Visualizing dendritic cell networks in vivo

Randall L Lindquist1,4, Guy Shakhar2,4, Diana Dudziak1, Hedda Wardemann1, Thomas Eisenreich1,3,Michael L Dustin2 & Michel C Nussenzweig1,3

In the steady state, dendritic cells (DCs) in the lymph node induce T cell tolerance to self antigens. Innate signals trigger the

maturation of tissue DCs, which migrate into lymph nodes and activate T cells. To examine DCs in vivo, we produced transgenic

mice whose DCs expressed enhanced yellow fluorescent protein. Two-photon microscopy of lymph nodes in live mice showed

that most of the steady-state DCs were enmeshed in an extensive network and remained in place while actively probing adjacent

T cells with their processes. Mature DCs were more motile than steady-state DCs and were rapidly dispersed and integrated into

the sessile network, facilitating their interaction with migrating T cells.

Dendritic cells (DCs) are specialized antigen-presenting cells thatare found in nonlymphoid and lymphoid organs and are essentialin initiating immune responses and maintaining tolerance1–6. Innonlymphoid organs, DCs serve as sentinels that capture and thencarry antigens to lymphoid organs, where they interact with specificT cells. In contrast, DCs that reside in lymphoid organs such aslymph nodes receive antigens from lymph, blood or immigrant DCs,relaying antigen from the periphery7–11. The outcome of antigenpresentation by DCs depends on the state of DC terminal differentia-tion or maturation1–6. In the steady state, immature DCs capture,process and present a variety of self antigens to T cells; sources ofantigen include serum proteins, extracellular matrix componentsand dead cells11–18. Antigen presentation by steady-state DCs estab-lishes T cell tolerance to prevent adverse reactions to self whenthe same antigens are presented in immunizing conditions6. Incontrast, DCs that have been stimulated to mature by inflammatorymediators, such as Toll-like receptor ligands, induce strong effectorT cell responses1–6.

Two-photon microscopy has been used to study mature DCsactively participating in immune responses in explanted intactlymph nodes or surgically exposed lymph nodes in anaesthetizedmice19–22. DCs were labeled with exogenous fluorescent dyes, and DCmaturation was induced during isolation or by deliberate treatmentwith adjuvants19–22. In those studies mature DCs ‘crawled’ in randomdirections, albeit at a slower speed than T cells, and they rapidlyextended and retracted long dendritic processes. However, steady-stateDCs were not examined.

To examine DC activity in the steady state, we produced transgenicmice expressing enhanced yellow fluorescent protein (EYFP) inCD11c+ DCs. We selected founders that had sufficient expressionfor imaging deep in the lymph node by two-photon laser-scanningmicroscopy and verified that the expression was restricted to the

appropriate cells. Here we report on the activity of steady-state andmature DCs in various regions in the lymph nodes of live mice.

RESULTS

CD11c-EYFP–transgenic mice

We produced transgenic mice expressing an EYFP23 reporter under thecontrol of the CD11c promoter24. We examined intact explantedlymph nodes from the progeny of two of the founders by two-photonfluorescence microscopy to determine their suitability for deep tissueimaging. CD11c-EYFPhi mice had many bright fluorescent cells atdepths of 175–300 mm; these cells were not visible in CD11c-EYFPlo

mice (Fig. 1a and Supplementary Videos 1 and 2 online). Inagreement with results of the two-photon experiments, brightlystained cells (EYFP+ cells) were also present in frozen sections fromlymph nodes only in the CD11c-EYFPhi mice, and these cells expressedendogenous CD11c, as determined by immunofluorescence(Fig. 1b,c). By flow cytometry, we found that the EYFP+ cells brightenough to be visualized intravitally were CD11c+CD19�CD3�CD86+

major histocompatibility complex (MHC) class II–positive DCs(Fig. 1d and Supplementary Fig. 1 online). A small percentage of Bcells and T cells also expressed EYFP, but the amount of EYFP expressedby these cells was at least one log lower than that expressed by DCs andwas consequently below the threshold of detection in the microscope(determined by comparison with CD11c-EYFPlo; Fig. 1 and Supple-mentary Fig. 1 online). Thus, the fluorescent cells visualized by two-photon microscopy in CD11c-EYFPhi mice corresponded to DCs withhigh expression of CD11c, which could be visualized in lymph nodes atdepths of up to 300 mm (Supplementary Videos 1 and 2 online).

