In vitro hematopoiesis produces a distinct class of immature dendritic cells from spleen progenitors with limited T cell stimulation capacity

Ben Quah, Keping Ni and Helen C. O’Neill

School of Biochemistry and Molecular Biology, Australian National University, Canberra, ACT 0200, Australia

Correspondence to: H. C. O’Neill. E-mail: helen.oneill{at}anu.edu.au
Transmitting editor: M. Miyasaka


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
The study of dendritic cells (DC) has been hampered by the difficulty of isolating rare cells for analysis of their phenotype and function. Interpretation of the DC lineage has been largely influenced by studies on cell populations which can be readily isolated and amplified in the presence of cytokines. Long term cultures (LTC) from murine spleen have been shown to support continuous in vitro hematopoiesis of DC dependent on interaction with a stromal cell monolayer. LTC-DC represent a single, stable class of DC derived by constant turnover of spleen DC progenitors maintained within stroma. They represent a resident DC population in spleen. The functional characteristics of LTC-DC have been studied in terms of capacity to stimulate T cells and response to activation by environmental stimuli. LTC-DC have many morphological, phenotypic and functional properties reflecting an immature or partially mature, marginal zone-like CD4CD8 splenic DC subset. They are highly endocytic and can process and present protein antigen to naive hen egg lysozyme (HEL)-specific MHC-II-restricted TCR-Tg CD4+ T cells. They do not, however, induce T cell proliferation in a mixed lymphocyte reaction. LTC-DC do not respond in a typical fashion to common DC activators like LPS and CD40L. They upregulate MHC-I and CD80/CD86 but not MHC-II and CD40. They reflect an endogenous, immature DC subset in spleen with properties distinct from immature DC located in peripheral tissues.

Keywords: antigen presentation, dendritic cells, hematopoiesis, T cells


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Many subsets of dendritic cells (DC) have been identified which differ in phenotype and function. Over time these subsets have been classified variably in terms of phenotype, tissue-specific origin and state of activation or maturation (1). The generally accepted model is that DC located in peripheral tissues are immature and that after exposure to antigen in the presence of inflammatory factors these cells mature and migrate to lymphoid organs where they present antigen for T cell stimulation. This model has been well supported by studies on DC which capture antigen in sites such as skin, lung and gut (24). Migration of antigen-carrying DC from peripheral sites of infection into draining lymph nodes is associated with maturation of DC and is a key event in the priming of the immune response (5). DC maturation occurs upon exposure to microbes or inflammatory factors. Mature DC show upregulation of cell surface markers like CD80/CD86, MHC-II and CD40 and capacity to activate naive T cells. In contrast, immature DC are thought to play an important role in tolerance induction (6,7). Evidence that immature DC subsets are also located within lymphoid tissues such as spleen and lymph nodes (8) supports an important role for DC in the maintenance of self tolerance. Immature DC are characterized by high endocytic capacity and poor capacity to process and present antigen to T cells. DC in the steady-state are expected to be immature and play an important role in sampling the environment and maintaining tolerance.

Attempts to classify the DC subsets in different tissues are continuing with a view to solving the puzzle of DC development. However, the isolation of fresh murine DC is tedious, involving large amounts of tissue and labour intensive isolation procedures (9). This can result in significant leukocyte impurities and unintentional removal of some DC subsets (10). As a result, the classification of DC into different subsets and states of maturation is a very difficult task. Many in vitro protocols have been developed to propagate DC from progenitors using cocktails of cytokines. In mice, this involves isolation of progenitors from organs like bone marrow, blood and liver, and culture of cells with combinations of factors like granulocyte macrophage colony stimulating factor (GM-CSF), tumor necrosis factor-{alpha} (TNF-{alpha}) and IL-4 (11). Adoption of these in vitro cultures has led to significant advances in DC biology (12). However, addition of growth factors like TNF-{alpha} as well as the process of culturing cells in vitro can induce activation of DC with loss of their capacity to endocytose antigen for T cell stimulation (13). In vitro activation of DC can complicate the functional and phenotypic identification of DC subsets within tissue sites.

More recently, dendritic-like cells have been generated from precursor cells without the need for added cytokines, provided they are cultured in the presence of stromal cell monolayers containing endothelial cells. In one in vitro model, endothelial cells supported the development of DC from blood-derived monocytes (14). Human umbilical vein endothelial monolayers supported the differentiation of DC from blood monocytes within 2 days of culture by a process involving trans-endothelial migration (14). Long term cultures (LTC) established from mouse spleen in the absence of exogenous cytokines are a proven method for generating large numbers of DC where hematopiesis is supported by stromal cell monolayers comprising endothelial and fibroblastic cells (1518). LTC support the continuous differentiation of spleen progenitors into non-adherent cells displaying characteristics common to DC (1518). The production of DC in LTC has been confirmed by preparation of subtracted libraries representing precursor and progeny cells produced in LTC. Sequencing of differentially expressed clones indicated that cell differentiation within LTC was marked by upregulation of genes common to DC (19). Multiple LTC have been shown to reproducibly generate DC independently of GM-CSF (20). LTC represent a niche environment for reproducible hematopoiesis of spleen DC from progenitors contained within a stromal cell monolayer (17,18). The ease with which cells are produced in LTC would suggest that DC generated in LTC reflect the normal in situ production of a common class of DC in spleen.

Lymphoid organs are known to contain a major population of immature DC that play an important homeostatic role in tolerance. Mature DC in these organs are thought to enter from peripheral sites carrying antigen for T cell activation (8,21). The majority of spleen DC in both mouse and human have been shown to represent immature DC (8). Spleen is known to contain three subpopulations of DC identifiable on the basis of the expression of the CD4 and CD8 markers: CD4CD8 DC, CD4CD8+ (CD8+) DC and CD4+CD8 (CD4+) DC (22). The CD8+ DC subset which specifically expresses CD205 is thought to represent a class of DC located in T cell areas (23), while the CD4CD8 reside in the marginal zone (24). This study is concerned with spleen-derived DC produced in LTC and their functional characterization in relation to known DC subsets in spleen. It demonstrates that LTC-DC are representative of a functionally distinct endogenously produced immature DC population resident in spleen which is highly endocytic, refractory to activation by common DC activators and has only weak capacity to activate T cells.


    Methods
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Cell isolation from mice
Inbred mice were bred in the John Curtin School of Medical Research (Australian National University, Canberra, Australia) under specific pathogen-free conditions. BALB/c, B10.A(2R) and C57BL/6J male and female mice aged between 5 and 15 weeks were used throughout this study. TCR-transgenic (TCR-Tg) mice expressing TCR {alpha} and ß chains from the 3A9 T cell hybridoma specific for hen egg lysozyme (HEL) peptide presented by IAk (25) and bred on to the CBA/H (H-2k) background were kindly provided by Chris Goodnow (Australian National University). Mice were housed and handled according to the guidelines of the ANU Animal Experimentation Ethics Committee. Spleens or lymph nodes were harvested from mice euthanazed by cervical dislocation. Single cell suspensions were prepared by forcing tissue through a fine wire mesh using a syringe plunger followed by repeated pipetting in culture medium. RBC depletion involved cell lysis in 5 ml lysing buffer [0.14 M NH4Cl, 0.017 M Tris-base (pH 7.5)] for 5 min at 20°C followed by three washes in ice-cold medium.

