of May 1, 2018.This information is current as

Cells and Then by T and B Cells+Stromal Growth First by CD11c−Vascular

Coordinated Regulation of Lymph Node

Kumar, Sha Tian, Martin Lipp and Theresa T. LuSusan Chyou, Fairouz Benahmed, Jingfeng Chen, Varsha

http://www.jimmunol.org/content/187/11/5558doi: 10.4049/jimmunol.1101724October 2011;

2011; 187:5558-5567; Prepublished online 26J Immunol 

MaterialSupplementary

4.DC1http://www.jimmunol.org/content/suppl/2011/10/26/jimmunol.110172

Referenceshttp://www.jimmunol.org/content/187/11/5558.full#ref-list-1

, 29 of which you can access for free at: cites 62 articlesThis article

        average*  

4 weeks from acceptance to publicationFast Publication! •    

Every submission reviewed by practicing scientistsNo Triage! •    

from submission to initial decisionRapid Reviews! 30 days* •    

Submit online. ?The JIWhy

Subscriptionhttp://jimmunol.org/subscription

is online at: The Journal of ImmunologyInformation about subscribing to

Permissionshttp://www.aai.org/About/Publications/JI/copyright.htmlSubmit copyright permission requests at:

Email Alertshttp://jimmunol.org/alertsReceive free email-alerts when new articles cite this article. Sign up at:

Print ISSN: 0022-1767 Online ISSN: 1550-6606. Immunologists, Inc. All rights reserved.Copyright © 2011 by The American Association of1451 Rockville Pike, Suite 650, Rockville, MD 20852The American Association of Immunologists, Inc.,

is published twice each month byThe Journal of Immunology

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

The Journal of Immunology

Coordinated Regulation of Lymph Node Vascular–StromalGrowth First by CD11c+ Cells and Then by T and B Cells

Susan Chyou,*,† Fairouz Benahmed,*,† Jingfeng Chen,*,† Varsha Kumar,*,† Sha Tian,*,†

Martin Lipp,‡ and Theresa T. Lu*,†,x

Lymph node blood vessels play important roles in the support and trafficking of immune cells. The blood vasculature is a component

of the vascular–stromal compartment that also includes the lymphatic vasculature and fibroblastic reticular cells (FRCs). During

immune responses as lymph nodes swell, the blood vasculature undergoes a rapid proliferative growth that is initially dependent

on CD11c+ cells and vascular endothelial growth factor (VEGF) but is independent of lymphocytes. The lymphatic vasculature

grows with similar kinetics and VEGF dependence, suggesting coregulation of blood and lymphatic vascular growth, but lym-

phatic growth has been shown to be B cell dependent. In this article, we show that blood vascular, lymphatic, and FRC growth are

coordinately regulated and identify two distinct phases of vascular–stromal growth—an initiation phase, characterized by up-

regulated vascular–stromal proliferation, and a subsequent expansion phase. The initiation phase is CD11c+ cell dependent and

T/B cell independent, whereas the expansion phase is dependent on B and T cells together. Using CCR72/2 mice and selective

depletion of migratory skin dendritic cells, we show that endogenous skin-derived dendritic cells are not important during the

initiation phase and uncover a modest regulatory role for CCR7. Finally, we show that FRC VEGF expression is upregulated

during initiation and that dendritic cells can stimulate increased fibroblastic VEGF, suggesting the scenario that lymph node-

resident CD11c+ cells orchestrate the initiation of blood and lymphatic vascular growth in part by stimulating FRCs to upregulate

VEGF. These results illustrate how the lymph node microenvironment is shaped by the cells it supports. The Journal of

Immunology, 2011, 187: 5558–5567.

Lymph nodes are sites of adaptive immune responses, al-lowing Ag-specific T and B cells to efficiently detectcognate Ag present in draining tissues and to interact with

one another. Blood vessels are critical to lymph node function,because they support the metabolic needs of the nodes and deliver,via the high endothelial venules (HEVs), recirculating lymphocytesto the lymph node parenchyma. The blood vasculature is suspendedalong with the lymphatic vasculature within a reticular network ofcollagen-rich fibrils covered by fibroblastic reticular cells (FRCs).The lymphatic sinuses control entry of Ags and APCs from thedraining tissues and the egress of lymphocytes from the lymphnodes, whereas the reticular network regulates lymphocyte lo-calization, migration, and homeostasis (1–4). During immune re-sponses, when lymph nodes can swell to many times their origi-

nal size, the different elements of the vascular–stromal compart-ment also grow. The feeding arteriole delivering arterial flow isremodeled, and the HEVs and other portions of the microcircu-lation undergo a proliferative expansion, resulting in increaseddelivery and entry of blood-borne lymphocytes (5–10). The lym-phatic vasculature also expands, allowing for increased migratorydendritic cell entry (11–13). The reticular network undergoes ap-parent expansion as well, presumably to accommodate the greatlyincreased cellularity in the stimulated lymph node (14). Under-standing how the blood vascular growth is regulated and whetherthe growth is coordinately regulated with that of other cellularelements of the vascular–stromal compartment may help to under-stand how immune responses are orchestrated.We have previously shown using flow cytometry that the total

population of endothelial cells (consisting of a mix of 85% bloodvascular endothelial cells and 15% lymphatic endothelial cells)in lymph nodes demonstrate upregulation of proliferation within2 d after immunization and that this proliferation is accompanied bymarked expansion of endothelial cell numbers in subsequent days.The initial upregulation of proliferation is dependent on CD11c+

cells, because depletion of CD11c+ cells abrogates this upregu-lation. Injection of bone marrow-derived dendritic cells (BMDCs)can drive upregulation of proliferation at day 2 even in RAG12/2

mice, suggesting that dendritic cells can initiate vascular expan-sion in a manner independent of T and B cells (8). Studies focusedon the lymphatic vasculature showed that lymphatic expansion isdetectable by day 4 and that this expansion is dependent on B cellsat this time point (11, 12), pointing to a role for lymphocyteseither during initiation of lymphatic growth or during lymphaticexpansion. Whether these studies together represent conflictingdata on the role of lymphocytes in vascular growth, regulation thatis unique to blood versus lymphatic endothelial cells, or whetherthey reflect an early lymphocyte-independent initiation phase

*Autoimmunity and Inflammation Program, Hospital for Special Surgery, New York,NY 10021; †Division of Pediatric Rheumatology, Hospital for Special Surgery, NewYork, NY 10021; ‡Department of Molecular Tumor Genetics and Immunogenetics,Max Delbruck Center for Molecular Medicine, Berlin 13125, Germany; and xDepart-ment of Microbiology and Immunology, Weill Cornell Medical College, CornellUniversity, New York, NY 10021

Received for publication June 10, 2011. Accepted for publication September 25,2011.

This work was supported by National Institutes of Health Grants R01-AO068900 andR01 AI079178 and a Lupus Research Institute award (to T.T.L.).

Address correspondence and reprint requests to Dr. Theresa T. Lu, Hospital forSpecial Surgery, 535 East 70th Street, New York, NY 10021. E-mail address: lut@hss.edu

The online version of this article contains supplemental material.

Abbreviations used in this article: DT, diphtheria toxin; DTR, diphtheria toxin re-ceptor; FDG, fluorescein-di-b-D-galactopyranoside; FRC, fibroblastic reticular cell;b-gal, b-galactosidase; HEV, high endothelial venule; iBALT, induced BALT; KO,knockout; PNAd, peripheral node addressin; VEGF, vascular endothelial growthfactor; WT, wild-type.

Copyright� 2011 by TheAmericanAssociation of Immunologists, Inc. 0022-1767/11/$16.00

www.jimmunol.org/cgi/doi/10.4049/jimmunol.1101724

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

followed by a lymphocyte-dependent expansion phase has notbeen fully addressed, but Liao and Ruddle (12) showed that earlyphenotypic changes on HEVs are T and B cell independent,whereas later alterations to HEV are B cell dependent, suggestingtwo phases of vascular regulation that could potentially also existfor the regulation of vascular growth.The lymph node vasculature is intimately associated with FRCs,

