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Originally published online as doi:10.1189/jlb.1103588 on April 9, 2004

Published online before print April 9, 2004
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(Journal of Leukocyte Biology. 2004;75:1102-1110.)
© 2004 by Society for Leukocyte Biology

Short-term Flt3L treatment effectively mobilizes functional macaque dendritic cells

Natalia Teleshova*, Jennifer Jones*,1, Jessica Kenney*, Jeanette Purcell{dagger}, Rudolf Bohm{dagger}, Agegnehu Gettie{ddagger} and Melissa Pope*,2

* Center for Biomedical Research, Population Council, New York, New York;
{dagger} Tulane National Primate Research Center, Tulane University, Covington, Louisiana; and
{ddagger} Aaron Diamond AIDS Research Center, New York, New York

2 Correspondence: Center for Biomedical Research, Population Council, 1230 York Avenue, New York, NY 10021. E-mail: mpope{at}popcbr.rockefeller.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In vivo administration of soluble Flt3L increases dendritic cell (DC) numbers to favor improved DC targeting of vaccine antigens, augmenting vaccine efficiency. In addition to confirming the effectiveness of human Flt3L in macaques, we strove to determine the optimal regimen to elevate numbers of functional DCs. Circulating DCs were identified within lineage–human leukocyte antigen-DR+ cells, which comprised CD11c–CD123+ plasmacytoid DCs (PDCs) and CD123– cells including CD11c+CD123– myeloid DCs as well as CD11c–CD123– cells. Traditionally, DCs have been monitored 1–2 days after 10- to 14-day treatments with Flt3L (100 µg/kg/day). We demonstrate that although standard treatment increased macaque DC percentages, as little as 5–7 days of treatment was sufficient, if not more effective at mobilizing DCs. Moreover, DC frequency continued to escalate over the ensuing days, peaking at ~4 days post 7 days of treatment and ultimately decreasing thereafter. As expected, there was a more pronounced increase in the percentages and actual numbers of CD123– cells (CD11c+ and CD11c– subsets) compared with PDCs. Flt3L-mobilized DCs exhibited slightly increased CD80/CD86 expression but typically still that of immature DCs and were resilient to freeze-thawing. Overnight culture activated the cells, up-regulating CD80/CD86 expression as well as interleukin-12 release, typically being boosted by CD40L. This was even more apparent for enriched DC cultures. These data verify that peak mobilization of large numbers of functional macaque DCs occurs a few days, not immediately, after short-term Flt3L dosing. This has important implications for improved DC-targeting vaccine strategies to prevent infection with human immunodeficiency virus and other pathogens.

Key Words: monkeys • PDCs • MDCs • expansion • CD80 • CD86


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Increasing evidence emphasizes the need to ensure that vaccine antigens must be aptly presented to the immune system for maximal vaccine efficacy and that directly targeting the antigens to dendritic cells (DCs) represents an extremely promising strategy in this regard [1 2 3 ]. DCs represent a minimal proportion of the leukocytes in the blood and tissues, underscoring the necessity to design methods to efficiently target these rare cells for maximal immune activation. Circulating DCs exist in what is termed an immature state and must be activated into mature DCs to ensure appropriate immune activation for effective protection against a pathogen [4 ]. DC activation occurs as a result of triggering via tumor necrosis factor (TNF)–TNF receptor (TNFR) and Toll-like receptor (TLR) family members [2 ]. Maturation of DCs encompasses the up-regulation of costimulatory and adhesion molecule expression and increased secretion of interleukin (IL)-12 and IL-15, together favoring activation of potent T and B cell responses.

Two major populations of DCs exist, the CD11c+CD123–/low myeloid-derived DCs (MDCs) and CD11c–CD123high plasmacytoid-derived DCs [PDCs; DCs typically lack markers of natural killer (NK) cell, T cell, B cell, and monocyte lineages (Lin–)]. MDCs and PDCs respond to stimuli (CD40L, bacteria, viruses; refs. [5 6 7 8 9 ]) by up-regulating the expression of CD80 and CD86 coincident with their elevated immunostimulatory function. PDCs respond to influenza and herpes simplex viruses [5 6 7 ] or immunostimulatory oligodeoxynucleotides (ISS-ODNs) [10 11 12 ] by secreting enormous amounts of type I interferons (IFNs), notably IFN-{alpha}, which can drive the differentiation and activation of MDCs [13 , 14 ] and PDCs [15 ] biasing toward strong T helper cell type 1 responses [15 16 17 ]. Recent evidence revealed that MDCs also produce type I IFNs in response to viral infection [18 ]. Immunodeficiency viruses, human immunodeficiency virus [19 , 20 ], and simian immunodeficiency virus (SIV; N. Teleshova et al., submitted) also induce MDC/PDC-enriched populations to secrete IFN-{alpha}. High levels of DC-produced type I IFNs likely play an important role in innate immunity and also provide a link to augment the adaptive immune system [14 , 21 22 23 24 ].