In vitro generation of DC in spleen LTC
Primary spleen LTC were established as described previously (1518) in supplemented DMEM (Gibco BRL, Grand Island, NY) containing 10% fetal calf serum (sDMEM) as detailed previously (18). After several weeks of culture with medium change as needed, LTC were selected which contained a stromal cell monolayer of fibroblasts and endothelial cells that continually supported the proliferation and differentiation of progenitors into non-adherent dendritic-like cells. LTC sublines were generated by dissociating stroma in LTC using plastic scrapers and transferring aliquots of these together with non-adherent cells into new tissue culture flasks. Cultures were easily maintained with medium change every 2–3 days. LTC generated as many as 1 x 106 non-adherent cells per 75 cm2 of stroma every 2 days. Non-adherent cells were collected at medium change for analysis. Cells produced in LTC have been extensively characterized as DC (1520,26). Results presented here are representative of many replicate analyses on DC produced by multiple LTC derived from two different strains of mice.

For treatment, LTC-DC were cultured for 24 h without stroma in medium alone, or in the presence of various agents. Treatments included 10 µg/ml LPS (Sigma, MO, USA) and/or 1/100 dilution of CD40L titrated to produce maximal B cell proliferation. CD40L was prepared as the supernatent of paraformaldehyde-fixed CD40L-baculovirus-infected Sf9 cells, kindly provided by Virginia McPhun (Australian National University). For antigen pulsing, LTC-DC (106) were cultured without stroma for 24 h in the presence of 10 µg/ml HEL (Sigma) with or without 10 µg/ml LPS. Cells were washed twice in PBS (pH 7.4) to remove soluble protein antigen before addition to cultures of purified T cells.

Electron microscopy
Resin-embedded sections of non-adherent cells from LTC were prepared for transmission electron microscopy (TEM). Cells (107) were fixed overnight in 2% gluteraldehyde/0.1 M sodium cacodylate buffer (pH 7.4), washed in sodium cacodylate buffer (pH 7.4), stained in 1% osmium tetroxide for 1.5 h, repeatedly washed in ddH2O and stained in 2% uranyl acetate for 2 h. They were then washed in ddH2O and dehydrated in increasing concentrations of acetone. Cells were infiltrated with Spurr’s resin and cut into thin (80 nm) sections using a diatome diamond cutter followed by staining with lead citrate (2.6% lead nitrate, 3.5% sodium citrate, 0.16 M sodium hydroxide) for 15 min. After washing with ddH2O, sections were air-dried and viewed in a Hitachi 7000 transmission electron microscope (Hitachi, Japan) at 60 Kv.

Antibody staining and analysis of cells
Crystal violet (CV) has been found to quench the high autofluorescence of DC which can interfere with detection of bound fluorochrome-conjugated antibodies by flow cytometry (27). LTC-DC were treated with 2 mg/ml crystal violet for 7 min on ice followed by three washes in PBS. Prior to staining with specific antibody, cells were incubated for 15 min on ice in sDMEM/0.1% NaN3 containing 40 µg/ml of ‘Fc block’ or antibody 2.4G2 specific for Fc{gamma}II/IIIR (CD32/CD16) (Pharmingen, San Diego, CA). This prevents non-specific binding of both primary antibody and avidin-PE. This step was excluded for staining with antibody to Fc{gamma}II/IIIR, or if second stage reagents bound to 2.4G2.

For staining, saturating concentrations of antibody were added to cells in a total volume of 50 µl sDMEM/0.1% NaN3 for 30 min on ice followed by three washes with PBS/0.1% NaN3. If necessary, cells were incubated for 30 min on ice with a second stage fluorochrome-conjugated reagent, either FITC-conjugated sheep IgG anti-mouse Ig (Serotec, Oxford, UK), FITC-conjugated goat F(ab)'2 anti-rat Ig (Serotec) or avidin-PE (Pharmingen) in sDMEM/0.1% NaN3, followed by three washes in PBS/0.1 %NaN3. Cells were resuspended in 200 µl PBS/0.1% NaN3 for analysis. For multi-colour staining, the same incubation and washing procedure was repeated for the addition of other specific antibodies and fluorochrome-conjugated secondary reagents as required. Specificity of antibody was monitored through use of isotype matched control antibodies or second stage reagents alone. Antibodies with the following specificity were purchased from Pharmingen as affinity purified reagents: CD11c (HL3), Thy1.2 (30H-12), B220 (RA3-6B2), CD34 (RAM34), CD8{alpha} (53-6.7), CD4 (GK1.5), CD11b (M1/70), MHC-I (11-4.1), MHC-II (AF6 120.1 and TIB120), MHCII (TIB120), CD80 (16-10A1), CD86 (GL1), CD40 (3/23), TCR-Vß8.2 (F23.2) CD40L (MR1), TCR-Vß8.2 (F23.2) and Fc{gamma}II/IIIR (FcR; 2.4G2). Some antibodies with the following specificity were prepared as the supernatent of hybridoma cells received from the American Type Culture Collection (Rockville, MD, USA): DEC-205 (NLDC-145), DC (33D1), macrophages (F4/80).

Flow cytometry was performed in a FACSort (Becton Dickinson, CA, USA). Cell debris was gated out using a forward scatter (FSC) threshold of 50 and 10,000–50,000 events were collected for analysis. Data analysis was performed using CellQuest software (Becton Dickinson) and involved post-acquisition gating to obtain information on cell subsets. For analysis of lymphocytes amongst a mixed leukocyte population, cells were gated out on the basis of FSC and side scatter (SSC) properties. Lymphocyte blastogenesis was assessed on the basis of FSC above background and cell division by dilution of the intracellular dye, 5-(and 6-)carboxyfluorescein diacetate succinimidyl ester (CFSE; Molecular Probes, OR, USA). In some experiments, 5 µl of 100 µg/ml propidium iodide (PI) was added to 0.2 ml cells in PBS prior to flow cytometry for discrimination of dead cells. For total cell counts, 1000 CaliBRITE beads (Becton Dickinson) were added to 200 µl of cell solution for estimation of total cell number in solution based on ratio of cell subsets to beads in FSC and SSC analysis of collected data.

Endocytosis assay
LTC-DC were assessed for capacity to endocytose FITC-OVA (Molecular Probes). Cells were washed and placed on ice for 10 min before addition of 100 µg/ml OVA-FITC in a total volume of 100 µl sDMEM. Cells were then incubated for 45 min at 37°C. Control cells were kept on ice for 45 min. Endocytosis was halted by addition of 100 µl of ice-cold PBS/0.1% NaN3. Cells were washed three times and resuspended in PBS/0.1% NaN3 for analysis of fluorescence uptake by flow cytometry.