because the FRCs ensheathe the vessels as well as the network ofcollagen-rich fibrils that provide the infrastructure for the lymphnodes (15). The FRCs express chemokines and cytokines neces-sary for lymphocyte compartmentalization and homeostasis (16,17), provide a substrate upon which lymphocytes migrate (18),and help to promote peripheral T cell tolerance (19). The reticularnetwork may help regulate vascular activity, because the com-partment between the FRCs and the fibrils and vessels they en-sheathe comprise a conduit system that allows for the delivery ofsmall molecules through the reticular network to the blood ves-sels. This conduit system receives afferent lymphatic flow, andmolecules generated in the draining tissue or, potentially, by theFRCs of the reticular network can be delivered to the blood ves-sels (4, 15, 20). The FRCs are also likely to directly regulatevascular activity. The integrity of FRC organization around HEVsis associated with alterations in vascular permeability (21). GP38/podoplanin is highly expressed on lymphatic endothelial cells butat lower levels, and it also marks FRC populations throughout thelymph node (17, 22–24). Analysis of vascular endothelial growthfactor (VEGF) reporter mice as well as of isolated FRCs showedthat gp38+ FRCs, especially the ones immediately around and inthe vicinity of blood and lymphatic vasculature, are the main ex-pressers of VEGF within lymph nodes (9). VEGF regulates homeo-static endothelial cell proliferation and is also required for theproliferative expansion of both the blood vascular and lymphaticvasculature after immunization (8, 9, 11, 13). VEGF protein levelsare upregulated by 2-fold as early as day 1 after lymph nodestimulation, and this increase is sensitive to CD11c+ cell depletion,suggesting that CD11c+ cells may mediate upregulation of endo-thelial cell proliferation at least in part by inducing upregulationof VEGF (8). GP38+ FRCs are the main expressers of VEGFmRNA both at homeostasis and at day 2 after immunization (9),but the source of the additional VEGF in the lymph nodes afterimmunization has been unclear. Immunochemical staining forVEGF localizes the protein to B cell areas after immunization, andthis localization is dependent on lymph node B cells, suggestingthat B cells may be a potential source of or repository for VEGF(11). A role for B cells is further supported by increased blood andlymphatic vascular growth in transgenic mice expressing humanVEGF in B cells (25). In contrast, density of in situ hybridizationsignals and real-time PCR for VEGF mRNA show no increaseafter immunization, leading to the idea that VEGF may betransported in from distal sources such as the inflamed skin (13).Understanding whether FRCs proliferate and expand along withendothelial cells and whether they upregulate VEGF expressionafter immunization would help to understand the regulation oflymph node vascular growth.A further unresolved issue in the regulation of vascular growth

in stimulated lymph nodes is the identity of CD11c+ cells thatmediates the initial upregulation of endothelial cell proliferation(8). CD11chiMHCIImed cells appear to contribute in part, becausetheir depletion attenuates upregulation of endothelial cell prolif-eration at day 2, but the majority of the effect is likely to bemediated by CD11cmed cells that include CD11cmedMHCIIhi skin-derived cells and CD11cmedMHCIImed cells (21). As mice wereimmunized in the footpad, a potential scenario is that skin den-dritic cells are stimulated to migrate to the draining node to induce

upregulation of endothelial cell proliferation. This scenario is alsoattractive because afferent lymphatic flow has been implicated toregulate HEV function (26–28). The requirement for skin den-dritic cells or their migration to lymph nodes, however, has notbeen tested.In this study, we first examine the role of T and B cells in lymph

node blood vascular growth and show that the growth is made upof a T/B cell-independent and CD11c+ cell-dependent initiationphase, followed by a T and B cell-dependent expansion phase.Lymphatic endothelial cells and FRCs proliferate and expand co-ordinately with the blood vascular endothelial cells. We showthat skin-derived dendritic or other CD11c+ cells do not mediatethe upregulation of endothelial and FRC proliferation during theinitiation phase, although our experiments uncover a regulatoryrole for CCR7-dependent radioresistant cells in controlling themagnitude of endothelial and FRC proliferation. Furthermore, weshow in this article that FRCs upregulate VEGF expression duringthe initiation phase and that dendritic cells can stimulate fibro-blastic cells to upregulate VEGF, supporting the thought thatCD11c+ cells induce upregulation of blood vascular and lymphaticendothelial cell proliferation by stimulating FRCs to upregulateVEGF. These results extend our understanding of the regulationof lymph node vascular growth and underscore the coordinatedregulation of the individual elements of the vascular–stromal com-partment.

Materials and MethodsMice

C57BL/6 mice between 6 and 12 wk of age were obtained from The JacksonLaboratory (Bar Harbor, ME), Taconic Farms (Hudson, NY), or NationalCancer Institute (Frederick, MD). RAG12/2 (29) and CD11c-diphtheriatoxin receptor (DTR) mice (30) were originally from The Jackson Labo-ratory and bred in our facility. mMT and bd TCR-deficient mice were fromThe Jackson Laboratory. CCR72/2 mice (31) and Langerin-DTR mice(32) were as described previously. VEGF-lacZ mice were originally ona CD-1 background and crossed over 10 generations to C57BL/6 mice forour studies (9, 33). All animal procedures were performed in accordancewith the regulations of the Institutional Animal Use and Care Committeeof the Hospital for Special Surgery.

Mouse treatments and immunizations

For immunization with OVA in CFA (OVA/CFA), mice received hindfootpad injection of 20 ml OVA/CFA as described previously (8). Micereceiving BMDCs were injected in hind footpads with 1 3 106 cells/footpad in 20 ml volume as described previously (8).

For proliferation studies, mice were given i.p. injections of 2 mg BrdU at18 and 1 h prior to sacrifice and 0.8 mg/ml BrdU in the drinking waterbetween injections as described previously (8, 21).

CD11c-DTR mice express simian DTR that allows for depletion ofCD11c+ cells upon diphtheria toxin (DT) administration (30). Depletion isas described previously (8, 21). Mice were injected in one footpad withindicated amount of DT (Calbiochem, San Diego, CA, or List BiologicalLaboratory, Campbell, CA) in 10–15 ml volume. At 8 h later, they wereimmunized with OVA/CFA in the same footpad. To control for the DTinjection, control CD11c-DTR mice were injected with inactive mutant DTcross-reactive material (CRM 197; Calbiochem).

For T cell depletion in mMT mice, 500 mg anti-CD4 (clone GK1.5) andanti-CD8 (clone 53-6.72) or isotype controls (all from BioXCell, WestLebanon, NH) were injected i.p. at days 25, 24, 23, 0 (day of immu-nization), and +3 (34). Depletion was verified by flow cytometry, withCD4+ T cells depleted by 97% and CD8+ T cells depleted by 92% indraining nodes.

BMDC preparation

BMDCs were generated as described previously (8). Briefly, bone marrowcells were cultured with 8–10% supernatant from GM-CSF–expressingJ558L cells and matured with LPS on day 7. They were harvested on day8, washed, and injected at 1 3 106 cells/hind footpad. BMDC preparationswere depleted of contaminating Gr-1+ neutrophils using MACS magneticselection (Miltenyi Biotec, Auburn, CA).

The Journal of Immunology 5559

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

Flow cytometry analysis

For flow cytometric analysis of endothelial cell and FRC numbers andproliferation, we prepared single-cell suspensions and stained cells asdescribed previously (8). In brief, lymph nodes were digested with colla-genase type II (Worthington, Lakewood, NJ) and stained with Abs forflow cytometry. Abs used are against CD45 (BD Biosciences, San Jose,CA), CD31 (BD Biosciences), peripheral node addressin (PNAd; BD Bio-sciences), gp38 (BioLegend, San Diego, CA, or Developmental StudiesHybridoma Bank, Iowa City, IA), and BrdU (Invitrogen, Carlsbad, CA).The unconjugated anti-PNAd is detected using fluorophore-conjugatedanti-rat IgM (Jackson ImmunoResearch Laboratories, West Grove, PA).Cells were analyzed using a FACSCanto (BD Biosciences) and CellQuestPro (BD Biosciences) software.

Flow cytometric analysis of b-galactosidase (b-gal) activity in VEGF-lacZ mice is as described previously (9). Briefly, the cells were stained forextracellular markers and then incubated with the b-gal substrate fluo-rescein-di-b-D-galactopyranoside (FDG) (Invitrogen) in water for 1 min toallow entry of FDG and subsequent cleavage and release of fluorescein.The reaction was stopped by addition of 5-fold more media and placementon ice, and samples were immediately run on the flow cytometer.

BMDC-3T3 cell cocultures

NIH-3T3 cells were cultured in DMEM (Mediatech, Herndon, VA) con-taining 10% heat-inactivated calf serum (Hyclone, Logan, UT) as describedpreviously (9). Cells were plated at 5 3 103 cells/well in 24-well plates,and the next day, the medium was changed and Gr-1–depleted BMDCs orthe contaminating Gr-1+ fraction was added at 3 3 105/well 3 2 d. Su-pernatant was collected for VEGF ELISA, which was performed as de-scribed previously using a kit (DuoSet mouse VEGF; R&D Systems) (9).For real-time PCR of the 3T3 cells, cells were trypsinized, and 3T3 cellswere isolated from adherent BMDCs by magnetic depletion of CD45+

cells, yielding .96–99% 3T3 cells. RNA was extracted and subjected toreal-time PCR for VEGF and normalized to cyclophilin as described pre-viously (9).