Human studies suggest that DC numbers and function decrease with HIV disease progression [19 , 25 , 26 ], underscoring the need to boost their functionality for future immune therapies. The rhesus macaque system is being used to study the targeting of circulating DCs in vaccine strategies against HIV infection [3 ]. Recent progress identified typical MDCs and PDCs residing in macaque blood and tissues of naïve [27 , 28 ], SIV {Delta}nef-infected and wild-type-challenged or SHIV162P-infected macaques (N. Teleshova et al., submitted). Typically representing less than 1% of total leukocyte suspensions [27 , 28 ], macaque DCs respond comparably with TLR and TNF–TNFR triggering, much like their human counterparts. Previous murine [29 30 31 ] and human studies [32 33 34 ] demonstrated that subcutaneous (s.c.) administration of Flt3L significantly increased DC numbers in blood, lymphoid, and nonlymphoid tissues. In particular, the numbers of circulating CD11c+CD123–/low MDCs and CD11c–CD123high PDCs [32 33 34 ] increased ten- to 50-fold. Coates et al. [28 ] recently verified that a 10-day treatment with 100 µg human Flt3L per kg per day readily mobilized functional macaque DCs, increasing the numbers of Lin–human leukocyte antigen (HLA)-DR+ DCs from 0.82% to 10.4% (~13-fold increase) of the total blood leukocytes [28 ].

Herein, we sort to characterize Flt3L-mobilized macaque DCs and delineate the optimal time of Flt3L treatment as well as the peak of DC mobilization under such conditions. We show that as little as 5–7 days of treatment of healthy adult macaques with Flt3L increases DC numbers as well if not better than the traditional 10- to 14-day treatment. Moreover, percentages among peripheral blood mononuclear cells (PBMCs) as well as the actual numbers of Lin–HLA-DR+ cells per ml blood continued to rise after the cessation of treatment, and the peak increases were detected ~4 days after such short-term treatment. Increased numbers of CD11c–CD123– antigen-presenting cells (APCs) were also documented. Flt3L administration did not overtly activate the MDCs or PDCs, but both subsets up-regulated CD80 and CD86 expression upon culture, and the presence of CD40L enhanced this. This correlated with augmented IL-12 release by these cells. Flt3L-mobilized APCs were resilient to freezing and even exhibited enhanced viability upon in vitro culture. These data provide the first evidence for an effective, short-term Flt3L strategy that readily increases the numbers of healthy, functional DCs in macaque blood and will likely advance DC-targeted vaccine strategies against HIV as well as other pathogens.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Reagents
Human Flt3L (Chinese hamster ovary) and human CD40L trimer were kindly provided by Amgen (Thousand Oaks, CA). The reagents were stored at –80°C until used at the indicated doses. Thawed aliquots of CD40L were kept at 4°C for up to 12 weeks.

Animals and treatment
Adult male and female Indian rhesus macaques (Macaca mulatta) were colony-bred and housed at the Tulane National Primate Research Center (TNPRC; Covington, LA). At the commencement of these studies, all animals tested negative by polymerase chain reaction for simian type D retroviruses, simian T cell leukemia virus, and SIV. As some of the animals came from a previous study in which they were exposed to SHIV162P via the intravaginal route but had not reportedly become infected, we double-checked the plasma viral RNA levels in these animals during the study. Animals AM40, M010, P311, N833, AP37, and T693 were confirmed negative by branched DNA analyses (<250 copies/ml; Bayer Diagnostics, Emeryville, CA). However, AP85 showed up as a possible low-level infected animal with 298 viral copies per ml plasma (~2.5 years after infection). Animals were anesthetized with ketamine–HCl (10 mg/kg) before all procedures. Animals received daily s.c. injections of 100 µg/kg Flt3L for 5–14 days. Heparinized blood samples were taken at the indicated times (10 ml/kg/month), and superficial inguinal lymph nodes (LNs) were surgically removed before or after Flt3L treatment. Protocols were reviewed and approved by the Institutional Animal Care and Use Committee of the TNPRC. Animal care procedures were in compliance with the regulations detailed under the animal welfare act and in the Guide for the Care and Use of Laboratory Animals.

Cell isolation and culture
PBMCs were isolated from heparinized blood of treated and untreated macaques using Ficoll-Hypaque density gradient centrifugation (Amersham Pharmacia Biotec AB, Uppsala, Sweden). LN cell (LNC) suspensions were prepared using collagenase D digestion and mechanical disruption as described previously [35 , 36 ]. Total single-cell suspensions were further enriched for DCs by depleting Lin+ cells expressing CD3, CD8, CD11b, CD14, and CD20. This was achieved using the Miltenyi AutoMACS System (Miltenyi Biotec, Auburn, CA), where cells were first labeled with fluorescein isothiocyanate (FITC)-conjugated antibodies (Ab) against CD3 (clone SP34); CD8 (clone SK1) and CD20 (clone L27), followed by incubation with anti-FITC, -CD11b (anti-mouse/human), -CD14 (anti-human), and -CD19 (anti-human) beads (Miltenyi Biotec); and collection of the flow-through Lin– fraction (N. Teleshova et al., submitted). Whole-cell suspensions or Lin– DC-enriched cell mixtures were then analyzed by flow cytometry before and after in vitro culture.