Preparation of purified CD4+ T cells from 3A9 TCR-Tg mice
A purified population of CFSE labelled CD4+ T cells was prepared from 3A9 TCR-Tg mice. A single cell suspension of spleen cells was prepared. Pelleted cells (<=107/ml) were labelled by resuspension in 1 ml sDMEM and addition of CFSE while vortexing to give a final concentration of 5 µM. Cells were then incubated at room temperature for 5 min before RBC depletion. CD4+ T cells were then isolated by depletion of unwanted cell subsets using magnetic Dynabeads coupled to sheep Ig anti-rat Ig (Dynal, Oslo, Norway). The procedure involved labelling unwanted cells with the following combination of rat antibodies: B220 (RA3-6B2), CD8{alpha} (53-6.7), CD11b (M1/70), MHC-II (TIB120), DEC-205 (NLDC-145), DC (33D1), macrophages (F4/80). Specific antibody was absorbed to cells for 30 min on ice. Cells were then washed three times in ice-cold sDMEM by centrifugation. Sheep Ig anti-rat Ig Dynabeads were added to cells for 30 min at 4°C with rotation using a bead to cell ratio of 4:1. Bead-coated cells were exposed to a magnet for 2 min, allowing aspiration of CD4+ T cells in solution. Purification typically resulted in >=90% purity of cells as assessed by antibody staining and flow cytometry.

Mixed lymphocyte reactions
To assess lymphocyte proliferation through DNA synthesis, RBC-depleted spleen cells were cultured in triplicate at 2 x 105 cells/well in 96-well plates together with diluting concentrations of LTC-DC as stimulator cells in a total volume of 200 µl sDMEM. Concanavalin A was used as a control stimulator of lymphocytes. Prior to culture, LTC-DC were irradiated (20 Gy on a Cobalt60 source; Commonwealth Scientific and Industrial Research Organisation [CSIRO], Black Mountain, ACT, Australia). DNA synthesis was assessed over the last 16 h of a 72 h culture through addition of 1 µCi/well of 3H-thymidine (3H-T) (Amersham, Buckinghamshire, UK). Cells were harvested on to glass fibre filters then saturated in MicroScint scintillation fluid (Packard, CT, USA) for measurement of label incorporation in a Top-Count scintillation counter (Packard). Responses were reported as mean c.p.m. ± SE of the mean of triplicate samples. Controls included responders or stimulators alone. Background 3H-T incorporation due to stimulators alone was <500 c.p.m.


    Results
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Morphology and ultrastructure of DC produced in LTC
Non-adherent cells produced in LTC were predominately spherical, internally complex and displayed dendrites visible under light microscopy (Fig. 1a and b). These cell populations and their supporting monolayer of stromal cells have maintained similar morphological characteristics over the course of this and previous studies (15,18). LTC represent a highly reproducible culture system for production of a population of DC of consistent morphology, phenotype and function.



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Fig. 1. DC produced in LTC are suspended above a supporting stromal monolayer. (a and b) Light microscopic examination of LTC from B10.A(2R) mouse spleen. (a) Non-adherent cells in focus above a stromal monolayer of fibroblast and endothelial cells. (b) Non-adherent cells display membrane processes and granular cytoplasm. Bars in each micrograph represent relative size in µm. (c and d) TEM analysis of lead citrate stained resin-embedded sections of non-adherent cells from LTC (80 nm thick) derived from B10.A(2R) mice. (c) Whole cell cross sections. (d) Intracellular ultrastructure of cells. Bars in each micrograph represent relative size in µm.

 
Non-adherent cells produced in LTC can be characterized as DC on the basis of their unique morphology and ultrastructure using TEM. Most cells were large with an irregular outline and an average diameter of 20 µm (Fig. 1c). Cell nuclei were predominantly irregular, eccentric and spherical or kidney-shaped, containing a peripheral rim of chromatin. The majority of cells displayed high cytoplasmic to nuclear ratio, with many mitochondria and abundant endosomes of varying density (Fig. 1d). Non-adherent cells produced in LTC have morphology and ultrastructure consistent with a metabolically active, endocytic, dendritic-bodied cell, analogous to previously described DC (28).

Phenotype of DC produced in LTC
In the absence of a definitive cell surface marker, DC are discriminated from other leukocytes through expression of multiple markers (29). Markers expressed by LTC non-adherent cells were analysed by labelling cells with specific antibodies and measurement of antibody binding by flow cytometry. Light scatter properties revealed that the majority of LTC non-adherent cells displayed high FSC consistent with their large size and high SSC consistent with cellular complexity. PI staining of cells showed that LTC maintain a population of ~25% dead cells reflecting the turnover of cells in a continuous culture system.

Antibody staining and flow cytometry were used to assess surface marker expression after gating to exclude dead cells and cell debris (Fig. 2). Antibody activity was titrated initially on positively staining control cell populations isolated from spleen (data not shown). Non-adherent cells produced in LTC expressed high levels of CD11c, a common marker on DC (9). Some cells expressed low levels of DEC-205, a marker associated with DC present in T cell areas of spleen (30). Very few LTC cells expressed the molecule recognized by the 33D1 antibody, thought to be expressed at low levels by marginal zone DC (24). Markers for lymphoid cells including Thy1.2, B220, CD8{alpha} and CD4 were not expressed. Cells did however express high levels of the myeloid marker CD11b, but did not express the macrophage/myeloid marker F4/80. Cells expressed moderate levels of MHC-I and CD80 but the expression of CD40 and CD40L was low or undetectable. Cells expressed high levels of CD86 typical of mature DC (29) but high levels of Fc{gamma}II/IIIR typical of immature DC. This staining pattern is typical of immature DC (9,31) except for the high expression of CD86. The majority of cells did not express MHC-II, but a subpopulation (6.9%) of LTC cells expressed high levels of MHC-II characteristic of mature DC (9). There was almost no expression of CD34, a marker of hemopoietic stem cells. If stem cells are produced in LTC, they are in very low numbers or contained within the stromal cell layer (26).



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Fig. 2. Cell surface marker expression and endocytic capacity of non-adherent LTC cells derived from B10.A(2R) mice. Cellular debris was gated out on the basis of FSC and SSC for all flow cytometry analysis of LTC cells. Cell surface expression of markers and endocytic capacity of LTC cells was analysed by flow cytometry. Marker expression was analysed by the binding of antibodies specific for CD11c (HL3), DEC-205 (NLDC-145), Thy1.2 (30H-12), DC (33D1), B220 (RA3-6B2), CD34 (RAM34), macrophages (F4/80), CD8{alpha} (53-6.7), CD4 (GK1.5), CD11b (M1/70), MHC-I (11-4.1), MHC-II (AF6 120.1), CD80 (16-10A1), CD86 (GL1), CD40 (3/23), CD40L (MR1) and Fc{gamma}II/IIIR (FcR; 2.4G2) relative to background binding of isotype controls or autofluorescence. Background binding is indicated by oval gates. LTC cells were stained with 2 mg/ml crystal violet for 7 min at 4°C to quench autofluorescence prior to antibody staining. Endocytic capacity was assessed by the uptake of 100 µg/ml FITC-OVA over 45 min at 37°C relative to background uptake at 4°C (oval gate). Percentages reflect proportion of cells within the adjacent gated region.