ResultsDependence on T and B cells separates initiation phase fromsubsequent phase of vascular expansion

We have shown previously that Tand B cells are dispensable for theinitial upregulation of endothelial cell proliferation at day 2 afterlymph nodes are stimulated with footpad injection of BMDCs.After day 2, proliferation is accompanied by a marked expansion ofendothelial cell numbers by day 5 (8, 21), and we asked whether Tand B cells were also dispensable for this subsequent expansion.We examined endothelial cells as we have previously done, gatingon CD452CD31+ cells (Fig. 1A). At day 5 after BMDC injection,RAG12/2 lymph nodes showed no growth in cellularity (Fig. 1B),and total endothelial cell numbers showed no expansion (Fig. 1C)and a poor rate of proliferation (Fig. 1D), suggesting that T and/orB cells are critical for mediating the vascular expansion thatoccurs after day 2. This result combined with our previous resultspoint to two separate phases of vascular growth—an early T andB cell-independent initiation phase, whereby endothelial cell pro-liferation is upregulated, followed by a T and/or B cell-depen-dent expansion phase.To confirm that these two phases were not specific to stimulation

with BMDCs, we immunized wild-type (WT) and RAG12/2 micewith OVA/CFA. In these experiments, we also included additionalmarkers of endothelial cells to examine the regulation of differentsubsets of endothelial cells in more detail. We have previouslyused PNAd expression to mark HEV endothelial cells within theCD452CD31+ endothelial cell population (1, 9). In this article, weidentified HEV endothelial cells using PNAd again and furtherdivided the PNAd2 population into gp382 blood vascular endo-thelial cells and gp38+ lymphatic endothelial cells (Fig. 1A) (17,22). Note that the gp38+ lymphatic endothelial cells make up onlya small percentage (∼10–15%) of the total endothelial populationin popliteal lymph nodes (Fig. 1A, 1F), whereas they make up∼30–45% of the endothelial cells in brachial nodes. In this article,

we will refer to PNAd+ endothelial cells as “HEV endothelialcells,” PNAd2 cells as a whole as “non-HEV mixed endothelialcells,” PNAD2gp382 cells as “non-HEV blood vascular endo-thelial cells,” and PNAD2gp38+ cells as “lymphatic endothelialcells.”At day 2 after OVA/CFA inWTmice, lymph node cellularity was

increased 4-fold from baseline (Fig. 1E). Total endothelial cellnumbers were not significantly increased (Fig. 1F), but HEV en-dothelial cells doubled in numbers (Fig. 1F). The RAG12/2 miceshowed a similar pattern of lymph node expansion (Fig. 1E) (WT= 4.4-fold [60.8]; knockout [KO] = 3.3-fold [61.3]; p = 0.05) andHEV expansion at day 2 (Fig. 1F). All endothelial cell types inboth WT and RAG12/2 mice showed a marked increase in pro-liferation by day 2 (Fig. 1G). These results confirmed that theinitiation of lymph node vascular growth is T and B cell inde-pendent and also established that the T and B cell independence isconsistent for all endothelial cell subsets examined.By day 5 after OVA/CFA, lymph nodes were greatly enlarged

(Fig. 1H) (18-fold [67.2]), and total endothelial cell numbers inWT mice expanded by ∼4-fold, with the HEV subset expandingmost dramatically and consequently accounting for ∼50% of thetotal endothelial cells (Fig. 1I). In contrast to the unimportance ofT and B cells at day 2 and similar to the findings in Fig. 1B, lymphnodes were only modestly expanded in size (Fig. 1H) (5-fold[62.8]; p , 0.01 compared with WT), and total endothelial cellnumbers in RAG12/2 mice were not expanded, although HEVandlymphatic subsets had a low level of expansion (Fig. 1I), with theHEV endothelial cells demonstrating an expansion similar to whathad been observed at day 2 (Fig. 1F). The proliferation rate inmost endothelial cell subsets was upregulated at day 5 comparedwith unimmunized RAG12/2 mice, but the increase in prolifera-tion was modest when compared with the increase in WT mice(Fig. 1J). Taken together, our results suggested that there are twodistinct phases of vascular growth after immunization: a T andB cell-independent initiation phase between days 0 and 2 and a Tand/or B cell-dependent vascular expansion phase after day 2through at least day 5.

FRCs proliferate and expand with endothelial cells

We examined the extent to which the gp38+ FRCs proliferate andexpand in number over time after OVA/CFA. We used the gatingstrategy shown in Fig. 2A. Similar to the pattern seen with en-dothelial cells, gp38+ FRCs in WT mice at day 2 after immuni-zation had not yet expanded but showed upregulated proliferation(Fig. 2B, 2C). By day 5, the gp38+ FRCs in WT mice were ex-panded in number and continued to proliferate (Fig. 2D, 2E). InRAG12/2 mice at day 2, the upregulation of gp38+ FRC prolif-eration from baseline was preserved and even enhanced (Fig. 2C).GP38+ FRCs continued to proliferate at higher than baseline ratesat day 5 in RAG12/2 mice, but the expansion in gp38+ FRCnumbers was completely abrogated (Fig. 2D, 2E), suggesting thepossibility that proliferated cells were unable to survive in theabsence of lymphocytes. Taken together, these results indicate thatgp38+ FRCs proliferate and expand in parallel with endothelialcells in immune-stimulated lymph nodes and that, similar to en-dothelial cells, the initial proliferation is T and B cell indepen-dent, whereas the later expansion is T and/or B cell dependent. Vas-cular growth, then, occurs in the context of a more generalizedvascular–stromal proliferation and expansion.

B cells and T cells together mediate vascular–stromalexpansion

To better understand the relative contributions of B and T cells tovascular–stromal expansion, we examined mice deficient in B or

5560 TWO PHASES OF REGULATING LYMPH NODE VASCULAR STROMAL GROWTH

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

T cells. B cell-deficient mMT mice at day 5 showed an increase inlymph node cellularity (Fig. 3A) and in HEV and lymphatic en-dothelial cell numbers from baseline (Fig. 3B) that was only par-tial relative to that of WT mice but greater than the expansionfound in RAG12/2 mice (Fig. 1I). Non-HEV blood vascular en-dothelial cells were reduced in number at baseline in mMT mice incomparison with that of WT mice (Fig. 3B), but the fold increasein expansion was the same as in wild-type mice (WT = 2.8-fold[61.3] and mMT = 3-fold [60.5]; p = 0.66 t test), indicating thatB cell absence did not affect immunization-induced expansion ofthis population. Proliferation rates of endothelial cells at day 5 inmMT mice were attenuated (Fig. 3C) but to a lesser extent than

that seen in RAG12/2 mice (compare with Fig. 1J). FRCs alsoshowed partial expansion in numbers in mMT mice (Fig. 3D), butsimilar to what was observed in RAG12/2 mice (Fig. 2E), FRCproliferation rates were similar to that of WT mice (Fig. 3E).B cell deficiency, then, resulted in attenuated expansion and pro-liferation of HEV and lymphatic endothelial cells and in attenu-ated expansion but normal proliferation rate of FRCs.bd TCR-deficient mice lacking ab and gd T cells had reduced

lymph node cellularity and numbers of endothelial cells both atbaseline and at day 5 after immunization when compared with WTmice (Fig. 3F, 3G). However, the fold increase in expansion in thenumbers of HEV and non-HEV blood vascular endothelial cells

FIGURE 1. Lymph node vascular growth is characterized by an initial T/B cell-independent initiation phase followed by a T/B cell-dependent expansion

phase. A, Flow cytometry plots showing subsetting of CD452CD31+ endothelial cells into PNAd+ HEV endothelial cells (HEV EC), PNAd2gp382 non-

HEV blood vascular endothelial cells (non-HEV BEC), and PNAd2gp38+ lymphatic endothelial cells (LEC). B–D, WT or RAG12/2 mice were injected

with 106 BMDCs in the hind footpads on day 0, and draining popliteal lymph nodes were examined on day 5. Unimmunized mice received either no

footpad injection or were sham injected with PBS. B, Lymph node cellularity as determined by count of lymph node cells. C, Number of total CD452

CD31+ endothelial cells as determined by flow cytometric analysis. D, Proliferation rate of CD452CD31+ endothelial cells as determined by the percent of

endothelial cells that are BrdU+. E–J, WT or RAG12/2 mice were immunized with OVA/CFA on day 0, and draining popliteal nodes were examined on day

2 (E–G) or on day 5 (H–J). Unimmunized mice received either no footpad injection or were sham injected with PBS. E and H, Lymph node cellularity. F

and I, Number of endothelial cells in each subset. G and J, Proliferation rate of endothelial cell subsets. For all studies except for the day 5 proliferation

studies, n = at least six mice per group over at least three experiments. For day 5 proliferation studies, n = 3 mice over two experiments for RAG12/2 PBS

group, and n = at least six mice per group over at least three experiments for all other groups. *p , 0.05 and **p , 0.01 in comparison with the same cells

in unimmunized controls of the same genotype using t test; +p , 0.05 and ++p , 0.01 in comparison with the same cells in WT mice receiving same

treatment.