The medium used for these studies was RPMI 1640 (Cellgro; Fisher Scientific, Springfield, NJ) containing 2 mM L-glutamine (Gibco-BRL Life Technologies, Grand Island, NY), 10 mM HEPES (Gibco-BRL Life Technologies), 50 µM 2-mercaptoethanol (Sigma Chemical Co., St. Louis, MO), penicillin (100 U/ml)–streptomycin (100 µg/ml; Gibco-BRL Life Technologies), and 1% heparinized human plasma. Cells were cultured at 1–2 x 105 cells per well of a 96-well, round-bottomed plate (MicrotestTM U-Bottom, Becton Dickinson, Franklin Lakes, NJ) in 200 µl medium or 5 x 105 cells per well of a 96-well, flat-bottomed plate (MicrotestTM 96, BD) in 250 µl medium. CD40L was added at a final concentration of 1 µg/ml. After 24 h of culture, cell-free supernatants were collected and stored at –20°C before enzyme-linked immunosorbent assay (ELISA) analysis. Cultured cells were also assayed by flow cytometry to assess phenotypic activation.

Fluorescence-activated cell sorter (FACS) analysis
Ab used in this study are listed in Table 1 . MDCs and PDCs were identified and characterized using three- or four-color flow cytometry. FITC–anti-Lin Ab (CD3, CD8, CD11b, CD14, CD20) were used with APC–anti-HLA-DR and PE–anti-CD123. Anti-CD56 is excluded here and replaced by anti-CD11b and -CD8 to stain NK cells, as macaque monocyte-derived DCs (moDCs) are CD16lowCD56+, unlike human moDCs [37 ], NK cells can lack CD56 (Jeffrey Lifson, personal communication), and we wanted to avoid removing CD16+CD56+ MDCs. A minor subset of CD8+ cells expressing moderate HLA-DR levels was identified within the Lin–HLA-DR+ cells (6.73%±2.27; data from three animals), which could reflect that CD8+ DCs have been excluded from these analyses. FITC–anti-Lin Ab, APC–anti-HLA-DR, Cy–anti-CD80 or -CD86, and PE–anti-CD123 combinations were used to monitor CD80 and CD86 expression by the various subsets. To better define MDC and PDC subsets, FITC–anti-Lin Ab, PerCP–Cy5.5–anti-HLA-DR, PE–anti-CD123, and APC–anti-CD11c Ab were used (Table 1) . DCs were defined within the Lin–HLA-DR+ gate (first gated on total leukocytes to exclude neutrophils), and then a gate identifying CD123+CD11c–, CD123–CD11c+, or CD123–CD11c– cells was applied to define DC percentages (see Fig. 2A ). Isotype-matched Ig controls were included for all experiments, and the mean fluorescence intensities (MFIs) of control Igs were routinely <1 log.


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  		Table 1. Monoclonal Ab Used in This Study

 


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  		Figure 2. Peak increases in DC mobilization occur 4 days after short-term Flt3L treatment. An example of gating techniques used to characterize the Lin–HLA-DR+ subsets is shown (A). PBMCs were gated according forward-scatter (FSC) and side-scatter (SSC) characteristics, and the neutrophils were largely excluded (R1, rectangular gate). A second gate (R2, oval gate) was set to identify Lin–HLA-DR+ cells within R1. CD123+ and CD123– subsets were identified within the combined R1 and R2 gates and monitored for CD80 or CD86 expression. (CD80 vs. CD123 staining is shown as an example.) Alternatively, combined CD11c and CD123 staining was performed to define the distinctive DC subsets within the Lin–HLA-DR+ cells (R1 and R2). Flt3L (100 µg/kg/d s.c.) was administered for 5 days (four animals) or 7 days (seven animals), and blood was sampled extensively during and for up to 24 days after the completion of treatment (set as the 0 time-point in each case). Lin–HLA-DR+ cells were identified in the PBMCs (A; and Fig. 1 ), and the percentages of Lin–HLA-DR+ cells at the indicated time points (x-axes) are plotted (B). DC percentages were determined before Flt3L (Pre) for comparison. Individual animal numbers are indicated in each panel. The averaged percentages of Lin–HLA-DR+ cells (burgundy bars) for the seven animals treated for 7 days are summarized (C; mean±SEM). This shows the baseline percentages pretreatment (Pre Flt3L) versus those at the peak 4-day time-point post the 7-day treatment (Post Flt3L). Lin–HLA-DR+ APCs were defined as CD123+ (123+, red bars) or CD123– (123–, green bars) cells (using PE–anti-CD123) to delineate the PDC- and MDC-containing fraction within total DC population, respectively. In addition, cells from two animals before and four animals after Flt3L treatment were analyzed by four-color flow cytometry to characterize the CD123– fraction (FITC–anti-Lin, PerCP–Cy5.5–anti-HLA-DR, PE–anti-CD123, and APC–anti-CD11c). The averaged percentages of Lin–HLA-DR+CD123–CD11c+ (123–11c+, blue bars) and Lin–HLA-DR+CD123–CD11c– (123–11c–, orange bars) are presented. The actual numbers of Lin–HLA-DR+, Lin–HLA-DR+CD123–, Lin–HLA-DR+CD123+, Lin–HLA-DR+CD123–CD11c+, and Lin–HLA-DR+CD123–CD11c– were calculated before and after Flt3L treatment (shown as 106 cells/ml blood; C). Numbers of Lin–HLA-DR+, CD123–, and CD123+ cells were calculated for five animals pre- and post-7-day Flt3L treatment. Two animals before Flt3L treatment and four animals after Flt3L treatment were used to determine the numbers of CD123–CD11c+ and CD123–CD11c– cells.