EYFP+ DC distribution in lymph nodes

In lymph node sections, many EYFP+ DCs were present in theperifollicular zone and T cell area, and occasional cells were present

Published online 14 November 2004; doi:10.1038/ni1139

1Laboratory of Molecular Immunology, 3Howard Hughes Medical Institute, The Rockefeller University, New York, New York 10021, USA. 2Program in MolecularPathogenesis and Department of Pathology, Skirball Institute of Biomolecular Medicine, New York University School of Medicine, New York, New York 10016, USA.4These authors contributed equally to this work. Correspondence should be addressed to M.C.N. (nussen@mail.rockefeller.edu) or M.L.D. (dustin@saturn.med.nyu.edu).

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in the B cell follicle and subcapsular space (Figs. 1 and 2); these cellswere not macrophages or granulocytes because they did not have highexpression of the cell surface markers MOMA-1 (Fig. 2a), CD11b(Fig. 2b), F4/80 (Fig. 2c) or Gr-1 (Fig. 2d). Unlike mature DCs thatrecently immigrated to the lymph node20,25, steady-state DCs were notpreferentially associated with high endothelial venules (HEVs; Fig. 2e,fand Supplementary Video 3 online). By immunofluorescence, EYFP+

DCs were heterogeneous for expression of MHC class II, the DCmarker DEC-205 and CD86 (Fig. 2g–i). There was a partial overlapbetween populations of cells expressing EYFP and MHC class II(Fig. 2g) as well as between populations that expressed EYFP andDEC-205 (Fig. 2h). As noted above, by flow cytometry, which is a

more sensitive technique, all EYFP+ cells expressed surface MHC classII, but they showed a broad range of expression of this antigen (Fig. 1dand Supplementary Fig. 1 online). We conclude that transgenicCD11c-EYFP DCs resemble their wild-type counterparts in expressingheterogeneous amounts of traditional markers of DC maturation26,27.

In vivo imaging of steady-state DCs

We used two-photon intravital microscopy to examine the inguinallymph nodes of anaesthetized mice. We maintained the inguinallymph node at 37 1C during imaging using a stage warmer and anobjective heater and confirmed the temperature with a temperatureprobe close to the lymph node. This system enabled the acquisition of

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Figure 1 Brightness and surface markers of CD11c-EYFP cells. (a) Two-photon fluorescence images descending along the z-axis through explanted lymph

nodes from CD11c-EYFPhi and CD11c-EYFPlo mice, with EYFP in green and second harmonics from collagen fibers in blue; depth from the apex of the

lymph node (in mm) is above the images. Only CD11c-EYFPhi cells can be visualized deep in the lymph node. Scale bars, 50 mm. (b) Fluorescence

micrographs of lymph node sections from wild-type (WT), CD11c-EYFPlo and CD11c-EYFPhi mice stained with anti-CD3 (red) and anti-B220 (blue); EYFP

fluorescence was visualized directly. Scale bars, 200 mm. (c) Confocal micrographs of lymph node sections immunostained for CD11c in red, with EYFP

fluorescence in green. Scale bars, 10 mm. (d) Flow cytometry histograms showing surface marker expression on EYFP+ cells from CD11c-EYFPhi mice,

gated on cells bright enough to detect intravitally (Supplementary Fig. 1 online). Results are representative of at least two experiments. MHC II, MHC

class II. Control, isotype control antibody.

EYFP MOMA-1 B220 EYFP CD11b CD3 EYFP Gr-1 B220 EYFP PECAM B220 EYFP HEV CD3EYFP F4/80 B220

EYFP CD86 EYFP CD86EYFP DEC-205 B220 EYFP DEC-205 B220EYFP MHC II EYFP MHC II

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Figure 2 CD11c-EYFP mice examined by immunofluorescence. Confocal micrographs of lymph node sections stained for various cell surface markers in

red (above images) and counterstained in blue with CD3 or B220 to provide anatomical definition in the lymph node. Squares linked to arrows indicate

areas magnified at right. Scale bars, 200 mm (low-resolution images) or 100 mm (high-magnification insets). Results are representative of at least

three experiments.

