluronan, the primary ligand for LYVE-1, is degraded in lymph nodes after release from its proteoglycan binding partners in the interstitial tissue matrix (4). Furthermore, LYVE-1 itself can function in vitro as an endocytic receptor for hyaluronan and is expressed abundantly in vivo within lymph node sinus endothelium (22). We therefore investigated the effect of LYVE-1 gene deletion on hyaluronan homeostasis by measuring the steady-state levels of the glycosaminoglycan in different tissues. The results (Fig. 4A) showed no significant difference in the concentrations of hyaluronan in skin, muscle, intestine, and lung between the two groups of animals, and only trace levels within liver. This was also confirmed by probing tissue sections from wild-type and knockout mice with a biotinylated bHABP, which revealed similar distributions of hyaluronan within, e.g., the dermis of the ear (Fig. 4B and C) as well as in liver, lymph node, and intestine (not shown). Finally, a comparison of the serum hyaluronan levels of normal (0.783 ± 0.081 µg/ml; n = 12) and knockout (0.762 ± 0.048 µg/ml; n = 12) mice indicated no difference due to LYVE-1 gene deletion.


Figure 4
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  		FIG. 4. Hyaluronan levels in LYVE-1 gene-targeted mice. (A) The levels of hyaluronan in tissues from wild-type and LYVE-1–/– (KO) mice were determined by a competitive ELISA-like assay using biotinylated bHABP as described in Materials and Methods. (B) Frozen sections of ear tissue stained with bHABP/Alexa488 streptavidin (green) and podoplanin-Alexa594-conjugated goat anti-hamster 594 (red) to label hyaluronan and lymphatic vessels, respectively, show no significant differences between wild-type and LYVE-1 gene-targeted mice.

Leukocyte populations are unaltered in LYVE-1–/– mice. Given the abundant expression of LYVE-1 in normal lymph node sinus endothelium and the role of secondary lymphoid tissue in regulating resident and circulating leukocyte populations, we considered the possibility that LYVE-1 deficiency might have an impact on leukocyte development and/or compartmentalization. Hence, we carried out a comprehensive analysis of the number and composition of major leukocyte populations, including the CD4+ and CD8+ (helper and cytotoxic) T-cell subsets, CD19+ B cells, NK1.1/Ly49b+ NK cells, Ly6G/Gr1+ neutrophils, and F4/80/CD11b+ monocytes/macrophages in peripheral blood, spleen, thymus, and lymph nodes by use of flow cytometry. The results (Fig. 5A and B; also, see Fig. S2 in the supplemental material) revealed no significant differences between any of these leukocyte populations within either wild-type mice or homozygous/heterozygous LYVE-1 gene-targeted mice. In addition, ELISA determination of individual serum immunoglobulin classes (IgG, IgM, and IgA) and subtypes (IgG1, IgG2a, IgG2b, and IgG3) indicated no significant differences between wild-type and knockout mice (see Fig. S3 in the supplemental material). Together, these results indicate that LYVE-1 function is not required for the development or compartmentalization of immune cells within primary or secondary lymphoid tissue.


Figure 5
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  		FIG. 5. Leukocyte populations in LYVE-1 gene-targeted mice. (A and B) Leukocyte populations prepared from peripheral blood (A) and spleen (B) (see Materials and Methods) of wild-type and LYVE-1–/– (KO) mice were stained with directly conjugated antibodies as indicated and analyzed by flow cytometry. Values are the mean percentages of positive cells in the leukocyte gate ± standard errors (n = 3). Mono/Mac, monocytes/macrophages.

Migration of dendritic cells in afferent lymph. Based on homology with the leukocyte-homing receptor CD44, which supports rolling on blood vascular endothelium, we had speculated that LYVE-1 might mediate the adhesion or transmigration of cells entering the lymphatic vasculature (7, 10). A prominent example of such trafficking is the migration of antigen-presenting cells from tissue to draining lymph nodes during normal immune surveillance. To compare the process in normal mice with that in knockout mice, we measured the trafficking of epidermal dendritic cells to lymph node via afferent lymphatics using the technique of FITC skin painting. Dissection of the draining lymph nodes 18 h after dye application, followed by staining with the dendritic cell marker CD11c and flow cytometry, revealed that both the total CD11c-positive dendritic cell populations and the numbers of skin-derived CD11c/FITC double-positive cells were comparable in wild-type mice and both heterozygous LYVE-1+/– and homozygous LYVE-1–/– mice (Fig. 6A). Hence we conclude that LYVE-1 is not required for either entry or migration of dendritic cells through the afferent lymphatics.