 
ELISA
Cell-free supernatants collected from the variously cultured cells were analyzed for the presence of IL-12 p70 as well as free p40 subunit (BioSource International Inc., Camarillo, CA).

Statistical analysis
The nonparametric-matched Wilcoxon test, the simple-sign test, and the parametric-paired difference t-test were performed to evaluate CD80 and CD86 expression on Lin–HLA-DR+ cells before and after Flt3L treatment.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Optimum DC yields after Flt3L treatment
Initial studies were performed to verify that human soluble Flt3L would function in rhesus macaques to mobilize DCs as seen in humans [32 33 34 ], mice [29 30 31 ], and most recently, in monkeys [28 ]. Two healthy macaques received the standard s.c. dose of 100 µg/kg/day Flt3L for 14 days. Blood and LN samples were taken before and 1 day after the completion of treatment, and additional blood samples were taken during treatment. Circulating DCs were identified within Lin–HLA-DR+ cells. In all samples, APC subsets were defined as CD123+ PDCs and CD123– cells (comprising MDCs and other APCs) [27 , 28 ]. Typically, FITC–anti-Lin Ab were used with APC–anti-HLA-DR and PE–anti-CD123 Ab. In some experiments, FITC–anti-Lin Ab were combined with PerCP-Cy5.5–anti-HLA-DR, PE–anti-CD123, and APC–anti-CD11c Ab to identify CD123+CD11c– PDCs, CD123–CD11c+ MDCs, and CD123–CD11c– cells.

As expected, Flt3L treatment improved the percentages of circulating Lin–HLA-DR+ cells in macaque blood and LNs (Fig. 1 and Table 2 ). Elevated percentages of Lin–HLA-DR+ blood cells were evident within 10 days of treatment (day –4 relative to completion of treatment) but were more pronounced immediately after continued treatment for 14 days. Blood Lin–HLA-DR+ cell percentages increased 17- to 30-fold (Table 2) . Lin–HLA-DR+ cells are present at lower frequencies in LN compared with blood, and Flt3L had less impact on LN Lin–HLA-DR+ cells, increasing them by <threefold. As a result of the lower Lin–HLA-DR+ cell frequency in LNs, subsequent studies focused on blood APCs, where larger numbers of cells are available for more detailed analyses.



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  		Figure 1. Traditional 2-week Flt3L treatment mobilizes macaque Lin–HLA-DR+ cells. A healthy, uninfected macaque received daily 100 µg/kg doses of Flt3L administered s.c. for 14 days. The last day of treatment has been set as Day 0 for analyses. Blood samples were taken on commencement of treatment (Day –13), during treatment (Days –10 and –4), and 1 day after the cessation of treatment (Day 1). A LN biopsy was also taken before (Day –13) and after (Day 1) treatment. Circulating APCs were detected in PBMC and LNC suspensions by staining with FITC–anti-Lin Ab and APC–anti-HLA-DR. Lin–HLA-DR+ APCs are indicated by the oval gates in each panel and highlight the increasing percentages of Lin–HLA-DR+ cells as a result of Flt3L administration. The percentages of Lin–HLA-DR+ cells detected at each time point are summarized in Table 2 for this and a second Flt3L-treated animal. Isotype Ig controls were included in each experiment and showed MFIs of <1 log, with no detectable changes post-Flt3L treatment.

 

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  		Table 2. Flt3L Mobilization of Lin–HLA-DR+ Cells in Blood and LN

 
Identifying the optimal treatment time for Flt3L mobilization
Knowing that the traditional 14-day Flt3L strategy mobilizes macaque DCs, we investigated whether shorter periods of treatment would equally mobilize DCs. This would be important to save precious reagents as well as to lessen the treatments that the animals endured. In addition, we were interested in determining how long the DC numbers remained elevated after the completion of Flt3L treatment. In total, we treated four animals for 5 days and seven animals for 7 days with daily 100 µg/kg/day Flt3L s.c. administrations. All animals were monitored as frequently as possible during and after the treatment to assess more accurately the kinetics of DC mobilization.