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three-dimensional volumes in 15–30 s, including collagen fibersvisualized through second-harmonic imaging (SupplementaryVideo 4 online) and three fluorescence channels including enhancedcyan fluorescent protein (ECFP), enhanced green fluorescent protein(EGFP) and EYFP (Fig. 3a and Supplementary Video 3 online).Blood flow was verified by intravenous injection of rhodamine dextran(Fig. 3b and Supplementary Video 3 online). We found abundantrolling and attached cells in blood vessels that we verified as HEVs byin vivo direct staining with Alexa 546–conjugated MECA-79 antibody,an HEV marker (Fig. 3c), or by morphological criteria (Supplemen-tary Video 3 online). Lymph flow was also maintained, based ondrainage of subcutaneously injected rhodamine dextran through theefferent lymphatics (data not shown). Our blood and lymph flowresults were similar to those reported before28 and verified that theinguinal lymph node preparation could be used to assess the in vivodynamics of DCs in physiological conditions. Finally, we also exam-ined popliteal lymph nodes20 and found no differences in DC activityin the two preparations (data not shown).

Two-photon intravital microscopy of the inguinal lymph nodeshowed distinct populations of DCs (Fig. 4 and SupplementaryVideo 1 online). The most superficial EYFP cells were in thesubcapsular sinus (Fig. 4a). These DC had few dendrites and multiplelarge ruffles, which they extended and retracted rapidly (Supplemen-tary Videos 5 and 6 online). Subcapsular DCs were intercalatedamong collagen fibers (visualized by second harmonics) and relatively

sessile phagocytic macrophages, visualized by their uptake of66-kilodalton rhodamine-dextran29. The subcapsular DCs resembledEYFP cells on serosal surfaces outside the lymph node (Supplemen-tary Video 7 online), which may suggest that they were recentimmigrants from the periphery30.

In contrast, most of the DCs in the lymph node parenchyma werein extensive networks that are difficult to appreciate by conventionalhistology or intravital studies of immigrant mature DCs19–21. T cellzones and B cell follicles were demarcated by injection of CD11c-EYFPmice with EGFP-expressing T or B cells, respectively. DCs in deepparacortical networks interspersed with T cells showed extensive‘probing’, and more than 95% seemed to be firmly tethered to thenetwork and were thus sessile (Fig. 4b and Supplementary Videos 8and 9 online).

Three different populations of DCs were associated with the B cellfollicles: sessile perifollicular DCs between the subcapsular space andB cell follicle; DC clusters at the junction of T and B cell zones; andscattered DCs in the B cell follicle. The perifollicular DCs (Fig. 4cand Supplementary Videos 4 and 10 online) were relatively ‘dim’for EYFP, showed ‘probing’ motions and seemed well positioned toacquire antigens from lymph. In the junction between the T and B cell

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Figure 4 Anatomic distribution and activity of DCs in the lymph node. (a–f) Inguinal lymph nodes from CD11c-EYFP mice examined by intravital microscopy.

EYFP fluorescence is in green. Scale bars, 50 mm. (a) Subcapsular DCs surrounded by macrophages that have taken up rhodamine-labeled dextran. (b) DCs

in the T cell area (T-zone) of the lymph nodes, with transferred EGFP+ T cells ‘false-colored’ red. (c) DCs form a network surrounding the B cell follicles.

(d) DC cluster at the boundary between T cell and B cell areas (T-B interface). (e) Inguinal lymph node diagram indicates the anatomical locations of the

DCs in the surrounding images. (f) DCs in a B cell follicle (B-zone), with transferred EGFP+ B cells ‘false-colored’ blue. (g) ‘Crawling’ speeds of DCs vary

with anatomical location. Each point represents the average speed of one cell as calculated by the movement of the centroid; cells with velocities less than

2 mm/min are considered sessile. Horizontal lines indicate median speeds. Cells tend to arrest when incorporated into cell networks, although individual

fast-moving cells do occur in all locations. Results summarize at least three imaging sessions in each location.