Figure 6
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  		FIG. 6. Trafficking of dendritic cells and resolution of skin inflammation in LYVE-1 gene-targeted mice. (A) Analysis of dendritic cell antigen uptake and migration by FITC skin painting. FITC dissolved in acetone dibutylphthalate was applied to the shaved abdominal skin of wild-type, LYVE-1+/–, and LYVE-1–/– littermates (n = 3). The following day, axillary and inguinal lymph nodes were dissected, pooled together as left and right nodes, stained with PE-labeled CD11c, and analyzed by flow cytometry. Values are mean percentages ± standard errors for positive cells from the left and right draining lymph nodes from three mice (n = 3 x 2). (B) Delayed-type hypersensitivity response in LYVE-1 gene-targeted mice. Wild-type and LYVE-1–/– (Lyve-1KO) littermates were sensitized by a topical application of oxazolone (Ox) on the abdomen and challenged 5 days later by an application of oxazolone to the left ear while the right ear was treated with vehicle only (ctrl). A second elicitation was performed 7 days after the first elicitation. Ear swelling was measured with a dial thickness gauge immediately before and at various times after elicitation. Values shown are mean absolute increases in ear thickness in 10–2 mm for four mice.

Development and resolution of skin inflammation. Each of the parameters described above was used to evaluate the role of LYVE-1 in normal tissue homeostasis. To determine whether LYVE-1 function is manifest under conditions of tissue stress, we subjected both wild-type and LYVE-1–/– mice to allergen-induced contact hypersensitivity, a condition that promotes T-cell-mediated skin inflammation characterized by local tissue edema and the mobilization of epidermal Langerhans cell migration to draining lymph nodes via afferent lymphatics. To provoke inflammation, mice were sensitized by the consecutive topical application of oxazolone on the shaved abdominal skin and challenged 5 days later by the application of oxazolone on the ear. Measurement of edema as determined by ear thickness and microscopic examination of ear tissue after treatment showed no difference in the responses of the two groups of mice in either the magnitude or the rate of resolution of inflammation (Fig. 6B). These results suggest that the presence of LYVE-1 on lymphatic endothelial cells is required neither for T-cell-mediated hypersensitivity nor for clearance of the attendant inflammation/edema.

Growth and metastasis of subcutaneous tumors. Finally, we considered the requirement for LYVE-1 integrity on tumor growth. The development of a lymphatic supply by either cooption or lymphangiogenesis is known to influence tumor fluid drainage and, consequently, tumor hydrostatic pressure. Hence, any defects in lymphatic vessel formation might be expected to limit tumor growth. To investigate these phenomena, we transplanted syngeneic (C57BL/6) B16F10 melanoma cells subcutaneously into wild-type and LYVE-1–/– mice and monitored tumor diameter over a 5-week period. The results showed there was no obvious difference in tumor growth rates between the two groups of animals (Fig. 7A). We also investigated growth and lymph vessel development in a Lewis lung carcinoma that had been engineered to overexpress the lymphangiogenic growth factor VEGF-C by retroviral gene transfer. Again, these tumors showed similar growth rates in both wild-type and LYVE-1 gene-targeted mice (Fig. 7B), and there were no obvious differences in either the number of lymphatics, their patterns, or the depths of their penetration into tumor tissue, as assessed by ß-Gal histochemistry (Fig. 7C). Together, these results indicate that LYVE-1 is not obligatory for either the growth or the apparent function of tumor-associated lymphatics.


Figure 7
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  		FIG. 7. Growth and lymphangiogenesis of subcutaneous tumors in LYVE-1–/– mice. (A) B16F10 melanoma cells (0.5 x 106) were injected subcutaneously into wild-type (n = 9), LYVE-1+/– (het) (n = 10), and LYVE-1–/– (ko) (n = 8) mice. Values represent mean tumor diameters measured after 5 weeks. (B) Lewis lung carcinoma cells retrovirally transfected with VEGF-C were injected (1 x 106) subcutaneously into LYVE-1 knockout, heterozygous, and wild-type littermates (three mice per group), and tumor volume was determined subsequently (T, +14 days) as described in Materials and Methods. (C) Fourteen-day Lewis lung carcinoma tumors from heterozygous and LYVE-1 knockout mice stained with X-Gal to visualize intratumoral lymphatics.