The gating strategies used to define the subsets are shown in Figure 2A . The percentages of Lin–HLA-DR+ cells identified by flow cytometry at each time-point are displayed in Figure 2B and 2C . After 5 days of treatment, there was little increase in the Lin–HLA-DR+ cell frequency immediately after cessation of treatment, as evidenced by the low percentages shown 2 days after the end of treatment. However, Lin–HLA-DR+ cell percentages increased with time up to 13–25% (average of ~12-fold increase) 4–6 days after the completion of the 5-day treatment (Fig. 2B , left). Similar and even more striking results were obtained after only 7 days of Flt3L treatment, where greater increases were typically observed over time (Fig. 2B , right). The peak of Lin–HLA-DR+ cell percentages appeared to occur ~4 days after stopping the 7-day treatment. The average percentage of Lin–HLA-DR+ cells at the day-4 peak-point after 7 days of Flt3L was 36.34%, ranging from sixfold up to 53.8-fold (18.4-fold average) more circulating Lin–HLA-DR+ cells compared with the levels before Flt3L (Fig. 2B , right, and Fig. 2C , left). Lin–HLA-DR+ cell percentages tended to begin to decrease within 5–6 days and had returned to baseline levels within 2–3 weeks post-treatment.

More detailed analysis of the cells by flow cytometry confirmed that the CD123+ PDCs represented only a small fraction of the Lin–HLA-DR+ cells, with ~93% of the Lin–HLA-DR+ cells being CD123– (Fig. 2A and 2C) . These ratios were similar after Flt3L treatment; however, there was an overall increase in the percentages of CD123– cells (~99%). Examining the percentages within the mobilized Lin–HLA-DR+ fraction, there was about a 1.5-fold increase in CD123+ PDCs compared with the ~13-fold increase in the CD123– MDC-containing subset. Within the CD123– subset, 56.8% ± 1.9 (two animals) were CD11c+ MDCs before Flt3L treatment (comparable with the 60.4%±3.9 CD11c+ MDCs detected in another eight naïve animals from a separate study; N. Teleshova et al., submitted). It is interesting that the frequency of CD11c+ MDCs appeared to decrease after the 7-day Flt3L treatment (33.76%±8.76, six animals; Fig. 2C , left). However, when the actual numbers were calculated per ml blood based on PBMC yield and DC percentages, the total numbers of each DC subset increased after Flt3L treatment. The total numbers (cells/ml) of Lin–HLA-DR+, Lin–HLA-DR+CD123–, and Lin–HLA-DR+CD123+ subsets increased by 51-, 57-, and fivefold, respectively. Of note, there was a 39-fold increase in the numbers of Lin–HLA-DR+CD123–CD11c+ MDCs and a 124-fold increase in the numbers of Lin–HLA-DR+CD123–CD11c– cells (Fig. 2C , right). Thus, short-term Flt3L treatment predominantly mobilized CD123– cells.

Mobilized DCs are immature, resilient to freezing, and can be enriched by lineage depletion
To characterize the Flt3L-mobilized DCs more carefully, the Lin–HLA-DR+ cells were stained with anti-CD80 or -CD86 in combination with anti-CD123 to delineate the levels of costimulatory molecules on the respective subsets (e.g., Fig. 2A ). Circulating CD123– MDC-containing fraction and CD123+ PDCs express low levels of CD80 and CD86 (Fig. 3 ). Even after Flt3L mobilization, these cells maintained low-level expression of these molecules (Fig. 3) . However, upon analysis of the MFIs of multiple samples, it was clear that the APCs in the blood of Flt3L-treated animals exhibited slightly elevated CD80/CD86 levels. These levels were still representative of immature DCs (MFIs <100) but may reflect partial activation as a result of Flt3L treatment. The differences in CD80 and CD86 expression before and after Flt3L treatment did not reach statistical significance.



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  		Figure 3. Low-level CD80 and CD86 expression by Flt3L-mobilized DCs. PBMC samples were taken before (Pre) and 4 days after (Post) 7-day treatment with Flt3L and analyzed by four-color flow cytometry (Fig. 2A) . Cells were stained with FITC–anti-Lin Ab, APC–anti-HLA-DR, PE–anti-CD123, and Cy–anti-CD80 or -CD86. CD80 and CD86 expression was monitored on Lin–HLA-DR+ cells as well as Lin–HLA-DR+CD123–and Lin–HLA-DR+CD123+-gated cells. The average MFIs (±SEM) of CD80 and CD86 expression by Lin–HLA-DR+ cells versus the CD123– and CD123+ cells from seven animals are shown pre- and post-Flt3L treatment.