 
An important characteristic of immature DC is their capacity to take up antigen through endocytosis. This property is thought to enable DC to sample their surroundings for potential pathogens, but is rapidly down-regulated upon cell maturation (12). Non-adherent cells produced in LTC were incubated with OVA-FITC and monitored for fluorescent uptake by flow cytometry after incubation at 37°C versus 0°C. At 37°C, cells were able to efficiently take up soluble OVA-FITC. Greater than 95% of cells absorbed the protein after 45 min of culture compared with ~0.5% at 0°C, a temperature which inhibits endocytic ability (Fig. 2).

The majority of non-adherent cells present in LTC display many properties consistent with a predominately immature DC characterized by high endocytic capacity, high expression of Fc{gamma}II/IIIR and low expression of the molecules MHC-II, CD40 and CD40L. Most cells are apparently immature yet express high levels of the costimulatory molecule CD86 common to mature DC. A small subset of mature MHC-II+ cells was also detected.

Responsiveness of LTC-DC to activation
One question addressed was whether non-adherent cells produced in LTC respond to factors commonly known to activate DC. After exposure to LPS and CD40L, subsets of immature DC have been shown to undergo functional maturation, rapidly downregulating receptors for endocytosis and reducing their endocytic capacity (12). They also upregulate expression of immunostimulatory molecules such as MHC and CD80/86, important for antigen presentation and lymphocyte activation.

LTC-DC were collected and recultured on the plastic surface of tissue culture flasks in the absence of stroma. Cells were cultured for 24 h either in medium alone or in medium supplemented with activators. Cells were then collected and assessed for marker expression and endocytic capacity in comparison with cells freshly isolated from a stroma-dependent LTC. Activation conditions included reculture in the absence of stroma on plastic tissue culture flasks alone, or with added LPS or CD40L or both LPS and CD40L. Each of these conditions is a known potent activator of DC (13,31).

Non-adherent cells produced in LTC upregulated MHC-I and CD86 expression upon reculture in medium alone (Fig. 3a). This result suggested that the stromal environment of LTC did not lead to activation of developing DC. In fact, when cells were collected and cultured in the presence of known DC activators like LPS and CD40L, or a combination of these two agents, there was no further upregulation of marker expression upon reculture, indicating no further activation of cells. Only slight upregulation of CD40 and CD40L was detected and again these effects were not dramatically enhanced by addition of LPS or CD40L (Fig. 3a). Upregulation of CD40L could be a result of soluble CD40L adhering to LTC cells in culture even though cells were extensively washed prior to staining. Each of the treatments led to partial but very weak downregulation of endocytic capacity and of Fc{gamma}II/IIIR expression. This is also known to occur during the process of DC maturation (29).



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Fig. 3. LTC-DC display limited capacity to modulate marker expession and endocytic capacity upon stimulation. LTC-DC, namely 2RBQ (a) or X1 (b), were stimulated by reculture in the absence of stroma in fresh tissue culture flasks for 24 h in medium alone or in medium containing LPS (10 µg/ml) and/or CD40L (1/100 dilution, which gives maximal B cell proliferation). Marker expression and endocytic capacity was assessed in recultured cells relative to freshly isolated untreated LTC non-adherent cells by flow cytometry. Marker expression on LTC cells (solid lines) was analysed with monoclonal antibodies specific for MHC-I (11-4.1), MHC-II (AF6 120.1), CD80 (16-10A1), CD86 (GL1), CD40 (3/23), CD40L (MR1) and Fc{gamma}II/IIIR (FcR; 2.4G2) relative to background binding of isotype control antibodies or medium controls (grey shading). Endocytosis was assessed by the uptake of FITC-OVA over 45 min at 37°C (solid lines) relative to background uptake at 4°C (grey shading). For cells recultured in medium alone (a) dashed lines represent marker expression on fresh LTC cells. For LPS/CD40L treated cells (a and b), dashed lines represent marker expression on LTC cells recultured in medium alone. Numbers in (a) represent mean fluorescence intensity (MFI) values after treatment (solid lines). Numbers in (b) represent MFI before and after treatment.

 
The changes induced by LPS were found to vary slightly for non-adherent cells produced in different LTC. In the majority of cases, there was no upregulation in MHC-II expression. However, on three separate occasions, LPS was found to induce a slight upregulation of MHC-II above that induced by reculture of cells in medium alone (see Fig. 3b). This was also associated with noticeable upregulation of MHC-I and CD86 on non-adherent cells collected from these cultures.

LTC-DC can be induced to modulate marker expression and endocytic capacity, including upregulation of MHC and CD86 together with downregulation of Fc{gamma}II/IIIR and endocytic capacity. However, these effects were only minor, with treated cells still maintaining high endocytic capacity, high Fc{gamma}II/IIIR expression and weak expression of CD80, CD40 and CD40L and weak or no expression of MHC-II. Furthermore, treatment with TNF-{alpha} did not induce phenotypic changes in LTC cells (data not shown). In general, non-adherent cells derived from LTC are relatively refractory to common DC activators although they do show some maturation in response to treatment with activating agents. The in vitro production of DC in stroma-dependent LTC gives rise to immature DC most of which are not activated by the culture procedure.

Capacity to stimulate antigen-specific naive T cells
One important characteristic of immature DC is their capacity to take up and process antigen for presentation and subsequent activation of antigen-specific naive T cells. This effect can be dramatically enhanced by prior maturation of DC (9,33). The lymphostimulatory capacity of LTC-DC has not previously been shown to include MHC-restricted antigen-specific activation of naive T cells. With the relatively low level of MHC-II expression on the majority of LTC-DC, it was unclear whether cells could process and present protein antigen in order to stimulate naive CD4+ T cells. Since a minor subpopulation (6.9%) of LTC cells is shown to express high levels of MHC-II along with CD86 (Fig. 2), it is possible that LTC can develop into DC with immunostimulatory capacity within the LTC environment. The addition of LPS as a DC activator would be expected to upregulate immunostimulatory markers on cells and so increase the T cell response generated.

Non-adherent cells were collected from LTC and pulsed with HEL protein. Cells were then tested for their ability to stimulate purified CD4+ T cells from 3A9 TCR-Tg mice. Purified CD4+ T cells display the TCR derived from the 3A9 hybridoma specific for a HEL peptide presented in the context of IAk (25). T cells were purified from the spleen of TCR-Tg mice by antibody-mediated negative depletion to remove unwanted cell types. This protocol was chosen to avoid stimulation of T cells which occurs in positive selection methods. The purification method involved removal of MHC-II+ APC including DC, to preclude induction of T cell responses by endogenous APC. The depleted cell population showed enrichment for T cells having ~90% CD3{epsilon}+ cells and <=1% MHC-II+ cells (Fig. 4a). All gated CD4+ T cells expressed the TCR-Vß8.2 epitope of the transgene (Fig. 4a). The MHC-specificity of the response was tested by comparing responses generated by HEL-pulsed DC collected from LTC established from B10.A(2R) (IAk) mice and from C57BL/6J (IAb) mice. The inclusion of control IAb-positive stimulators also controlled for stimulation of T cells by any residual HEL taken up and presented by residual endogenous APC. Some cells were also treated with LPS at the time of antigen pulsing to determine the effect of LPS in activating LTC-DC to be more immunostimulatory.