Figure 3 Two-photon intravital microscopy of

inguinal lymph nodes. (a) Three-dimensional

reconstruction from two-photon microscopy data

showing the anatomic distribution of perifollicular

and deeper EYFP+ DCs (green) relative to the

lymph node capsule (second-harmonics signals

from collagen are in blue). (b) Three-dimensional

reconstruction from two-photon microscopy datawith EYFP+ DCs (green) and blood vessels (red)

visualized by intravenous injection of rhodamine-

labeled dextran. (c) Maximum intensity projection

of a 20-mm-thick volume showing EYFP+ DCs and

HEVs (red) visualized by injection of labeled

MECA-79 antibody. Scale bar, 50 mm.

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zones, DCs were present in sessile clusters that varied in size up to80 mm across and contained sessile EYFP– lymphocytes (Fig. 4d andSupplementary Video 11 online; discussed below); DCs in clustershad very dynamic membrane extensions and seemed to activelycapture lymphoid cells from their surroundings (SupplementaryVideo 12 online). EYFP-bright DCs were sparsest in the B cell folliclesbut they were on average the most motile DC population in the lymphnode (Fig. 4f,g). We conclude that in the steady state, DCs in lymphnodes form previously unappreciated groups of cells that can bedistinguished based on their activity. These include a minority ofhighly motile discrete cells and extensive assemblies of interconnectedsessile cells in networks and clusters.

Quantitative analysis of steady-state DC movement

Measurement of DC motility is challenging because active ‘probing’ ofthe DC means that the three-dimensional center of fluorescence(centroid) for a DC is continually and rapidly shifting independentof displacement (actual movement through the tissue). This can beshown with different colors to demonstrate the outline of a single DCat 30-second intervals (Supplementary Fig. 2 online). Because thesemovements are rapid, the rate at which three-dimensional imagevolumes are acquired in the tissue has a large effect on the measuredinstantaneous and average speeds. This makes it difficult to comparespeed measurements between different studies and also means thatstudies with high sampling rates may overestimate DC speed.To derive speeds that are related to displacement in the tissue,we applied a filter that rejected small movements in each stepand only recorded movements that were greater than a threshold of

2.5 mm (Supplementary Fig. 2 online).Although we obtained three-dimensionaldata sets every 30 s, this filter made thethree-dimensional analysis of the tracks rela-tively independent of the sampling rate andyielded a speed better related to displacementof the DC over time, a parameter that shouldbe more comparable between studies (Sup-plementary Fig. 2 online). Other studies havemade use of plots of displacement versus thesquare root of time to derive roughly linearplots consistent with a random ‘walk’20,21,31.The slope of these plots can be used todetermine a motility coefficient31, which isindependent of the data acquisition rate. Weused displacement as a filter at the single-celllevel by defining a ‘sessile’ cell as a cell thatdid not leave a sphere 20 mm in diameterwithin 30 min.

Most steady-state DCs in T cell zones weresessile, with a median speed slower than1 mm/min, whereas their motility coefficientwas 1.2 mm2/min (Fig. 4g, SupplementaryFig. 2 online and data not shown). Perifolli-cular DCs were mostly sessile, with a medianspeed of 1.6 mm/min (Fig. 4g). DCs in the Bcell follicle were the fastest moving, with amedian speed of 4 mm/min and relatively fewsessile cells (Fig. 4g). There were sparseindividual DCs that were ‘crawling’ in thefollicle and may have been traversing it(Supplementary Video 10 online). In almostall cases, cells that were sessile by displace-

ment thresholds had a nonzero speed, mostly because of ‘probing’movements too large to be filtered out by the 2.5-mm threshold.

Steady-state lymphocyte-DC interaction

The DC clusters in the T cell–B cell interface areas seemed to activelycapture round cells (Supplementary Video 12 online), which oftenremained in the clusters for the duration of imaging (430 min). Todetermine the cellular composition of DC clusters, we labeled thicksections of fixed inguinal lymph nodes with fluorescent antibodiesand examined three-dimensional reconstructions of confocal images(Supplementary Video 13 online). Clusters contained EYFP DCs,B cells and T cells (Supplementary Fig. 3 online). Because of theirtight packing, it was impossible to count the DCs in a cluster with theEYFP signal alone. We therefore visualized nuclei with the nuclearstain TO-PRO-3 in combination with CD3 and CD19 immunostain-ing (Fig. 5) and generated reconstructions as described above; clusterstypically contained up to eight DCs plus three to five B and T cells perDC on the interior (n ¼ 54 clusters measured). Lymphocytes in theseclusters were atypically round and sessile, rather than being polarizedand mobile like B and T cells in the B cell follicles and T cell zones.Because DC clusters were not always present, and because theyincorporate both B and T cells, we speculate clusters might be involvedin later stages (such as B cell–T helper cell interactions) of ongoingimmune responses.