 
We also tested the sensitivity of these cells to freezing and thawing to evaluate whether it was feasible to immediately store cells for later analysis. PBMCs collected 4 days after 7 days of Flt3L treatment (three separate animals) were processed immediately for FACS analysis, and the remainder of the cells was frozen (up to 20x106 cells/ml in 1.8 ml cryotubes). After 6–12 months in liquid nitrogen, the cells were thawed and analyzed by FACS. A representative FACS profile is shown in Figure 4A , where data on freshly isolated PBMCs versus frozen PBMCs from the same animal are provided. The significant Lin–HLA-DR+ subset is highlighted in each population. Examination of the percentages of these subsets in each sample revealed that freezing had little impact overall on Lin–HLA-DR+ cell percentages (Fig. 4B) . One animal exhibited a slight decrease, but the other two basically showed no changes in Lin–HLA-DR+ cell frequency after freezing. This observation was also repeated on two separate occasions with replicate samples of cells from AK66 collected 1 day post-Flt3L treatment (not shown).



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  		Figure 4. Recovery of Flt3L-mobilized Lin–HLA-DR+ cells from frozen PBMCs. PMBCs from three Flt3L-treated animals (AG64, N833, and AK66; 4 days post a 7-day dose), were analyzed immediately, or aliquots were frozen and stored in liquid nitrogen. Cells were thawed and evaluated for DC recovery by three-color flow cytometry. Representative FACS profiles from fresh (upper) versus frozen (lower) cells from animal N833 are provided (A). The circular gates denote the Lin–HLA-DR+ cells in each preparation. The average percentages of Lin–HLA-DR+ cells within the PBMCs of all three animals (frozen cells from AG64 were analyzed twice) are displayed (B).

 
Although there is a significant increase in Lin–HLA-DR+ cells following Flt3L treatment, we wanted to determine if we could enrich the DCs to more closely monitor the functional capacities of these cells (below). To achieve this, PBMCs were depleted of Lin+ cells using the Miltenyi autoMACS System (Miltenyi Biotech, Auburn, CA), leaving the Lin– DC-enriched fraction behind. PBMCs were routinely depleted of CD3+CD8+CD11b+CD14+CD20+ cells using combinations of directly labeled beads or FITC-labeled Ab followed by anti-FITC beads, as indicated in Materials and Methods. A representative FACS profile is shown (Fig. 5A ) detailing the Lin–HLA-DR+ APCs in PBMCs before depletion of Lin+ cells and the significantly enriched Lin–HLA-DR+ APC population in the Lin– fraction. Starting from such high percentages of Lin–HLA-DR+ cells (~36%), Lin–HLA-DR+ APC enrichment was very successful, typically yielding >80% Lin–HLA-DR+ APCs with no additional manipulation (Fig. 5B) . Analysis of the Lin–HLA-DR+ APCs confirmed that the percentages of CD123– MDC-containing fraction (~98.6% CD123– cells) and CD123+ PDCs (~1.4%) were maintained throughout this procedure (compare Figs. 2C and 5B ), and there appeared to be no significant enrichment or loss of either subset.



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  		Figure 5. Effective DC enrichment by removal of Lin+ cells. Taken at the peak of DC mobilization, PBMCs were depleted of Lin+ cells using the autoMACS system by removing CD3+CD8+CD11b+CD14+CD20+ cells. FACS analysis of FITC–anti-Lin Ab and APC–anti-HLA-DR-stained PBMCs before and after depletion of Lin+ cells (Lin– cells) confirmed that the Lin–HLA-DR+ DCs (circular gates) could be significantly enriched in this way (A). The mean percentages (±SEM) of Lin–HLA-DR+ cells (APCs) in total cells, Lin–HLA-DR+CD123+ (CD123+), and Lin–HLA-DR+CD123– cells (CD123–) within the Lin–HLA-DR+-gated Lin– cells from four separate animals (AG64, AP85, and M010 4 days after a 7-day treatment and T693, 6 days after a 5-day treatment) are provided (B).

 
DCs from Flt3L-treated macaques exhibit normal responsiveness to DC stimuli
Macaque DCs respond to classical TNF–TNFR and TLR stimuli [28 , 37 38 39 ]. Using this knowledge, we cultured PBMCs or DC-enriched Lin– cells in the presence and absence of human CD40L trimer and monitored activity at the level of membrane phenotype and IL-12 release. CD80 and CD86 levels expressed by Lin–HLA-DR+ APCs in PBMCs or in Lin– cells dramatically increased after overnight culture in medium, and overall, the inclusion of CD40L had little additional influence on this (Fig. 6 ). Note that the CD40L-induced up-regulation of CD80/CD86 was significantly more uniform in the Lin– cells compared with the medium-cultured cells or PBMCs (Fig. 6A) . Although the average MFIs were slightly lower, even Lin–HLA-DR+ APCs within previously frozen PBMCs readily up-regulated these molecules during culture (Fig. 6B) . Similar elevation of CD80/CD86 was observed on cultured Lin– cells (Fig. 6A and 6B) . It is interesting that further analysis of frozen samples revealed that Lin–HLA-DR+ APC percentages actually increased slightly after culture relative to the total leukocyte population (Fig. 6C) .