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Fig. 4. LTC-DC stimulate antigen-specific naive CD4+ T cells from TCR-Tg mice. Non-adherent cells collected from LTC were assessed for capacity to stimulate purified CD4+ T cells from 3A9 TCR-Tg mice. (a) CD4+ T cells were enriched from lymph node and spleen by labelling cells with rat antibodies specific for MHC-II (IAbIEk: TIB 120), B220 (RA3-6B2), macrophages (F4/80), CD11b (M170), DC (33D1), DC (DEC-205) and CD8{alpha} (53–6.72) and magnetic removal of unwanted cells using sheep anti-rat Ig Dynabeads. The resulting population was highly enriched for CD3{epsilon}+ T cells and mostly depleted of MHC-II+ APC. This was confirmed by staining with the AF6 120.1 anti-MHC-II antibody which recognizes a different epitope to TIB 120 used in the T cell purification procedure. CD4+ T cells expressed the transgene product TCR-Vß8.2, as detected by binding of specific antibody (F23.2). (b) LTC-DC were pulsed with HEL protein (10 µg/ml) in the presence and absence of LPS (10 µg/ml) for 24 h. Cells were washed and 104 cells were assessed for capacity to stimulate 105 purified CD4+ TCR-Tg T cells. Untreated and LPS-treated B10.A(2R) (IAk) LTC cells were used as stimulators. Controls included untreated C57BL/6J (IAb) LTC cells and LPS-treated C57BL/6J (IAb) LTC cells or medium alone. Response to stimulation was measured using flow cytometry by upregulation of CD69, downregulation of CD3{epsilon}, blastogenesis (FSC) and proliferation (decrease in CFSE) at 0.5 and 3 days after culture. PI (25 µg/ml) was included to identify viable CFSE-labelled cells. Specificity of antibody labelling was monitored by background binding of isotype control antibody or medium alone (Nil). Numbers shown represent cell percentages in gated sectors. *Indicates MFI change in CD3{epsilon} expression by CD69+ cells relative to CD69 cells in the same plot.

 
In order to measure T cell responsiveness, TCR-Tg CD4+ T cells were examined by flow cytometry for upregulation of expression of the early activation marker CD69, downregulation of CD3{epsilon} (34), blastogenesis assessed as an increase in FSC and proliferation assessed by reduction in the mean fluorescence intensity (MFI) of the intracellular dye, CFSE. Measurement was performed at 0.5 and 3 days after co-culture with HEL-pulsed LTC cells or medium alone as a control. Naive HEL peptide-specific TCR-Tg T cells responded well to HEL-pulsed IAk-positive LTC cells. After 0.5 days, 27% of CD4+ T cells responded by upregulating CD69 expression above background (Fig. 4b). CD3{epsilon} expression was also reduced on CD69+ cells relative to CD69 cells with a reduction in MFI of 70 (Fig. 4b). At this early stage, 7.9% of cells showed signs of blastogenesis relative to controls. By 3 days, 40% of cells showed upregulation of CD69 with 51% of viable T cells (assessed by PI exclusion) showing blastogenesis. Of these T cell blasts, most had undergone at least one cell division and up to a maximum of four divisions as determined by reduction in CFSE intensity (Fig. 4b). T cell responses induced by IAb-positive LTC-DC were minor, showing only a 10% upregulation in CD69 expression at 3 days, blastogenesis in only 10% of cells and no cell proliferation indicated by CFSE staining. This confirmed an MHC-restricted response with no evidence for antigen presentation by endogenous APC present in the enriched T cell population.

LPS treatment of LTC-DC following antigen pulsing of LTC-DC had very little effect on T cell response (Fig. 4b). There was, however, early indication by 0.5 days of an increased T cell response following treatment of IAk-positive DC with LPS. This was indicated by a 17% increase in CD69 expression at 0.5 days which dropped to a 6% increase after 3 days. There was also evidence of greater reduction in the MFI of CD3{epsilon} on activated T cells. The effect of LPS treatment of LTC-DC on T cell responsiveness was noticeable at 0.5 days but was reduced by 3 days after stimulation. LTC-DC have capacity to take up and process proteins into peptides for presentation on MHC-II and to stimulate activation and proliferation of antigen-specific naïve CD4+ T cells. The activating effects of LPS on LTC-DC were noticeable but weak, which is consistent with the minor effects of LPS on the expression of CD86 and MHC-II seen in Fig. 3.

Inability of LTC-DC to stimulate an MLR
Consistent with their capacity to stimulate naive TCR-Tg CD4+ T cells, previous investigations have demonstrated that all tested LTC produce cells which can process and present conalbumin to the Th2 D10.G4.1 helper cell line (35). Despite this, LTC-DC derived from several cultures of B10.A(2R) mice were found to be incapable of stimulating an MLR in either allogeneic BALB/c mice or syngeneic C57BL/6J mice (Fig. 5). Responder lymphocytes from both strains could, however, respond to stimulation with concanavalin A.



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Fig. 5. LTC-DC fail to stimulate lymphocyte proliferation in an MLR. The lymphostimulatory capacity of LTC non-adherent cells was characterized in an allogeneic and a syngeneic MLR. RBC-depleted BALB/c or C57BL/6J responder spleen cells (1 x 105) were cultured with medium alone as a control, or with diluting numbers of 20 Gy irradiated B10.A(2R)-derived LTC-DC. Proliferation of responder spleen cells was measured by 3H-T incorporation over the last 16 h of a 72 h culture. The proliferation of responder spleen cells to diluting concentrations of concanavalin A (con A) is shown as a positive control. Data represent mean CPM ± SE (n = 3) after subtraction of background due to stimulators alone (<500 c.p.m.).