The Journal of Immunology 5561

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

were similar (for HEV, WT = 12-fold [68] and KO = 12-fold [69],p = 0.96; for non-HEV, WT = 4-fold [61.5] and KO = 4-fold[61.5], p = 0.87), suggesting that deficiency of T cells by them-selves did not prevent expansion of these populations. The foldincrease in lymphatic endothelial cell expansion was greater in bdTCR-deficient mice (WT = 7-fold [65] and KO = 17-fold [68];p = 0.02) echoing the recent findings by Koh and colleagues (35).Curiously, despite equal or greater expansion, HEV and lymphaticendothelial cell proliferation rates in T cell-deficient mice wereattenuated at day 5 (Fig. 3H), suggesting that T cells were requiredfor full proliferation of these populations, but this contributionwas not required by itself for full expansion. Non-HEV bloodvascular endothelial cell proliferation was reduced at baseline andafter immunization (Fig. 3H) and showed a similar fold increase inWT and bd TCR-deficient mice (WT = 25-fold [67.7] and KO =36-fold [611.7]; p = 0.11), consistent with the idea that T celldeficiency by itself did not affect non-HEV blood endothelial cellproliferation. In contrast to the endothelial cells, FRC expansionand proliferation were both reduced (Fig. 3I, 3J). In summary,T cell deficiency resulted in attenuated HEV and lymphatic pro-liferation without attenuation of expansion and in attenuated FRCproliferation and expansion.As B cell-deficient mice had only a partial decrease in vascular–

stromal expansion and T cell-deficient mice showed no decreasein vascular expansion, we considered that the nearly complete lackof vascular–stromal expansion in RAG12/2 mice might reflectpartially redundant roles for B and T cells. We depleted CD4 and

CD8 cells in mMT mice, and the resulting lack of both B andCD4/8 T cells led to small lymph nodes and complete abrogationof vascular–stromal expansion (Fig. 3K–M). Taken together, theseresults suggest that B and T cells together are required for vas-cular–stromal expansion.

CD11c+ cells are required for the blood vascular, lymphatic,and FRC proliferation during the initiation phase

We have previously shown that depletion of all CD11c+ subsets byfootpad injection of 100 ng DT into CD11c-DTR mice greatly re-duces the upregulation of endothelial cell proliferation normallyseen at day 2 after immunization (8, 21). In this study, we askedwhether CD11c+ cells were important for the proliferation ofboth blood and lymphatic endothelial cells and also for the gp38+

FRCs. As we have previously shown, footpad administration of100 ng DT resulted in relative depletion of all CD11c+ subsets, butespecially of CD11chiMHCIImed and CD11cmedMHCIIhi subsets(Fig. 4A, 4B), and also resulted in reduced proliferation of totalendothelial cells at day 2, with the HEV cells being less sensitiveto and requiring greater levels of CD11c+ cell depletion than non-HEV–mixed endothelial cells (Fig. 4C) (8, 21). Of the non-HEV–mixed endothelial cells, both non-HEV blood vascular endothelialcell and lymphatic endothelial cell proliferation were sensitive toCD11c+ cell depletion (Fig. 4C). Upregulation of FRC prolifera-tion was also attenuated upon CD11c+ cell depletion (Fig. 4D).These results indicated that the proliferation of blood vascular andlymphatic endothelial cells and FRCs during the initiation phaseare coregulated and are suggestive of a scenario whereby CD11c+

cells orchestrate a program of vascular–stromal proliferation.

Skin-derived dendritic cells are not required forvascular–stromal proliferation during the initiationphase and CCR7 plays a modest regulatory role

The CD11c+ cells that mediate the vascular–stromal proliferationduring the initiation phase could be resident within the lymphnodes or could derive from the draining skin. The CD11cmed

MHCIIhi dendritic cells are a candidate population of CD11c+

cells (21) and include migratory skin-derived dendritic cells thatrequire CCR7 to enter lymph nodes (31, 36, 37), so we askedabout the importance of dendritic cell migration from the skinduring the initiation phase by examining CCR72/2 mice. Con-sistent with the role of CCR7 in T and B cell entry into lymphnodes at homeostasis (31, 38), unstimulated brachial nodes weresmaller in CCR72/2 mice than in WT mice. However, the cellu-larity of the draining popliteal lymph nodes at day 2 after OVA/CFA was similar to that of WT mice (Fig. 5A), suggesting thatrecruitment of lymphocytes after immunization was not CCR7dependent. CD11cmedMHCIIhi dendritic cells were reduced in num-ber as expected in CCR72/2 mice (Fig. 5B, 5C). CCR72/2

mice had more HEV endothelial cells in nondraining brachialnodes and more HEV and non-HEV blood vascular endothelialcells in the draining popliteal nodes (Fig. 5D), suggesting a regu-latory role for CCR7-dependent cells. Endothelial cell prolifera-tion rates in both nondraining and draining lymph nodes wereequal to that of WT mice (Fig. 5E). GP38+ FRC numbers andproliferation were not affected in CCR72/2 mice (Fig. 5F, 5G).These results suggested that CCR7-dependent migration of den-dritic cells was not required to mediate the upregulation of en-dothelial and FRC proliferation during the initiation phase andwere, instead, suggestive of a regulatory role for CCR7-dependentcells.We asked further whether any defect in vascular–stromal pro-

liferation could be observed if we separated out the contribu-tions of radiosensitive and radioresistant CCR7-dependent cells.

FIGURE 2. FRCs proliferate and expand with the endothelial cells. WT

or RAG12/2 mice were immunized with OVA/CFA on day 0 and draining

popliteal nodes were examined on day 2 (B, C) or day 5 (D, E). A, Gating

scheme to identify gp38+ FRCs in flow cytometry plots. B and D, Number

of CD452CD312gp38+ FRCs at denoted time points. C and E, FRC

proliferation as measured by BrdU uptake at denoted time points. For day

5 proliferation studies, n = 3 mice over two experiments for RAG12/2 PBS

group, and n = at least six mice per group over at least three experiments

for all other groups. **p , 0.01 in comparison with unimmunized controls

of the same genotype using unpaired t test; +p , 0.05 in comparison with

WT mice receiving same treatment. For all studies except for the day 5

proliferation studies, n = at least six mice per group over at least three

experiments.

5562 TWO PHASES OF REGULATING LYMPH NODE VASCULAR STROMAL GROWTH

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

Langerhans cells derived from the epidermis and a CD11b+ sub-population of dermal dendritic cells are among the radioresistantdendritic cells in the skin (39, 40), whereas cells such as Langerin+

DEC205+ dermal dendritic cells are radiosensitive (41–43). Wegenerated mixed bone marrow chimeras to examine the contri-bution of radiosensitive dendritic and other non-T–non-B CCR7-dependent cells. Irradiated WT mice were reconstituted with 80%CCR72/2RAG12/2 and 20% WT bone marrow. These chimerashad a less robust lymph node enlargement when compared withCCR7-sufficient chimeras at day 2 after OVA/CFA (SupplementalFig. 1A) but had unchanged endothelial cell and FRC numbers andproliferation (Supplemental Fig. 1B–E), suggesting that radio-sensitive CCR7-dependent non-T–non-B cells mediated the fullextent of initial lymph node swelling but otherwise did not affectendothelial or FRC proliferation.To test the potential role of radioresistant CCR7-dependent cells,

we reconstituted CCR72/2 recipients with WT bone marrow. Inthese chimeras irrespective of the genotype, B cells and CD11chi

MHCIImed cells were .99% donor derived, indicating successfulreplacement of radiosensitive cells with donor cells. CD11cmed

MHCIIhi cells were 91% donor derived in the draining lymphnodes of WT→WT chimeras, reflecting a 9% contribution of theradioresistant Langerhans cells and dermal dendritic cell subset atday 2 after immunization. In the WT→CCR7 chimeras, the donor-derived cells were increased to .98%, likely reflecting the lackof migration to the lymph node of the radioresistant CCR7-dependent dendritic cells. The CD11cmedMHCIImed cells were95% donor-derived in chimeras, regardless of recipient genotype.WT→CCR72/2 chimeras had larger draining and nondrainingnodes than WT→WT chimeras (Supplemental Fig. 1F). TheWT→CCR72/2 chimeras had more HEV endothelial cells in thenondraining brachial nodes and more HEV and non-HEV bloodendothelial cells in the draining popliteal nodes (SupplementalFig. 1G), resembling the phenotype in the CCR72/2 mice. Pro-liferation of non-HEV blood vascular endothelial cells and ofgp38+ FRCs was increased in the draining nodes of WT→CCR72/2

chimeras when compared with the nodes of WT→WT chimeras(Supplemental Fig. 1H, 1J). FRC numbers were unchanged(Supplemental Fig. 1I). These results suggested that radioresistantCCR7-dependent cells do not contribute to the upregulation of

FIGURE 3. B and T cells together

mediate the expansion phase. mMT mice

(A–E) and bd TCR2/2 (F–J) and con-

trols were immunized with OVA/CFA

on day 0, and draining popliteal nodes

were examined on day 5. CD4 and

CD8 T cell-depleted mMT mice (K–M)

were immunized as above but examined

on day 4. A, F, and K, Lymph node

cellularity. B, G, and L, Number of en-

dothelial cells in each subset. C and H,

Proliferation rate of endothelial cells in

each subset. D, I, and M, Numbers of

gp38+ FRCs. E and J, Proliferation rate

of gp38+ FRCs. For mMT mice and bd

TCR2/2; n = 5–6 mice in each group

over three experiments. For T cell-

depleted mMT mice, n = 5 mice in

each group over two experiments. *p ,0.05 and **p, 0.01 in comparison with

the same cells in unimmunized controls

of the same genotype using t test; +p ,0.05 and ++p , 0.01 in comparison with

the same cells in WT mice receiving

same treatment.