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  		Figure 6. CD80 and CD86 up-regulation by Flt3L-mobilized Lin–HLA-DR+ cells upon in vitro culture. Blood was collected 1 day or 4 days post-7 days of Flt3L treatment, and cells were analyzed immediately, or PBMC aliquots were frozen. Freshly isolated PBMCs, previously frozen PBMCs, or Lin– cells were cultured (or not; None) in medium (Med) or CD40L (1 µg/ml) for 24 h. Cultured and uncultured cells were assayed by three-color staining with FITC–anti-Lin Ab, APC–anti-HLA-DR, and Cy–anti-CD80 or -CD86. (A) Representative histograms showing CD80 and CD86 expression by Lin–HLA-DR+ cells in PBMCs and Lin– cell preparations from cells taken 4 days post-treatment. Blue lines show the isotype controls, and the red lines show the CD80/CD86 expression. (B) The MFIs of CD80 and CD86 expression by gated Lin–HLA-DR+ cells (4 days post-treatment) are shown for each preparation (three to four animals, mean±SEM). (C) The Lin–HLA-DR+ APC percentages from frozen PBMCs before and after culture in medium or CD40L are shown. Cells were assayed from three separate donors taken 4 days post-Flt3L treatment (AK66, AG64, and N833; AG64 was assayed on two separate occasions) and frozen samples from AK66 (1 day post-Flt3L) were assayed on two different occasions. The second assaying of the same animals is indicated by #2 in the key. Mean Lin–HLA-DR+ APC percentages of these six samples are indicated by the horizontal bars in each condition.

 
An additional measure of DC/APC activation was performed by monitoring the release of IL-12 by activated DCs. Previous studies in humans [10 , 40 ] and macaques [28 ] have shown that CD40L activation of MDCs and PDCs induces IL-12 production. Here, we tested whether Lin–HLA-DR+ APCs in blood collected 4 days post a 7-day Flt3L treatment would release IL-12 in response to stimulation. Some IL-12 is detected when PBMCs from untreated animals were cultured with CD40L, contrasting the significant levels released by PBMCs from Flt3L-treated animals in response to CD40L (Fig. 7 , left). CD40L-induced IL-12 levels ranged from 0 to 211 pg/ml secreted by cells from untreated animals up to 130–550 pg/ml from the cells of Flt3L-treated monkeys. As predicted, even higher levels of IL-12 were secreted by DC-enriched Lin– cells after overnight culture in medium (35–990 pg/ml), and this was augmented in the presence of CD40L (93–1276 pg/ml; Fig. 7 , right).



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  		Figure 7. Activated Flt3L-mobilized DCs secrete IL-12. PBMCs and Lin– cells prepared from blood drawn 4 days after a 7-day Flt3L treatment were cultured in the presence and absence of 1 µg/ml CD40L for 24 h. After culture, the cell-free supernatants were collected and stored at –20°C before being analyzed by ELISA. Mean (±SEM) IL-12 levels (pg/ml) produced are graphed for each cell preparation from four different donor animals.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
HIV vaccines are fittingly tested in the SIV–rhesus macaque model to evaluate the capacity of immune responses to prevent new or control established infections with immunodeficiency viruses [41 ]. Previous studies in the macaque using moDCs or skin-derived DCs confirmed the similarities between human and macaque DC systems in basic biology as well as in how they interact with immunodeficiency viruses [37 , 38 , 42 43 44 45 46 47 ]. Circulating blood DCs have been identified in macaques [27 , 28 ], and these cells respond to classical DC stimuli akin to human DCs [28 ], further supporting the use of the macaque system to explore DC-targeted HIV vaccines as well as to ascertain the role of distinct DC subsets in HIV transmission and disease progression. In agreement with an earlier report [28 ], we show herein that ~7% of the Lin–HLA-DR+ cells were CD123+ PDCs, and ~93% were CD123– cells, which comprised ~57% CD11c+ MDCs as expected [28 , 32 , 33 , 48 ].

DCs are a vital component of the immune system, and antigen must be processed and presented by properly activated DCs to guarantee that strong, active immunity against the foreign antigen is induced [2 , 4 ]. Under this premise, progressively more strategies are being explored to encourage direct targeting of DCs in vivo with vaccine antigens to improve vaccine efficacy. As a result of the rarity of DCs circulating in the periphery, it is necessary to specifically home in on DCs via approaches targeting DC-restricted surface molecules [49 ] or increase the numbers of circulating DCs to render them more easily accessed by the incoming vaccine antigens. The latter is being investigated in a number of ways using recombinant factors that mobilize cells from the bone marrow to elevate the DC numbers circulating in the periphery [29 , 33 , 50 , 51 ]. Earlier, more extensive studies, first in mice [29 30 31 ] and then in humans [32 33 34 ], demonstrated how s.c. Flt3L administration mobilized DCs readily, increasing their numbers up to 50-fold above the basal levels. Flt3L increased MDC and PDC subsets [32 33 34 ], and MDC numbers are augmented more dramatically than PDCs. Here, and in the recent report from Coates et al. [28 ], mobilization of functional DCs has been confirmed to occur comparably in the macaque in response to human Flt3L. What is more, we now show that a considerably shorter treatment time is sufficient to similarly mobilize DCs and that the peak in DC percentages occurs within a few days of completing the treatment, not immediately afterwards, which is when researchers typically monitored DC numbers after longer times of Flt3L dosing [33 , 51 ]. Analogous data can be achieved using a human Flt3L–IgG2 fusion protein to mobilize DCs in rhesus macaques and sooty mangabeys (Mark Feinberg, personal communication).