 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
This study has revealed that LTC generate non-adherent cells with unique DC properties. Cells display many of the structural, phenotypic and functional properties commonly used to define DC. Short cytoplasmic extensions, irregular nuclei with a peripheral rim of chromatin and high mitochondrial and endosomal content characterize LTC-DC and are known characteristics of DC (28), particularly those in an immature state (12). LTC-DC have a CD11c+ CD11b+ DEC-205–/low 33D1 F40/80 CD4 CD8{alpha} phenotype. They also have high expression of Fc{gamma}II/IIIR, low expression of MHC-II, CD80 and CD40 and high endocytic capacity, all typical of immature DC. Cells of similar phenotype have been identified in murine spleen (9, unpublished data). Their high expression of CD86 and a capacity to stimulate TCR-Tg T cells suggests attainment of some mature DC properties. Low to undetectable expression of DEC-205 is indicative of DC residing outside the T cell area of spleen (30), suggesting they may represent a marginal zone type DC in spleen (36). LTC cells do not however express molecules detected by the 33D1 antibody, reportedly expressed by spleen marginal zone DC (24). LTC-DC could represent an immature CD11c+CD11b+CD8{alpha}DEC-205–/low marginal zone type DC that has undergone partial maturation with DEC-205 upregulation and downregulation of CD4 and F4/80 (23). However, they also express high levels of CD11b and do not express CD8{alpha}. LTC could provide an environment that promotes DC development to a point where cells attain the properties of a T cell area type DC ready for lymphocyte modulation. This proposal is depicted in Fig. 6.



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Fig. 6. LTC-DC represent a partially mature DC. Model for the development of splenic DC incorporating a ‘transitional stage’ represented by LTC-DC which have many features common to marginal zone DC. They have dendritic form, express CD11c, MHC and costimulatory molecules, high capacity to endocytose antigen and to stimulate lymphocytes, including MHC-II-restricted naive CD4+ T cells. The expression marker profile of LTC-DC: CD4CD8{alpha}CD11b+DEC-205–/lowF4/8033D1: could be consistent with a transient ‘partially mature’ marginal zone type DC with surface expression of DEC-205 and capacity to modulate T cell responses and surface expression of DEC-205.

 
DC produced in LTC are not transformed cell lines. Cells are newly generated with high turnover from progenitors maintained within the stromal monolayer (26). LTC-DC represent immature/partially mature DC derived from self-renewing DC progenitors present in spleen. The phenotype of cells produced in LTC has remained remarkably stable over time and over many cultures. While plasticity of marker expression has been associated with DC following in vitro culture (9), it is also important to note that most culture procedures also induce DC differentiation and maturation which can be associated with changes in marker expression. In vitro propagation produces a population of cells of a single stable phenotype which could represent endogenously produced immature DC which have not yet acquired markers like CD4 and CD8 which classify distinct subsets in spleen. CD8{alpha} was previously thought to be a stable surface marker on a subpopulation of murine DC in the T cell area of spleen (9,36), but has recently been shown to vary in expression during DC differentiation (23). CD8{alpha} DC develop into CD8{alpha}+ DC in a process involving upregulation of DEC-205 concomitant with downregulation of CD11b, F4/80 and CD4 (23). This appears to be a normal developmental/maturational process of DC, with CD8{alpha} DC resident in the marginal zone where they trap antigen, subsequently developing into CD8{alpha}+ DC in T cell areas where they modulate lymphocyte responses. To date we have not identified signals which can induce differentiation of LTC-DC into the commonly identified spleen DC subsets which differ in expression of CD4 and CD8 (37). The stromal environment of LTC would appear to have limitations in that it supports DC production to an early stage in development.

LTC-DC have high expression of Fc{gamma}II/IIIR and a high capacity to endocytose protein antigen, typical of immature DC. In contrast, they have no capacity to stimulate lymphocyte proliferation in an MLR but can process protein antigen for presentation on MHC-II for stimulation of antigen-specific TCR-Tg CD4+ T cells, a property of mature DC. A partially mature phenotype for LTC-DC is consistent with high endocytic capacity and limited T cell activation capacity. Non-adherent LTC cells consistently express high levels of CD86 but low to very low levels of other immunostimulatory markers like MHC-I, MHC-II, CD80, CD40L and CD40. All of these markers are reportedly weakly expressed on immature DC but are upregulated upon DC maturation (12,31).

Despite their immature characteristics, LTC-DC are capable of stimulating antigen-specific responses from T cell clones and TCR-Tg T cells. Previously it was shown that DC in a range of maturation states including immature DC derived from bone marrow could process and present antigen to T cells (38). It is not clear, however, why LTC-DC cannot stimulate T cells in an allogeneic MLR, one of the first functional features identified in DC (39). Lack of T cell responses in an MLR might be a consequence of low expression of MHC-II on most LTC-DC, coupled with differences in activation thresholds of the responding T cell populations. It could also be argued that this is due to differences in the ligand avidity of the TCR on a clonal T cell population like D10.G4.1 or TCR-Tg T cells versus T cell populations that have a diverse TCR repertoire as in an MLR. However, recent studies have demonstrated that the response in T cells generated by DC in an allogeneic MLR may not be as clear cut as direct presentation of alloantigens by the stimulating DC (40). It was demonstrated that in order for allogeneic DC to stimulate a full response in MLR, the presence of syngeneic DC present in the responding leukocyte population was required (40). It was shown that allogeneic DC transferred MHC-II to syngeneic DC through CSN particles of <=200 nm in diameter, which endowed these syngeneic DC to stimulate T cells in a MLR (40). LTC-DC may not produce the factors required for this response to occur.

While a small subset of non-adherent LTC-DC expresses high MHC-II and clearly has capacity to initiate MHC-II-restricted T cell responses in 3A9 TCR-Tg mice, the majority of cells lack expression of MHC-II even after stimulation with LPS and CD40L. This contrasts with the majority of reports on DC (12). Lack of MHC-II expression could be a result of the in vitro culture conditions used to propagate LTC-DC. Freshly isolated DC from skin have been shown to dramatically decrease MHC-II synthesis during in vitro culture, with cessation after 3 days, although MHC-II is stably expressed on the cell surface (41). LTC-DC do synthesize MHC-II components (unpublished data). It is thought that with their high endocytic capacity, LTC-DC lose surface MHC-II in the rapid turnover of their plasma membrane. Recent evidence has shown that expression of MHC-II on the surface of DC is tightly controlled by the rate of endocytosis, with highly endocytic cells reabsorbing and degrading MHC-II/peptide complexes much faster than mature weakly endocytic DC (42).

LTC-DC have low propensity to respond to factors known to mature or activate commonly isolated subsets of DC. LPS treatment, CD40 crosslinking and TNF-{alpha} treatment have limited or no influence on either marker expression or endocytic function of LTC-DC. Furthermore, while reculture of LTC cells enhanced their expression of MHC-I and CD86, other markers like MHC-II and CD80 failed to upregulate. It has been demonstrated that DC treated with IL-10 (43) or TGF-ß (44) become less responsive to maturation signals. Analysis of gene expression has so far indicated that these two factors are not produced in LTC (unpublished data). Immature DC have also been propagated from BM with low doses of GM-CSF that are resistant to the maturational effects of LPS, CD40 ligation and TNF-{alpha} treatment (45). It has been suggested that these DC may induce T cell tolerance (41). LTC-DC clearly produce proliferative responses in TCR-Tg T cells. However, whether the end result of this T cell response is immunogenic or leads to abortive T cell proliferation involved in removing T cell clones (46,47) has not yet been addressed. This could occur if LTC-DC can stimulate regulatory T (Treg) cells. The outcome of T cell stimulation in terms of polarization of the T cell response to Th1, Th2 or Treg cells is currently under investigation.