The Journal of Immunology 5563

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

vascular–stromal proliferation but play a modest regulatory role,controlling the magnitude of lymph node cellularity, endothelialcell numbers, and non-HEV blood vascular endothelial cell andFRC proliferation.Although CCR7-mediated migration of skin dendritic cells to

lymph nodes was not necessary for initiation of lymph node vas-cular growth, cells in the skin can potentially regulate lymphnode events by delivery of molecules to the lymph node via theafferent lymphatics (44, 45). We tested directly whether dendriticand other CD11c+ cells from the skin were required for the vas-cular–stromal proliferation seen during initiation. We adminis-tered a low dose (25 ng) of DT locally to the footpad, reasoningthat the DT should deplete local skin CD11c+ cells and, at this lowdose, be less effective at reaching the lymph node to depleteCD11c+ cells located within the node. The DT resulted in a 70%reduction of CD11cmedMHCIIhi cells in the lymph node whileother populations were preserved in number (Fig. 6A, 6B), con-sistent with the idea that we had mostly depleted skin-derivedmigratory dendritic cells. Endothelial cell proliferation was notaffected (Fig. 6C), suggesting that the depleted skin-derived den-dritic cells were not important in mediating the vascular prolif-eration during the initiation phase.

Dendritic cells stimulate fibroblasts to upregulate VEGFexpression

We asked whether FRCs upregulated VEGF expression after im-munization by using VEGF-lacZ reporter mice to compare VEGFmRNA expression before and after immunization. FDG cleavageand release of fluorescein were used to measure b-gal activity inthe gp38+ FRCs of VEGF-lacZ mice. Because there is a low rateof continued FDG cleavage over the time required to run a sampleon the flow cytometer, we ran the samples from the immunizedmice prior to running the samples from unimmunized mice. Thispotentially underestimated any increase in b-gal activity uponimmunization but ensured that any increase seen in the samplesfrom immunized mice was not an artifact of a longer incubationperiod. GP38+ FRCs at day 2 after OVA/CFA had greater b-galactivity than FRCs in unimmunized mice (Fig. 7A), and similarresults were obtained at day 3 and with BMDC-stimulated mice(data not shown).

CD11c+ cells have been shown to be localized near or directlyon FRCs (18, 21, 46, 47), and we have previously shown theCD11c+ cells are also enriched in regions rich in VEGF-expressing FRCs (9), raising the possibility that dendritic or otherCD11c+ cells could interact with FRCs to promote VEGF upreg-ulation. To test this possibility, we added BMDCs to culturesof 3T3 fibroblasts, which we have previously shown to share someof the characteristics of gp38+ FRCs, including gp38, lymphotoxinb receptor, and VEGF expression (9). BMDCs were depleted ofcontaminating granulocytes using magnetic selection to yielda dendritic cell-rich fraction (DC fraction) that was 90% pure anda granulocyte-rich fraction (Gr fraction) that was relatively den-dritic cell poor but still contained ∼30% dendritic cells (data notshown). At baseline, the dendritic cell and Gr fractions bothshowed very low levels of VEGF expression, whereas 3T3 cellsexpressed high levels of VEGF (Fig. 7C). Addition of the dendriticcell fraction to the 3T3 cells induced a .2-fold rise in VEGFlevels (Fig. 7C). This effect was specific for the DC fraction,because culturing of the fibroblasts with Gr fraction cells did not

FIGURE 4. CD11c+ cells mediate upregulation of proliferation of blood

and lymphatic endothelial cells and of FRCs during the initiation phase.

CD11c-DTR mice were injected with 100 ng DT or control inactive mutant

(CRM) toxin in the right footpad at 8 h before OVA/CFA injection in the

same footpad on day 0. Draining popliteal nodes were examined on day 2.

A, Flow cytometry plots showing relative depletion of all CD11c+ cell

subsets with DT. B, Number of cells per lymph node in the gates shown in

A. C, Proliferation rate of endothelial cell subsets as measured by BrdU

uptake. D, Proliferation rate of gp38+ FRCs. n = 6 mice/condition over four

experiments. *p , 0.05 and **p , 0.01 in comparison with the same cells

in CRM-treated mice using unpaired t test.

FIGURE 5. CCR7-dependent cells do not mediate upregulation of en-

dothelial and FRC proliferation during the initiation phase. WTor CCR72/2

mice were immunized with OVA/CFA on day 0 and draining popliteal nodes

and nondraining brachial nodes were examined on day 2. A, Lymph node

cellularity. B, Number of dendritic cells per lymph node in the gates shown

in C. C, Flow cytometry plot showing relative lack of CD11cmedMHCIIhi

cells in CCR72/2 popliteal lymph nodes at day 2 after OVA/CFA. D, Num-

ber of endothelial cells in all subsets. E, Proliferation rate of endothelial

cells. F, Number of gp38+ FRCs. G, Proliferation rate of gp38+ FRCs.

Comparisons were made between WT and CCR72/2 samples using upaired

t test. For proliferation determination, n = 5 mice/group over two experi-

ments; for lymph node, dendritic cell, endothelial cell, and FRC numbers, n =

at least seven mice per group over at least three experiments. *p , 0.05,

**p , 0.01 in comparison with the equivalent cells in WT mice.

5564 TWO PHASES OF REGULATING LYMPH NODE VASCULAR STROMAL GROWTH

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

result in VEGF increases (Fig. 7C). The fibroblasts showed up-regulated VEGF mRNA expression after exposure to the DC frac-tion (Fig. 7D). These results support the possibility that dendriticcell stimulation of FRCs may contribute to the increased VEGFlevels found in stimulated lymph nodes.

DiscussionOur results presented in this article further our understanding ofhow the vasculature grows after lymph node stimulation. Bloodvascular and lymphatic endothelial cells proliferate and expandin a concerted manner with FRCs, and there is a CD11c+ cell-dependent and T/B cell-independent initiation phase character-ized by upregulation of proliferation that is followed by a T andB cell-dependent expansion phase that results in greater num-bers of endothelial cells and FRCs. We further extend our under-standing of how CD11c+ cells may orchestrate the initiation phase,finding that skin-derived dendritic cells are not the requiredCD11c+ cells that mediate the initiation phase, that gp38+ FRCsare stimulated to upregulate VEGF after immunization, and thatdendritic cells can directly stimulate fibroblast-type cells to up-

regulate VEGF, suggesting the scenario that resident CD11c+ cellsupregulate blood vascular and lymphatic endothelial cell prolif-eration in part by stimulating FRCs to produce more VEGF.The T and B cell independence during the initiation phase,

followed by T and B cell dependence during the expansion phasemirrors the kinetics of innate and then adaptive immune cell ac-tivation during immune responses and suggests a scenario wherebyinnate immunity begins to stimulate vascular–stromal growth thatis only fully completed if there is Ag-specific activation of lym-phocytes. Upon activation, the lymphocytes promote vascular–stromal expansion and the vascular–stromal expansion presumablyensures that the microenviroment can fully support the generationof effector and memory T and B cells. More experiments will berequired to fully understand whether Ag-specific lymphocytesare necessary.B cells have been shown to mediate lymphatic expansion after

acute immunization (11, 12) and HEV expansion in a viral in-fection model (48), and our current findings showing a partialcontribution by B cells are consistent with these studies. CD4T cells are important for feeding arteriole remodeling (49) andsuppress lymphatic expansion (35), and our findings of T cellcontributions echo these findings. In this study, our experimentsreveal that B and T cells together contribute to mediate full vas-cular–stromal expansion and that there is partial redundancy orcompensation between T and B cells. The normal vascular ex-pansion in bd TCR-deficient mice suggested that B cells can fullycompensate for the lack of T cells, whereas the partial defect invascular expansion in B cell-deficient mice suggested that B cellspossess unique properties for which T cells can only partiallycompensate. For FRCs, expansion was partially affected in both Band T cell-deficient mice, suggesting additive effects or partialredundancy of B and T cells.The data were suggestive for a role for lymphocytes in mediating

vascular–stromal expansion by actively promoting cell survival inaddition to and somewhat independent of promoting cell prolif-eration. For example, although T cell-deficient mice had normalHEV and non-HEV blood vascular and FRC expansion, they hada partial proliferation defect, indicating that T cells make uniquecontributions to vascular–stromal proliferation. The normalexpansion in the face of reduced proliferation may reflect thepossibility that the proliferation defect was too mild to affectexpansion, or it may reflect a role for B cells in compensatingfully for the proliferation defect in part by promoting enhancedcell survival. Such a separation between cell proliferation and cellsurvival was also seen in the mMT mice, because FRCs showed