Moreover, this translated into a fivefold increase in the numbers of CD123+ PDCs and a 39-fold increase in CD11c+ MDC numbers (cells/ml) 4 days after the 7-day treatment. In addition, there was a 124-fold increase in the numbers of the CD11c–CD123– APCs. Whether these CD11c–CD123– cells represent double-negative DC subsets or precursors and/or CD20– B cells not identified in the lineage cocktail needs to be determined.

DCs mobilized by this short-term treatment are functional, responding to a classical DC stimulus, CD40L, by secreting IL-12 and increasing the expression of costimulatory CD80 and CD86 molecules. This is especially encouraging for the use of smaller amounts of the mobilizing agent (in this case, Flt3L) and reflects a regimen that is clearly more acceptable for an individual to tolerate. In fact, a single dose of Flt3L administered intratracheally to rats was recently shown to increase DCs and T cells in the local tissues, peaking 3 days later and resulted in enhanced Ab responses following immunization with tetanus toxoid [52 ]. The fact that less agent is needed to effect significant DC mobilization is also heartening for the potential application of DNA vaccines in which the Flt3L gene is incorporated, where less protein would be expected to be expressed but might be sufficient for DC mobilization. Early reports favor the possibility that DNAs encoding Flt3L can mobilize DCs and enhance responses against peptide [53 ] or DNA [54 ] vaccines.

Although it has been reported that additional stimulation of Flt3L-mobilized DCs was necessary to activate the immune system appropriately [55 , 56 ], it seems that under some conditions, Flt3L-mobilized DCs can enhance the immune responses against an antigen even in the absence of additional activation [53 , 54 ]. The route of Flt3L administration, the route of immunization, and/or the type of antigen may influence this. Our studies reveal that although Flt3L-mobilized DCs appear immature, they may have slightly elevated CD80/CD86 levels (compared with the levels detected before treatment), indicative of some activation. These costimulatory molecules are uniformly up-regulated at least tenfold, especially in enriched Lin– cell cultures exposed to CD40L (likely as a result of increased DC–DC communication), underscoring their responsiveness to stimuli. In separate studies, more extensively characterizing MDC and PDC function in uninfected versus SIV {Delta}nef-infected and wild-type-challenged or in SHIV162P-infected macaques that have not received Flt3L, similar DC activation occurs in response to culture with CD40L (N. Teleshova et al., submitted). However, in the absence of Flt3L treatment, the elevation of CD80/CD86 expression is less dramatic in response to medium culture and requires triggering via CD40L or ISS-ODNs to be impacted significantly. Of note, the peak MFIs after CD40L or ISS-ODN stimulation reach ~300 as opposed to the MFIs of ~1000 seen in Flt3L-mobilized DCs after culture in medium or CD40L. This further suggests that Flt3L treatment has heightened the sensitivity of DCs to activation, explaining how additional exogenous activation may be less critical for optimal presentation upon antigen exposure. Moreover, the Flt3L-mobilized DCs possess heightened viability as demonstrated by the increases in DC numbers relative to other leukocytes after overnight culture. This contrasts what we have observed in cells from non-Flt3L-treated animals (N. Teleshova et al., submitted) and suggests Flt3L boosts DC viability.

Therefore, there is increasing evidence that human Flt3L can augment DC and possibly other APC numbers in nonhuman primates. Furthermore, we have identified that short-term treatment is sufficient to mobilize DCs and that the peak of the increase in DC frequency occurs within a few days of completing treatment. Such strategies will likely advance the targeting of DCs with putative HIV vaccines in vivo as well as allowing more detailed analysis of the role of DCs in HIV transmission.


   ACKNOWLEDGEMENTS
 
This work received significant support from the Elizabeth Glaser Pediatric AIDS Foundation as well as from NIH Grants R21 AI52060 and R01 AI40877 (to M. P.) and the TNPRC Base Grant RR00164. M. P. is an Elizabeth Glaser Scientist. N. T., J. J., and J. K. contributed equally to this work. The authors thank Irving Sivin for his assistance with statistical analysis.


   FOOTNOTES
 
1 Current address: Department of Medicine, University of Alabama at Birmingham, 1530 3rd Avenue South, FOT 1203, Birmingham, AL 35294-3412. Back

Received November 25, 2003; revised January 26, 2004; accepted February 26, 2004.


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 ABSTRACT
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 DISCUSSION
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