Mature DC interactions with endogenous networks

As steady-state DCs acted differently than mature migratory DCsobserved before19–21, we directly compared these two cell populations

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Figure 5 DC clusters include more than one DC. Series of confocal images optically sectioning an

inguinal lymph node, with EYFP in green, DNA in red (TO-PRO 3), and CD3 and CD19 in blue. Arrows

indicate nuclei from DCs in a single cluster. DCs (A–H) show no CD3 or CD19 staining. Numbers above

images indicate the depth of the optical plane in the tissue section.

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by intravital imaging of the inguinal lymphnode. We injected purified, lipopolysaccha-ride (LPS)–activated ECFP+ DCs32 intrader-mally into the flanks of CD11c-EYFP mice asdescribed20. We examined inguinal lymphnodes histologically and by intravital micro-scopy from 12 to 72 h after DC injection.Transferred DCs were sparse in the inguinalnodes for up to 18 h; by 24 h there was visibleswelling of the lymph node, and ECFP DCshad migrated into the lymph node andsettled most densely at the interface betweenthe B and T cell zones (Fig. 6a). Althoughthis location brought them near HEVs, ECFPDCs were not directly associated with HEVs,based on immunostaining with MECA-79antibody (Fig. 6a). DCs were present throughout the T cell area at24 h and later time points (Fig. 6a).

We tracked ECFP+ and EYFP+ cells in the T cell zones as describedabove (Supplementary Fig. 2 online). In agreement with the histologyresults, ECFP+ DCs were not associated with HEVs and by 48–72 hwere evenly dispersed throughout the T cell area (SupplementaryVideo 14 online). At 24 and 48 h, the transferred mature ECFP DCsmoved faster than the EYFP+ DCs (P o 0.001), whereas at 72 h therewas no significant difference (Fig. 6b,c and Supplementary Video 14online). The motility coefficient for the mature DC decreased withtime after transfer, as reported before20 (from 1.88 mm2/min to 1.36 mm2/min), and both the immigrant and transferredpopulations contained sessile cells (o2 mm/min) at all time points(Fig. 6c). Furthermore, the median speed of the endogenous EYFP+

DCs increased with time after injection from 1 mm/min to 2.1 mm/minbetween 24 and 72 h, whereas the motility coefficient decreased from1.2 mm2/min to 0.9 mm2/min. This suggests that the endogenous cellshad more ‘probing’ and local dynamics without a corresponding

increase in movement through the tissue. This increase in local‘probing’ may have been due to the LPS injected along with theactivated DCs20. At 96 h, the numbers of transferred DC in the lymphnode dropped off sharply, probably indicating cell death. We concludethat mature DCs migrating to the lymph node are initially moremotile than steady-state DCs, but they eventually join the sessile DCnetworks after distributing themselves throughout the node.

DISCUSSION

We found that in the steady state, only a minor fraction of the DCs inthe lymph nodes ‘crawled’. Most DCs formed dense networks of cellsthat touched each other with the tips of their processes, which showedrapid and extensive ‘probing’. The DC networks surrounded the B cellfollicles on all sides and extended into the T cell zones, but wereparticularly dense in the border zone between the T and B cell follicle,where T cell–dependent immune responses are initiated33,34. In thisarea, DCs often adhered closely to one another, creating tightly packedclusters that incorporated T and B lymphocytes, which we speculate

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Figure 6 Transferred ECFP DCs gradually

incorporate into the endogenous DC network.

Splenic DCs were purified by immunomagnetic

selection from the spleens of ECFP+ mice

injected subcutaneously with a tumor secreting

Flt3 ligand, and 4 � 106 to 6 � 106 cells

were injected intradermally along with LPS

into the flanks of CD11c-EYFP mice. Inguinallymph nodes were then examined by

immunofluorescence and intravital microscopy.