FIGURE 6. Depletion of skin-derived CD11c+ cells does not affect

lymph endothelial cell proliferation during the initiation phase. CD11c-

DTR mice were injected with 25 ng DT or CRM in the right footpad at 8 h

before OVA/CFA injection in the same footpad on day 0. Draining pop-

liteal nodes were examined on day 2. A, Flow cytometry plot showing

depletion of CD11cmedMHCIIhi cells with DT. B, Number of cells per

lymph node in the gates shown in A. C, Endothelial cell proliferation. *p,0.05 in comparison with same cells in CRM-treated mice using unpaired

t test; n = 6 mice/group over four experiments. CRM, control inactive

mutant.

FIGURE 7. VEGF is upregulated in FRCs in vivo and in fibroblasts cultured with dendritic cells. A and B, VEGF-lacZ mice were immunized with OVA/

CFA on day 0, and draining popliteal nodes were examined on day 2. b-Gal activity as an indicator of VEGF expression was measured as described in

Materials and Methods. A, Flow cytometry dot plots gated on CD452CD312 cells showing b-gal activity of gp38+ FRCs from an immunized WT mouse,

a sham-immunized VEGF-lacZ mouse, and an OVA/CFA-immunized VEGF-lacZ mouse. B, Graph shows mean fluorescence intensity of gp38+ FRCs in

the VEGF-lacZ mice. *p , 0.05 in comparison with unimmunized mice using paired t test. Each symbol represents one mouse, with matched symbols in

the unimmunized and OVA/CFA-immunized conditions representing a pair from a single experiment. Data were accumulated over four experiments. C and

D, 3T3 fibroblasts were cultured alone or with Gr-1 cell-depleted BMDCs (DC fraction) or the Gr1+ contaminating cells (Gr1 fraction) for 2 d. C, VEGF

levels in the culture supernatants. *p , 0.05 and **p , 0.01 t test. Each condition was plated in triplicate wells; bars, average value of the triplicate wells.

Results representative of at least five similar experiments. D, VEGF mRNA level in the 3T3 cells after culturing without or with DC fraction, as measured

by real-time PCR. Each bar represents average values of two experiments, with each symbol representing the value obtained in a single experiment.

The Journal of Immunology 5565

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

reduced expansion but WT levels of proliferation. Similarly, inRAG12/2 mice, the initial drive for FRC proliferation can be seento be intact even at day 5, but the expansion is completely pre-vented. Further work will be needed to test this idea, but cellproliferation and survival can be regulated by different sets ofmolecules (e.g., cyclins and Bcl-2 family members, respectively)and have been shown to be distinctly regulated. A recent exampleis in the case of mice deficient for p18, a negative modulator ofcell cycling, where plasmablasts proliferate highly but do notsurvive (50). Also, in the dissection of the role of the antiapoptoticmolecule survivin in balloon-mediated arterial injury, survivin wasrequired for platelet-derived growth factor-mediated cell survivalbut not for mitosis (51). It is possible that T and B cells can ac-tively regulate both cell proliferation and cell survival to promotevascular–stromal expansion.We have previously shown that CD11c+ cells are required for the

initiation of lymph node vascular growth (8), and in this article,we extend our findings to show that CD11c+ cells mediate up-regulation of blood vascular as well as lymphatic endothelial celland FRC proliferation. These results suggest that CD11c+ cellsinitiate the coordinated growth of the cellular elements of thevascular–stromal network. These findings may have implicationsfor understanding the role of CD11c+ cells in the growth of othertissues. Recent studies have shown that, in inflamed airways, de-pletion of CD11c+ cells during the period of induced BALT(iBALT) growth did not affect the number of iBALTs but resultedin smaller iBALTs (52). This role for CD11c+ cells in mediatingiBALT growth could potentially in part reflect the contribution ofinitiating CD11c+ cells to the growth of the vascular–stromalnetwork in these lymph node-like structures. Furthermore, the ben-eficial effect of adding dendritic cells to a tissue-engineeredlymph node (53, 54) may reflect increased dendritic cell-drivenproliferation of the fibroblast-type cells and lymphatics as well assupport blood vessel growth. In addition, CD11c+ cells have beenimplicated in vascular expansion in inflamed nonlymphoid tissuesand tumors (55–58), and our results suggest that CD11c+ cellsmay support the growth of these tissues beyond that of the bloodvascular endothelial cells.In this study, we have narrowed down the candidates for the

CD11c+ cells that mediate the initiation phase. CD11cmedMHCIIhi

cells include migratory dendritic cells that require CCR7 to mi-grate from skin and may also include monocyte-derived dendriticcells that enter lymph nodes via blood (31, 37, 59). Our resultsshowed that neither preventing lymph node entry of CD11cmed

MHCIIhi cells nor selectively depleting skin-derived CD11cmed

MHCIIhi cells resulted in reduced vascular–stromal proliferationand suggest that skin-derived dendritic cells likely do not mediatethe upregulation of vascular–stromal proliferation during the ini-tiation phase. These results combined with our previous resultsshowing only a partial role for CD11chiMHCIImed cells suggestthat a subpopulation of CD11cmedMHCIImed cells residing inlymph nodes is important for mediating the events of the initiationphase; experiments are ongoing to test this hypothesis.We show in this article that FRCs upregulate VEGF mRNA

expression at day 2 and that, in vitro, dendritic cells can stimulatefibroblast-type cells to upregulate VEGF expression. Our resultstogether suggest a scenario whereby, during the initiation phase,lymph node-resident CD11c+ cells stimulate FRCs to upregulateVEGF, which, in turn, induces upregulation of blood vascular andlymphatic endothelial cell proliferation. GP38+ FRCs can alsoexpress the lymphangiogenic molecule VEGF-C (60), and it islikely that CD11c+ cells alter FRC activity in additional ways tocontribute to the initiation of vascular expansion. Dendritic cellsare closely associated with the reticular network (18, 46, 47), and

our coculture experiments suggest that direct communication withFRCs could lead to VEGF upregulation. We have also shown arole for recruited blood-borne L-selectin–dependent cells in me-diating a portion of the dendritic cell-driven endothelial cellproliferation (8), so it is possible that CD11c+ cells also promoteVEGF upregulation in part by first promoting recruitment of L-selectin+ cells via increased blood delivery (5), increased vascularpermeability (7), or altered HEV cell adhesion molecule or che-mokine expression (12, 21, 61–63). Our results suggest that stim-ulation of blood and lymphatic vascular proliferation is mediatedby at least a tripartite interaction among CD11c+ cells, FRCs,and the endothelial cells.Taken together, our results add to the emerging concept that there

is a constellation of vascular–stromal alterations that occur withthe induction of an immune response in lymph nodes and high-light the coordinated regulation of vascular and stromal expansionby first innate and then adaptive immune cells. Lymph nodes areenlarged, and the vascular–stromal compartment can potentiallybe altered in autoimmune and lymphoproliferative diseases. Un-derstanding the extent of vascular–stromal alterations in stimu-lated lymph nodes, how these alterations are regulated and func-tion in immunity may lead to the targeting of the vascular–stromalcompartment in the development of novel therapeutics.

AcknowledgmentsWe thank all the members of the Lu laboratory for helpful discussions,

Dr. Sanjiv Luther for critical reading of the manuscript, Drs. Miriam Merad

and BernardMalissen for providing Langerin-DTRmice, and Dr. Sergio Lira

for providing CCR7 breeders.

DisclosuresThe authors have no financial conflicts of interest.

References1. von Andrian, U. H., and T. R. Mempel. 2003. Homing and cellular traffic in

lymph nodes. Nat. Rev. Immunol. 3: 867–878.2. Mueller, S. N., and R. N. Germain. 2009. Stromal cell contributions to the ho-

meostasis and functionality of the immune system. Nat. Rev. Immunol. 9: 618–629.

3. Lammermann, T., and M. Sixt. 2008. The microanatomy of T-cell responses.Immunol. Rev. 221: 26–43.

4. Junt, T., E. Scandella, and B. Ludewig. 2008. Form follows function: lymphoidtissue microarchitecture in antimicrobial immune defence. Nat. Rev. Immunol. 8:764–775.