Indeed, the finding that LTC-DC are refractory to further maturation with activation stimuli, like CD40L, LPS and TNF-{alpha}, is consistent with a model whereby LTC perpetually produce DC to a unique point in development. In support of this is evidence that DC have been shown to have a short ~2 day turnover time in spleen (48) and it is possible that the majority of splenic DC would die in an immature state with very few actually becoming mature cells. The inability of LTC-DC to respond to common DC activators would also be consistent with the presence of a distinct subset of immature DC in spleen which is refractory to activating factors known to activate DC located in other anatomical sites.


    Acknowledgements
 
This work was supported by grants to HO from the National Health and Medical Research Council of Australia, the Australian Research Council and the Clive and Vera Ramaciotti Foundation.


    Abbreviations
 
CFSE—5-(and 6-)carboxyfluorescein diacetate succinimidyl ester

CV—crystal violet

DC—dendritic cells

FSC—forward scatter

GM-CSF—granulocyte macrophage colony stimulating factor

HEL—hen egg lysozyme

LTC—long term cultures

MFI—mean fluorescence intensity

PI—propidium iodide

SSC—side scatter

sDMEM—supplemented DMEM

TCR—T cell receptor

TCR-Tg—TCR-transgenic

TEM—transmission electron microscopy

TNF—tumor necrosis factor


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

  1. Mellman, I. and Steinman, R. M. 2001. Dendritic cells: specialized and regulated antigen processing machines. Cell 106:255.[ISI][Medline]
  2. Randolph, G. J., Inaba, K., Robbiani, D. F., Steinman, R. M. and Muller, W. A. 1999. Differentiation of phagocytic monocytes into lymph node dendritic cells in vivo. Immunity 11:753.[ISI][Medline]
  3. Vermaelen, K. Y., Carro-Muino, I., Lambrecht, B. N. and Pauwels, R. A. 2001. Specific migratory dendritic cells rapidly transport antigen from the airways to the thoracic lymph nodes. J. Exp. Med. 193:51.[CrossRef][ISI][Medline]
  4. Huang, F. P., Platt, N., Wykes, M., Major, J. R., Powell, T. J., Jenkins, C. D. and MacPherson, G. G. 2002. A discrete subpopulation of dendritic cells transports apoptotic intestinal epithelial cells to T cell areas of mesenteric lymph nodes. J. Exp. Med. 191:435.[CrossRef]
  5. Banchereau, J. and Steinman, R. 1998. Dendritic cells and the control of immunity. Nature 392:245.[CrossRef][ISI][Medline]
  6. Lutz, M. B. and Schuler, G. 2002. Immature, semi-mature and fully mature dendritic cells: which signals induce tolerance or immunity? Trends Immunol. 23:445.[CrossRef][ISI][Medline]
  7. Steinman, R. M., Hawiger, D. and Nussenzweig, M. C. 2003. Tolerogenic dendritic cells. Annu. Rev. Immunol. 21:685.[CrossRef][ISI][Medline]
  8. Wilson, N. S., El-Sukkari, D., Belz, G. T., Smith, C. M., Steptoe, R. J., Heath, W. R., Shortman, K. and Villadangos, J. A. 2003. Most lymphoid organ dendritic cell types are phenotypically and functionally immature. Blood 102:2187.[Abstract/Free Full Text]
  9. Vremec, D. and Shortman, K. 1997. Dendritic cell subtypes in mouse lymphoid organs: cross-correlation of surface markers, changes with incubation, and differences among thymus, spleen, and lymph nodes. J. Immunol. 159:565.[Abstract]
  10. Martin, P., Del Hoyo, G. M., Anjuere, F., Arias, C. F., Vargas, H. H., Fernandez, L. A., Parrillas V. and Ardavin, C. 2002. Characterization of a new subpopulation of mouse CD8alpha+ B220+ dendritic cells endowed with type 1 interferon production capacity and tolerogenic potential. Blood 100:383.[Abstract/Free Full Text]
  11. Szabolcs, P., Moore, M. A. and Young, J. W. 1995. Expansion of immunostimulatory dendritic cells among the myeloid progeny of human CD34+ bone marrow precursors cultured with c-kit ligand, granulocyte-macrophage colony-stimulating factor, and TNF-alpha. J. Immunol. 154:5851.[Abstract/Free Full Text]
  12. Banchereau, J. and Steinman, R. M. 1998. Dendritic cells and the control of immunity. Nature 392:245.[CrossRef][ISI][Medline]
  13. Gallucci, S., Lolkema, M. and Matzinger, P. 1999. Natural adjuvants: endogenous activators of dendritic cells. Nat. Med. 5:1249.[CrossRef][ISI][Medline]
  14. Randolph, G. J., Beaulieu, S. Lebecque, S., Steinman R. M. and Muller, W. A. 1998. Differentiation of monocytes into dendritic cells in a model of transendothelial trafficking. Science 282:480.[Abstract/Free Full Text]
  15. Ni, K. and O’Neill, H. C. 1997. Long-term stromal cultures produce dendritic-like cells. Br. J. Haematol. 97:710.[ISI][Medline]
  16. Ni, K. and O’Neill, H. C. 1998. Hemopoiesis in long-term stroma-dependent cultures from lymphoid tissue: production of cells with myeloid/dendritic characteristics. In Vitro Cell Dev. Biol. Anim. 34:298.[ISI][Medline]
  17. Wilson, H. L., Ni, K. and O’Neill, H. C. 2000. Identification of progenitor cells in long-term spleen stromal cultures that produce immature dendritic cells. Proc. Natl Acad. Sci. USA 97:4784.[Abstract/Free Full Text]
  18. Wilson, H., Ni, K. and O’Neill, H. C. 2000. Proliferation of dendritic cells in long term culture is not dependent on granulocyte/macrophage colony stimulating factor. Expt. Haem. 28:193.
  19. Ni, K. and O’Neill, H. C. 2001. Development of dendritic cells from GM-CSF–/– mice in vitro: GM-CSF enhances production and survival of cells. Devel. Immunol. 8:133.
  20. Summers, K. L., Hock, B. D., McKenzie, J. L. and Hart, D. 2001. Phenotypic characterization of five dendritic cell subsets in human tonsils. Am. J. Pathol. 159:285.[Abstract/Free Full Text]
  21. Vremec, D., Pooley, J., Hochrein, H., Wu, L. and Shortman K. 2000. CD4 and CD8 expression by dendritic cell subtypes in mouse thymus and spleen. J. Immunol. 164:2978.[Abstract/Free Full Text]
  22. Martinez del Hoyo, G., Martin, P., Arias, C. F., Marin A. R. and Ardavin, C. 2002. CD8alpha+ dendritic cells originate from the CD8alpha-dendritic cell subset by a maturation process involving CD8alpha, DEC-205, and CD24 up-regulation. Blood 99:999.[Abstract/Free Full Text]