(a) Confocal micrographs of inguinal lymph

nodes immunostained for HEVs with MECA-79

antibody. Scale bars, 200 mm (low-resolution

panels) or 100 mm (high-magnification insets);

optical thickness is 10 mm in the ECFP channel

and 1.2 mm in all other channels. (b) Plots of

displacement versus the square root of time,

used to calculate the motility coefficient of

transferred ECFP+ cells and endogenous EYFP+

cells. Inguinal lymph nodes were imaged

by multiphoton microscopy and cells were

tracked as described above; the motility of

transferred DCs gradually decreases as the cells

incorporate into the endogenous DC network.

(c) ‘Scattergram’ of the average velocities of

individual transferred and endogenous DCs (time,

horizontal axis). Horizontal lines indicate medianspeeds. Data in b and c summarize at least four

imaging fields examined in each time point.

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may be involved in later stages of the immune response. Bothnetworks and clusters were dynamic, in that ‘crawling’ DCs couldbe seen joining groups of sessile cells. When we compared endogenousDCs with mature DC immigrants, we found that the LPS-activatedmature DCs moved faster than endogenous DCs at 24 and 48 h as theydispersed into the T cell zone. Nevertheless, many of the immigrantDCs were sessile at all time points, indicating that these immigrantshad joined the endogenous DC network. Our finding are consistentwith the observation that the speed of mature DC movement slowswith time after arrival in the lymph node20,21. We speculate thatmature DCs stop migrating as they join DC networks in the lymphnode and that this change in activity may be required for efficientantigen presentation in vivo.

Whereas DCs in the T cell zone are referred to anatomically as‘interdigitating reticulum cells’35, based on their reticular appearancein fixed tissue sections, and are described as ‘sessile’36 because they donot recirculate, the microscopic dynamics of steady-state lymph nodeDCs were unknown. Our experiments differ from other publishedreports on DC movement in vivo that showed no evidence of a sessilenetwork and instead reported random DC ‘crawling’ with averagespeeds of 2.7–6.6 mm/min in lymph nodes19–21,28,31. However, all ofthose studies involved labeling of a small number of mature activatedDCs, and therefore the steady-state DC network could not havebeen appreciated.

In the steady state, DC networks present T cells with an enormoussurface area rich in the self MHC class II–peptide complexes requiredto sustain T cell longevity37,38 and to maintain self-tolerance29. Selfand foreign antigens that arrive in lymph nodes from the lymph areprocessed simultaneously by many DCs in the network7–10,39, muchlike antigen delivery by antibody to DEC-205 (anti-DEC-205;refs. 40,41); this maximizes the surface area for presentation ofsuch antigens.

We found that mature DCs that arrived in lymph nodes fromtissues were more motile on average than steady-state DCs. The higheraverage speed of immigrant DCs may be promoted by stimulation ofcytoskeletal dynamics by Toll-like receptors or chemokine signaling30.We propose that faster-moving mature DC traverse the network beforejoining it, whereupon they become sessile. In the network, DCs arewell positioned to form stable interactions with antigen-specific T cellsor to relay antigen to other DCs in the network by releasing exosomesor apoptotic bodies9,42. When such cell fragments are ingested byseveral DCs in the network, the antigen-presenting surface area isexpanded, increasing the probability that a T cell migrating throughthe lymph node would meet a DC carrying its cognate antigen.

The backbone of the DC network is likely to be the reticular fibernetwork, a system of collagen bundles that is enveloped by reticularfibroblasts and undergoes considerable remodeling in response toinflammatory signals from tissues43,44. Immigrant DCs may migratewhen in contact with T cells and other DCs and join the network onlywhen they locate stromal cells or attachment points to collagenbundles that would provide greater anchorage.