5. Soderberg, K. A., G. W. Payne, A. Sato, R. Medzhitov, S. S. Segal, and A. Iwasaki.2005. Innate control of adaptive immunity via remodeling of lymph node feedarteriole. Proc. Natl. Acad. Sci. USA 102: 16315–16320.

6. Herman, P. G., I. Yamamoto, and H. Z. Mellins. 1972. Blood microcirculation inthe lymph node during the primary immune response. J. Exp. Med. 136: 697–714.

7. Anderson, N. D., A. O. Anderson, and R. G. Wyllie. 1975. Microvascularchanges in lymph nodes draining skin allografts. Am. J. Pathol. 81: 131–160.

8. Webster, B., E. H. Ekland, L. M. Agle, S. Chyou, R. Ruggieri, and T. T. Lu.2006. Regulation of lymph node vascular growth by dendritic cells. J. Exp. Med.203: 1903–1913.

9. Chyou, S., E. H. Ekland, A. C. Carpenter, T.-C. J. Tzeng, S. Tian, M. Michaud,J. A. Madri, and T. T. Lu. 2008. Fibroblast-type reticular stromal cells regulatethe lymph node vasculature. J. Immunol. 181: 3887–3896.

10. Mebius, R. E., J. Breve, A. M. Duijvestijn, and G. Kraal. 1990. The function ofhigh endothelial venules in mouse lymph nodes stimulated by oxazolone. Im-munology 71: 423–427.

11. Angeli, V., F. Ginhoux, J. Llodra, L. Quemeneur, P. S. Frenette, M. Skobe,R. Jessberger, M. Merad, and G. J. Randolph. 2006. B cell-driven lymphangio-genesis in inflamed lymph nodes enhances dendritic cell mobilization. Immunity24: 203–215.

12. Liao, S., and N. H. Ruddle. 2006. Synchrony of high endothelial venules andlymphatic vessels revealed by immunization. J. Immunol. 177: 3369–3379.

13. Halin, C., N. E. Tobler, B. Vigl, L. F. Brown, and M. Detmar. 2007. VEGF-Aproduced by chronically inflamed tissue induces lymphangiogenesis in draininglymph nodes. Blood 110: 3158–3167.

14. Katakai, T., T. Hara, M. Sugai, H. Gonda, and A. Shimizu. 2004. Lymph nodefibroblastic reticular cells construct the stromal reticulum via contact with lym-phocytes. J. Exp. Med. 200: 783–795.

5566 TWO PHASES OF REGULATING LYMPH NODE VASCULAR STROMAL GROWTH

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from

15. Gretz, J. E., A. O. Anderson, and S. Shaw. 1997. Cords, channels, corridors andconduits: critical architectural elements facilitating cell interactions in the lymphnode cortex. Immunol. Rev. 156: 11–24.

16. Cyster, J. G. 2005. Chemokines, sphingosine-1-phosphate, and cell migration insecondary lymphoid organs. Annu. Rev. Immunol. 23: 127–159.

17. Link, A., T. K. Vogt, S. Favre, M. R. Britschgi, H. Acha-Orbea, B. Hinz,J. G. Cyster, and S. A. Luther. 2007. Fibroblastic reticular cells in lymph nodesregulate the homeostasis of naive T cells. Nat. Immunol. 8: 1255–1265.

18. Bajenoff, M., J. G. Egen, L. Y. Koo, J. P. Laugier, F. Brau, N. Glaichenhaus, andR. N. Germain. 2006. Stromal cell networks regulate lymphocyte entry, migra-tion, and territoriality in lymph nodes. Immunity 25: 989–1001.

19. Lee, J.-W., M. Epardaud, J. Sun, J. E. Becker, A. C. Cheng, A.-r. Yonekura,J. K. Heath, and S. J. Turley. 2007. Peripheral antigen display by lymph nodestroma promotes T cell tolerance to intestinal self. Nat. Rev. Immunol. 8: 181–190.

20. Gretz, J. E., C. C. Norbury, A. O. Anderson, A. E. Proudfoot, and S. Shaw. 2000.Lymph-borne chemokines and other low molecular weight molecules reach highendothelial venules via specialized conduits while a functional barrier limitsaccess to the lymphocyte microenvironments in lymph node cortex. J. Exp. Med.192: 1425–1440.

21. Tzeng, T. C., S. Chyou, S. Tian, B. Webster, A. C. Carpenter, V. H. Guaiquil, andT. T. Lu. 2010. CD11chi dendritic cells regulate the re-establishment of vascularquiescence and stabilization after immune stimulation of lymph nodes. J.Immunol. 184: 4247–4257.

22. Breiteneder-Geleff, S., A. Soleiman, H. Kowalski, R. Horvat, G. Amann,E. Kriehuber, K. Diem, W. Weninger, E. Tschachler, K. Alitalo, and D. Kerjaschki.1999. Angiosarcomas express mixed endothelial phenotypes of blood and lym-phatic capillaries: podoplanin as a specific marker for lymphatic endothelium. Am.J. Pathol. 154: 385–394.

23. Farr, A. G., M. L. Berry, A. Kim, A. J. Nelson, M. P. Welch, and A. Aruffo. 1992.Characterization and cloning of a novel glycoprotein expressed by stromal cells inT-dependent areas of peripheral lymphoid tissues. J. Exp. Med. 176: 1477–1482.

24. Roozendaal, R., T. R. Mempel, L. A. Pitcher, S. F. Gonzalez, A. Verschoor,R. E. Mebius, U. H. von Andrian, and M. C. Carroll. 2009. Conduits mediatetransport of low-molecular-weight antigen to lymph node follicles. Immunity 30:264–276.

25. Shrestha, B., T. Hashiguchi, T. Ito, N. Miura, K. Takenouchi, Y. Oyama,K. Kawahara, S. Tancharoen, Y. Ki-I, N. Arimura, et al. 2010. B cell-derivedvascular endothelial growth factor A promotes lymphangiogenesis and highendothelial venule expansion in lymph nodes. J. Immunol. 184: 4819–4826.

26. Drayson, M. T., and W. L. Ford. 1984. Afferent lymph and lymph borne cells:their influence on lymph node function. Immunobiology 168: 362–379.

27. Hendriks, H. R., I. L. Eestermans, and E. C. Hoefsmit. 1980. Depletion ofmacrophages and disappearance of postcapillary high endothelial venules inlymph nodes deprived of afferent lymphatic vessels. Cell Tissue Res. 211:375–389.

28. Mebius, R. E., P. R. Streeter, J. Breve, A. M. Duijvestijn, and G. Kraal. 1991.The influence of afferent lymphatic vessel interruption on vascular addressinexpression. J. Cell Biol. 115: 85–95.

29. Mombaerts, P., J. Iacomini, R. S. Johnson, K. Herrup, S. Tonegawa, andV. E. Papaioannou. 1992. RAG-1–deficient mice have no mature B and T lym-phocytes. Cell 68: 869–877.

30. Jung, S., D. Unutmaz, P. Wong, G. Sano, K. De los Santos, T. Sparwasser, S. Wu,S. Vuthoori, K. Ko, F. Zavala, et al. 2002. In vivo depletion of CD11c+ dendriticcells abrogates priming of CD8+ T cells by exogenous cell-associated antigens.Immunity 17: 211–220.

31. Forster, R., A. Schubel, D. Breitfeld, E. Kremmer, I. Renner-Muller, E. Wolf,and M. Lipp. 1999. CCR7 coordinates the primary immune response by estab-lishing functional microenvironments in secondary lymphoid organs. Cell 99:23–33.

32. Kissenpfennig, A., S. Henri, B. Dubois, C. Laplace-Builhe, P. Perrin, N. Romani,C. H. Tripp, P. Douillard, L. Leserman, D. Kaiserlian, et al. 2005. Dynamics andfunction of Langerhans cells in vivo: dermal dendritic cells colonize lymph nodeareas distinct from slower migrating Langerhans cells. Immunity 22: 643–654.

33. Miquerol, L., M. Gertsenstein, K. Harpal, J. Rossant, and A. Nagy. 1999. Mul-tiple developmental roles of VEGF suggested by a LacZ-tagged allele. Dev. Biol.212: 307–322.

34. Kruisbeek, A. M. 2001. In vivo depletion of CD4- and CD8-specific T cells.Curr. Protoc. Immunol. Chp. 4, Unit 4.1.

35. Kataru, R. P., H. Kim, C. Jang, D. K. Choi, B. I. Koh, M. Kim, S. Gollamudi,Y. K. Kim, S. H. Lee, and G. Y. Koh. 2011. T lymphocytes negatively regulatelymph node lymphatic vessel formation. Immunity 34: 96–107.