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DISCUSSION
 
The near-ubiquitous expression and relative abundance of LYVE-1 in lymphatics in the adult and its appearance as one of the earliest lymphatic-specific genes during embryonic development had lead to the expectation that the receptor must play some fundamental role in maintaining either lymphatic architecture or normal lymphatic function (7, 19). Surprisingly, however, we found no developmental defects in LYVE-1-deleted mice. Rather, they displayed normal vessel morphology and patterning, as assessed by a comprehensive survey of different tissues by whole-mount histochemistry and immunohistology. Moreover, there was no evidence of edema, a condition associated with defective fluid drainage function that manifests as chylothorax or chylous ascites, respectively, in mice with targeted deletion of other genes expressed in lymphatic endothelium, such as those encoding the {alpha}9ß1 integrin (6) and the membrane sialomucin podoplanin (25). This was confirmed by lymphangiography with FITC-dextran and Evans blue dyes, which showed uptake and clearance both by superficial dermal lymphatics and by deeper collectors supplying the draining lymph nodes.

Recent studies in mouse embryogenesis have shown that LYVE-1 is present in cardinal vein endothelial cells before the initial separation of the hemovascular and lymphatic lineages and just after the first expression of VEGFR-3, whose integrity is essential not only for lymphatic but also for cardiac development (32). LYVE-1 is also present just prior to the expression of PROX-1, whose polarization in the cardinal vein signals the onset of budding and formation of the nascent lymph sacs at E9 to E10.5 (31, 32). The lack of any profound embryological phenotype in gene-deleted mice is therefore intriguing and suggests either that the expression of LYVE-1 at these early time points is not required for differentiation or lymphangiogenesis or that its loss can be compensated by other functionally similar molecules (also see below). Although such compensatory molecules have yet to be identified, they are unlikely to include CD44, as this molecule is not normally expressed by lymphatic endothelium and does not appear to be induced in the lymphatics of LYVE-1 knockout tissues (unpublished observation).

The lack of any defect in constitutive dendritic cell trafficking in LYVE-1–/– gene-deficient mice and their ability to resolve oxazolone skin contact-induced hypersensitivity and attendant edema formation argues against the role we originally proposed for the receptor as a gatekeeper for leukocyte adhesion/transmigration (10). In light of evidence that dendritic cells have the capacity to synthesize and present HA on their surfaces (17), we had anticipated that the migration of these cells would be at least partially disrupted in LYVE-1 knockout mice. However, one very recent finding from our laboratory relevant to this issue is that hyaluronan binding in lymphatic vessels in the context of normal tissues may be reversibly inactivated (18). Although we have not fully defined the molecular basis for inactivation, the emerging picture is of a reversible modification to LYVE-1 sugar chains that prevents interaction of the receptor with glycosaminoglycan while retaining expression at the cell surface. How this intriguing process is regulated is currently unknown. Perhaps the unique physiological function of LYVE-1, much like that of its homologue CD44, will be apparent only under specific conditions of tissue damage or disease.

A final possibility is that the loss of LYVE-1 in gene-deleted mice can be compensated by other HA receptors. The apparent maintenance of normal HA levels could be achieved through CD44 expressed on fibroblasts and tissue macrophages or the endocytic HARE molecule (hyaluronan receptor on endothelium) present in lymph node, spleen, and liver sinus endothelium, for example (34, 35). Further studies using LYVE-1/CD44 knockout mice and other double-knockout mice will be required to resolve this issue.


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ACKNOWLEDGMENTS
 
This work was supported by Unit funding from the United Kingdom Medical Research Council and by a project grant from Cancer Research UK (A399/C581).

We thank Anthony Dore, Nicholas Papadopoulos, Li Pan, Virginia Hughes, and Mary Simmons for assistance with flow cytometry and immunochemical staining; Ella Joffe for assistance with delayed-hypersensitivity experiments; and Jocelyn Holash and Irene Noguera for assistance with Lewis carcinoma experiments. We gratefully acknowledge David Valenzuela, Andrew Murphy and all the other members of the VelociGene team at Regeneron for the generation of the LYVE-1 knockout mice.


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FOOTNOTES
 
* Corresponding author. Mailing address: MRC Human Immunology Unit, Weatherall Institute of Molecular Medicine, John Radcliffe Hospital, Headington, Oxford OX3 9DS, United Kingdom. Phone: 44 1865 222313. Fax: 44 1865 222502. E-mail: djackson{at}hammer.imm.ox.ac.uk. Back

{triangledown} Published ahead of print on 13 November 2006. Back

{dagger} Supplemental material for this article may be found at http://mcb.asm.org/. Back

{ddagger} The contributions of both of these authors should be considered equal. Back


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Molecular and Cellular Biology, January 2007, p. 595-604, Vol. 27, No. 2
0270-7306/07/$08.00+0     doi:10.1128/MCB.01503-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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