Earlier studies suggested that DCs typically contact 500 T cells perhour (ref. 19); higher-resolution imaging showed that the ‘probing’motion of DC processes enables DCs to contact up to 5,000 T cells perhour (ref. 21). This ‘probing’ activity is dependent on the smallGTPases Rac1 and Rac2, and genetic experiments have confirmedthe importance of these pathways for presentation to T cells45. Ourdata suggest that the DCs maintain a fixed position in the network andscan the passing T cells by ‘probing’ with their dendrites. Thearrangement of DCs in a network throughout the T cell area forcesT cells to pass among many ‘probing’ DCs on their course through the

lymph node. This is different from models in which both T cells andDCs are engaged in extensive random searches, which are based onprior data with immigrant DCs21. Given the high motility coefficientof T cells, there is little disadvantage to fixing DCs in the network.However, it is important that mature DCs carrying antigen into thelymph node from the periphery disperse throughout the T cell zonesto increase the probability that a randomly migrating T cell willcontact a DC presenting its cognate antigen. We propose that thehigher motility of mature DCs functions to distribute the DCs and theantigen they carry throughout the T cell zones and thereby maximizethe likelihood of antigen-specific DC–T cell interactions.

METHODSTransgenic mice. EYFP-Venus23 was cloned into a CD11c promoter24. The

linearized construct was injected into C57BL/6-CBA F1 fertilized female

pronuclei, and progeny positive by PCR for the transgene were backcrossed

to C57BL/6 for at least six generations. C57BL/6 mice expressing ECFP on the

b-actin promoter were purchased from Jackson Laboratories46. Mice were used

for experiments at 4–6 weeks of age; all mice were housed in specific pathogen–

free conditions and were treated in accordance with protocols approved by the

Rockefeller University and New York University School of Medicine Institu-

tional Animal Care and Use Committees.

Immunofluorescence. Lymph nodes were fixed in PBS with 4% paraformal-

dehyde plus 10% sucrose and were cryoprotected in PBS plus 30% sucrose

before being embedded in optimum cutting temperature compound and being

frozen. Frozen tissue was sectioned (20 or 50 mm in thickness) on a microtome

and was fixed in acetone. All incubations were done in a humidified chamber.

Sections were blocked in 5% BSA and 10% serum in PBS in the presence of

anti-CD16/CD32 (BD Biosciences) and were sequentially blocked with excess

streptavidin and biotin (Vector Laboratories). For peroxidase amplification

with biotinyl tyramides (NEN), we quenched endogenous peroxidase activity

with 3% H2O2 in PBS before the initial blocking step, according to the NEN

protocol. The antibodies used were directly labeled anti-CD3e–Alexa 647, anti-

B220–Alexa 647 (CalTag), biotinylated anti-CD3e, anti-CD1b, anti-CD11c,

anti-CD19, anti-CD80, anti-CD86, anti-B220, anti-F4/80, anti-Gr-1, anti-I-

Ab, anti-PECAM, anti-rat IgM (BD Biosciences), MOMA-1 (BMA Biomedi-

cals), anti-CD11c, anti-CD45.2 (EBioscience), unconjugated MECA-79 (BD

Biosciences) and biotin and Alexa 647 conjugates of anti-DEC-205 produced

and conjugated by the Rockefeller University Monoclonal Antibody Core

Facility (New York, New York). Streptavidin-indocarbocyanine was from

Jackson ImmunoResearch; TO-PRO-3 was from Molecular Probes. Sections

were mounted in Fluoromount-G (Southern Biotech) and were stored at 4 1C.

Confocal microscopy. Confocal images were acquired on a Zeiss LSM 510

system with 458-, 488-, 514-, 543- and 633-nm excitation lines at the Rock-

efeller University Bio-Imaging Facility; EYFP and ECFP fluorescence were

visualized directly in all images. Tiled images were obtained with the motorized

stage using a 40� or 63� objective to capture the entire cut surface of the

lymph node, and images were exported into Adobe Photoshop for final

processing. Three-dimensional reconstructions were generated from Z-stacks

with Imaris (Bitplane).

Flow cytometry. Single-cell suspensions of lymph nodes were pretreated with

anti-CD16/CD32 and were stained with CD11c-allophycocyanin and biotiny-

lated antibodies specific for CD3, CD4, CD8a, CD11b, CD19, B220, CD86,

DEC-205 and MHC class II (I-Ab; Becton Dickinson Pharmingen). Living cells

were gated based on forward- and side-scatter on a FACSVantage SE (Becton-

Dickinson) and data were analyzed with FlowJo (Tree Star).