  23. Agger, R., Crowley, M. T. and Witmer-Pack, M. D. 1990. The surface of dendritic cells in the mouse as studied with monoclonal antibodies. Int. Rev. Immunol. 6:89.[Medline]
  24. Ho, W. Y., Cooke M. P., Goodnow, C. C. and Davis, M. 1994. Resting and anergic B cells are defective in CD28-dependent costimulation of naive CD4+ T cells. J. Exp. Med. 179:1539.[Abstract]
  25. Wilson, H. L. and O’Neill, H. C. 2003. Dynamics of dendritic cell development from precursors maintained in stroma-dependent long-term cultures. Immunol. Cell Biol. 81:144.[CrossRef][ISI][Medline]
  26. Ni, K. and O’Neill, H. C. 2000. Improved FACS analysis confirms generation of immature dendritic cells in long-term stromal-dependent spleen cultures. Immunol. Cell Biol. 78:196.[CrossRef][ISI][Medline]
  27. Steinman, R. M. and Cohn, Z. 1973. Identification of a novel cell type in peripheral lymphoid organs of mice. I. Morphology, quantitation, tissue distribution. J. Exp. Med. 137:1142.[ISI][Medline]
  28. Hart, D. N. 1997. Dendritic cells: unique leukocyte populations which control the primary immune response. Blood 90:3245.[Free Full Text]
  29. Inaba, K., Pack, M., Inaba, M., Sakuta, H., Isdell, F. and Steinman, R. M. 1997. High levels of a major histocompatibility complex II-self peptide complex on dendritic cells from the T cell areas of lymph nodes. J. Exp. Med. 186:665.[Abstract/Free Full Text]
  30. Pinchuk, L. M., Klaus, S. J., Magaletti, D. M., Pinchuk, G. V., Norsen, J. P. and Clark, E. A. 1996. Functional CD40 ligand expressed by human blood dendritic cells is up-regulated by CD40 ligation. J. Immunol. 157:4363.[Abstract]
  31. Kelleher, M. and Beverley, P. C. 2001. Lipopolysaccharide modulation of dendritic cells is insufficient to mature dendritic cells to generate CTLs from naive polyclonal CD8+ T cells in vitro, whereas CD40 ligation is essential. J. Immunol. 167:6247.[Abstract/Free Full Text]
  32. Turley, S. J., Inaba, K., Garrett, W. S., Ebersold, M., Unternaehrer, J., Steinman, R. M. and Mellman, I. 2000. Transport of peptide-MHC class II complexes in developing dendritic cells. Science 288:522.[Abstract/Free Full Text]
  33. Cai, Z., Kishimoto, H., Brunmark, A., Jackson, M. R., Peterson, P. A. and Sprent, J. 1997. Requirements for peptide-induced T cell receptor downregulation on naive CD8+ T cells. J. Exp. Med. 185:641.[Abstract/Free Full Text]
  34. O’Neill, H. C., Ni, K. and Wilson, H. 1999. Long-term stroma-dependent cultures are a consistent source of immunostimulatory dendritic cells. Immunol. Cell Biol. 77:434.[CrossRef][ISI][Medline]
  35. Leenen, P. J., Radosevic, K., Voerman, J. S., Salomon, B., van Rooijen, N., Klatzmann, D. and van Ewijk, W. 1998. Heterogeneity of mouse spleen dendritic cells: in vivo phagocytic activity, expression of macrophage markers, and subpopulation turnover. J. Immunol. 160:2166.[Abstract/Free Full Text]
  36. Shortman, K. and Liu, Y. J. 2002. Mouse and human dendritic cell subtypes. Nat. Rev. Immunol.2:151.[CrossRef][ISI][Medline]
  37. Veeraswamy, R. K., Cella, M., Colonna, M. and Unanue, E. R. 2003. Dendritic cells process and present antigens across a range of maturation states. J Immunol. 170:5367.[Abstract/Free Full Text]
  38. Steinman, R. M. and Witmer, M. D. 1978. Lymphoid dendritic cells are potent stimulators of the primary mixed leukocyte reaction in mice. Proc. Natl Acad. Sci. USA 75:5132.[Abstract]
  39. Bedford, P., Garner, K. and Knight S. C. 1999. MHC class II molecules transferred between allogeneic dendritic cells stimulate primary mixed leukocyte reactions. Int Immunol. 11:1739.[Abstract/Free Full Text]
  40. Kampgen, E., Koch, N., Koch, F., Stoger, P., Heufler, C., Schuler, G. and Romani, N. 1991. Class II major histocompatibility complex molecules of murine dendritic cells: synthesis, sialylation of invariant chain, and antigen processing capacity are down-regulated upon culture. Proc. Natl Acad. Sci. USA 88:3014.[Abstract]
  41. Villadangos, J. A., Cardoso, M., Steptoe, R. J., van Berkel, D., Pooley, J., Carbone, F. R. and Shortman K. 2001. MHC class II expression is regulated in dendritic cells independently of invariant chain degradation. Immunity 14:739.[CrossRef][ISI][Medline]
  42. Caux, C., Massacrier, C., Vandervliet, B., Barthelemy, C., Liu, Y. J. and Banchereau, J. 1994. Interleukin 10 inhibits T cell alloreaction induced by human dendritic cells. Int. Immunol. 6:1177.[Abstract]
  43. Bonham, C. A., Lu, L., Banas, R. A., Fontes, P., Rao, A. S., Starzl, T. E., Zeevi, A. and Thomson, A. W. 1996. TGF-beta 1 pretreatment impairs the allostimulatory function of human bone marrow-derived antigen-presenting cells for both naive and primed T cells. Transpl. Immunol. 4:186.[CrossRef][Medline]
  44. Lutz, M. B., Suri, R. M., Niimi, M., Ogilvie, A. L., Kukutsch, N. A., Rossner, S., Schuler, G. and Austyn, J. M. 2000. Immature dendritic cells generated with low doses of GM-CSF in the absence of IL-4 are maturation resistant and prolong allograft survival in vivo. Eur. J. Immunol. 30:1813.[CrossRef][ISI][Medline]
  45. Townsend, S. E. and Goodnow, C. C. 1998. Abortive proliferation of rare T cells induced by direct or indirect antigen presentation by rare B cells in vivo. J. Exp. Med. 187:1611.[Abstract/Free Full Text]
  46. Hawiger, D., Inaba, K., Dorsett, Y., Guo, M., Mahnke, K., Rivera, M., Ravetch, J. V., Steinman, R. M. and M. C. Nussenzweig. 2001. Dendritic cells induce peripheral T cell unresponsiveness under steady state conditions in vivo. J. Exp. Med. 194:769.[Abstract/Free Full Text]
  47. Kamath, A. T., Pooley, J., O’Keeffe, M. A., Vremec, D., Zhan, Y., Lew, A. M., D’Amico, A., Wu, L., Tough, D. F. and Shortman, K. 2000. The development, maturation, and turnover rate of mouse spleen dendritic cell populations. J. Immunol. 165:6762.[Abstract/Free Full Text]