36. Lopez-Bravo, M., and C. Ardavın. 2008. In vivo induction of immune responsesto pathogens by conventional dendritic cells. Immunity 29: 343–351.

37. Ohl, L., M. Mohaupt, N. Czeloth, G. Hintzen, Z. Kiafard, J. Zwirner,T. Blankenstein, G. Henning, and R. Forster. 2004. CCR7 governs skin dendriticcell migration under inflammatory and steady-state conditions. Immunity 21:279–288.

38. Okada, T., V. N. Ngo, E. H. Ekland, R. Forster, M. Lipp, D. R. Littman, andJ. G. Cyster. 2002. Chemokine requirements for B cell entry to lymph nodes andPeyer’s patches. J. Exp. Med. 196: 65–75.

39. Bogunovic, M., F. Ginhoux, A. Wagers, M. Loubeau, L. M. Isola, L. Lubrano,V. Najfeld, R. G. Phelps, C. Grosskreutz, E. Scigliano, et al. 2006. Identificationof a radio-resistant and cycling dermal dendritic cell population in mice andmen. J. Exp. Med. 203: 2627–2638.

40. Merad, M., M. G. Manz, H. Karsunky, A. Wagers, W. Peters, I. Charo,I. L. Weissman, J. G. Cyster, and E. G. Engleman. 2002. Langerhans cells renew

in the skin throughout life under steady-state conditions. Nat. Immunol. 3: 1135–1141.

41. Ginhoux, F., M. P. Collin, M. Bogunovic, M. Abel, M. Leboeuf, J. Helft,J. Ochando, A. Kissenpfennig, B. Malissen, M. Grisotto, et al. 2007. Blood-derived dermal langerin+ dendritic cells survey the skin in the steady state. J.Exp. Med. 204: 3133–3146.

42. Bursch, L. S., L. Wang, B. Igyarto, A. Kissenpfennig, B. Malissen, D. H. Kaplan,and K. A. Hogquist. 2007. Identification of a novel population of Langerin+

dendritic cells. J. Exp. Med. 204: 3147–3156.43. Poulin, L. F., S. Henri, B. de Bovis, E. Devilard, A. Kissenpfennig, and

B. Malissen. 2007. The dermis contains langerin+ dendritic cells that developand function independently of epidermal Langerhans cells. J. Exp. Med. 204:3119–3131.

44. Palframan, R. T., S. Jung, G. Cheng, W. Weninger, Y. Luo, M. Dorf,D. R. Littman, B. J. Rollins, H. Zweerink, A. Rot, and U. H. von Andrian. 2001.Inflammatory chemokine transport and presentation in HEV: a remote controlmechanism for monocyte recruitment to lymph nodes in inflamed tissues. J. Exp.Med. 194: 1361–1373.

45. Kunder, C. A., A. L. St John, G. Li, K. W. Leong, B. Berwin, H. F. Staats, andS. N. Abraham. 2009. Mast cell-derived particles deliver peripheral signals toremote lymph nodes. J. Exp. Med. 206: 2455–2467.

46. Luther, S. A., H. L. Tang, P. L. Hyman, A. G. Farr, and J. G. Cyster. 2000.Coexpression of the chemokines ELC and SLC by T zone stromal cells anddeletion of the ELC gene in the plt/plt mouse. Proc. Natl. Acad. Sci. USA 97:12694–12699.

47. Sixt, M., N. Kanazawa, M. Selg, T. Samson, G. Roos, D. P. Reinhardt, R. Pabst,M. B. Lutz, and L. Sorokin. 2005. The conduit system transports soluble anti-gens from the afferent lymph to resident dendritic cells in the T cell area of thelymph node. Immunity 22: 19–29.

48. Kumar, V., E. Scandella, R. Danuser, L. Onder, M. Nitschke, Y. Fukui, C. Halin,B. Ludewig, and J. V. Stein. 2010. Global lymphoid tissue remodeling duringa viral infection is orchestrated by a B cell-lymphotoxin–dependent pathway.Blood 115: 4725–4733.

49. Kumamoto, Y., L. M. Mattei, S. Sellers, G. W. Payne, and A. Iwasaki. 2011.CD4+ T cells support cytotoxic T lymphocyte priming by controlling lymphnode input. Proc. Natl. Acad. Sci. USA 108: 8749–8754.

50. Bretz, J., J. Garcia, X. Huang, L. Kang, Y. Zhang, K.-M. Toellner, and S. Chen-Kiang. 2011. Noxa mediates p18INK4c cell-cycle control of homeostasis inB cells and plasma cell precursors. Blood 117: 2179–2188.

51. Blanc-Brude, O. P., J. Yu, H. Simosa, M. S. Conte, W. C. Sessa, andD. C. Altieri. 2002. Inhibitor of apoptosis protein survivin regulates vascularinjury. Nat. Med. 8: 987–994.

52. Halle, S., H. C. Dujardin, N. Bakocevic, H. Fleige, H. Danzer, S. Willenzon,Y. Suezer, G. Hammerling, N. Garbi, G. Sutter, et al. 2009. Induced bronchus-associated lymphoid tissue serves as a general priming site for T cells and ismaintained by dendritic cells. J. Exp. Med. 206: 2593–2601.

53. Suematsu, S., and T. Watanabe. 2004. Generation of a synthetic lymphoid tissue-like organoid in mice. Nat. Biotechnol. 22: 1539–1545.

54. Okamoto, N., R. Chihara, C. Shimizu, S. Nishimoto, and T. Watanabe. 2007.Artificial lymph nodes induce potent secondary immune responses in naive andimmunodeficient mice. J. Clin. Invest. 117: 997–1007.

55. Fainaru, O., N. Almog, C. W. Yung, K. Nakai, M. Montoya-Zavala,A. Abdollahi, R. D’Amato, and D. E. Ingber. 2010. Tumor growth and angio-genesis are dependent on the presence of immature dendritic cells. FASEB J. 24:1411–1418.

56. Fainaru, O., A. Adini, O. Benny, I. Adini, S. Short, L. Bazinet, K. Nakai,E. Pravda, M. D. Hornstein, R. J. D’Amato, and J. Folkman. 2008. Dendriticcells support angiogenesis and promote lesion growth in a murine model ofendometriosis. FASEB J. 22: 522–529.

57. Nakai, K., O. Fainaru, L. Bazinet, P. Pakneshan, O. Benny, E. Pravda,J. Folkman, and R. J. D’Amato. 2008. Dendritic cells augment choroidal neo-vascularization. Invest. Ophthalmol. Vis. Sci. 49: 3666–3670.

58. Conejo-Garcia, J. R., F. Benencia, M. C. Courreges, E. Kang, A. Mohamed-Hadley, R. J. Buckanovich, D. O. Holtz, A. Jenkins, H. Na, L. Zhang, et al. 2004.Tumor-infiltrating dendritic cell precursors recruited by a b-defensin contributeto vasculogenesis under the influence of Vegf-A. Nat. Med. 10: 950–958.

59. Cheong, C., I. Matos, J. H. Choi, D. B. Dandamudi, E. Shrestha, M. P. Longhi,K. L. Jeffrey, R. M. Anthony, C. Kluger, G. Nchinda, et al. 2010. Microbialstimulation fully differentiates monocytes to DC-SIGN/CD209+ dendritic cellsfor immune T cell areas. Cell 143: 416–429.

60. Peduto, L., S. Dulauroy, M. Lochner, G. F. Spath, M. A. Morales, A. Cumano,and G. Eberl. 2009. Inflammation recapitulates the ontogeny of lymphoid stro-mal cells. J. Immunol. 182: 5789–5799.

61. Chen, Q., D. T. Fisher, K. A. Clancy, J. M. Gauguet, W. C. Wang, E. Unger,S. Rose-John, U. H. von Andrian, H. Baumann, and S. S. Evans. 2006. Fever-range thermal stress promotes lymphocyte trafficking across high endothelialvenules via an interleukin 6 trans-signaling mechanism. Nat. Immunol. 7: 1299–1308.

62. Guarda, G., M. Hons, S. F. Soriano, A. Y. Huang, R. Polley, A. Martın-Fontecha,J. V. Stein, R. N. Germain, A. Lanzavecchia, and F. Sallusto. 2007. L-selectin–negative CCR7– effector and memory CD8+ T cells enter reactive lymph nodesand kill dendritic cells. Nat. Immunol. 8: 743–752.

63. Martın-Fontecha, A., D. Baumjohann, G. Guarda, A. Reboldi, M. Hons,A. Lanzavecchia, and F. Sallusto. 2008. CD40L+CD4+ memory T cells migratein a CD62P-dependent fashion into reactive lymph nodes and license dendriticcells for T cell priming. J. Exp. Med. 205: 2561–2574.

The Journal of Immunology 5567

by guest on May 1, 2018

http://ww

w.jim

munol.org/

Dow

nloaded from