Cell transfer. To demarcate B cell follicles, we transferred 1 � 108 B cells

purified from EGFP+ C57BL/6 mice by magnetic depletion with anti-CD43

(Miltenyi Biotec). CD11c+ DCs were purified by immunomagnetic selection

from the spleens of ECFP+ mice injected with cells from a tumor expressing

Flt3 ligand and 4 � 106 to 6 � 106 cells injected intradermally along with

1 ng/ml LPS (Sigma) at four sites surrounding the inguinal lymph node. To

image the macrophages in the subcapsular sinus and blood vessels throughout

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the lymph node, we injected 1 mg of rhodamine-dextran (66 kilodalton;

Molecular Probes) in 100 ml saline subcutaneously in the base of the tail.

To examine high endothelial veins in vivo, we injected 10 mg of Alexa

594–labeled MECA-79 antibody intravenously.

Surgery. Mice were anaesthetized with 100 mg ketamine, 15 mg xylazine and

2.5 mg acepromazine per kg body weight and were kept anaesthetized with

hourly injections of half this dose. Mice were restrained on a stage warmer at

37 1C (BioTherm Micro S37; Biogenics) and the abdominal skin was incised

from the edge of the rib cage through the midline to the thigh. A skin flap was

separated from the abdominal muscle to expose the inguinal lymph node.

To stabilize, moisturize and maintain the lymph node at 37 1C, we placed the

skin flap on a thermoconductive base made of silicone elastomer (Sylgard 184;

Dow Corning) surrounding a core of thermoconductive putty (T-putty 502;

Thermagon). The connective and adipose tissue covering the lymph node was

removed by microsurgery that kept blood and lymph vessels intact, and a

saline-filled chamber consisting of a cover-slip glued to a nylon washer was

mounted on the skin flap over the lymph node. The chamber was laterally

secured by three ‘insect pins’ that pierced the skin and lodged in the silicone

base. The temperature was monitored with a probe (Bioptechs) placed near the

lymph node to ensure that it was at 37 7 1 1C. To compensate for anesthesia-

induced respiratory depression, we provided mice with 100% oxygen by mask

throughout the imaging session.

Intravital microscopy. The lymph node was imaged with a Bio-Rad Radiance

multiphoton microscope fitted with a Tsunami pulsed laser (Spectraphysics)

and controlled by Lasersharp2000 software (Bio-Rad) and a Nikon 40�objective (water immersed; numerical aperture, 0.8). For imaging of EYFP,

EGFP and ECFP, the excitation wavelength was set to 910 nm; for rhodamine,

870 nm was used. Band-pass filters optimized for the detection of EYFP (550/

30) and ECFP (510/30) and a 530 long-pass dichroic mirror were used to detect

fluorescent proteins; EGFP appeared in both channels. To create time-lapse

sequences, we typically scanned volumes of tissue 50 � 400 � 400 mm at 3-mm

Z-steps and 30-second intervals.

Data analysis. Cell movement was analyzed with Volocity software (Improvi-

sion). For cell speeds, the coordinates of each cell were calculated and tracked

over time; to eliminate motion artifacts caused by dendrite ‘probing’, pulse and

breathing, displacements smaller than 2.5 mm were filtered out of the cell tracks.

Despite efforts to filter high-frequency movement related to ‘probing’, DCs that

were distinctly sessile still seemed to have velocities up to 2 mm/min; this

velocity was therefore used as the threshold below which cells were considered

sessile. As cell speed showed an asymmetrical distribution, data are presented in

‘scattergrams’ and were compared with a nonparametric ‘bootstrapping’

procedure based on 10,000 resampling iterations47. P values less than 0.05

were considered significant.

Note: Supplementary information is available on the Nature Immunology website.

ACKNOWLEDGMENTSWe thank R. Steinman for discussions and W. Gan for guidance on two-photonmicroscopy. Supported by the National Institutes of Health (AI55037 to M.L.D.and AI051573 to M.C.N.), Irene Diamond Foundation (M.L.D.), RothschildFoundation (G.S.), Medical Scientist Training Program (GM07739 to R.L.L.),German Research Foundation (DU 548/1-1 to D.P.) and Howard HughesMedical Institute (M.C.N.).

COMPETING INTERESTS STATEMENTThe authors declare that they have no competing financial interests.

Received 14 September; accepted 22 October 2004

Published online at http://www.nature.com/natureimmunology/

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