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(Journal of Leukocyte Biology. 2002;72:285-296.)
© 2002 by Society for Leukocyte Biology

H1° histone and differentiation of dendritic cells. A molecular target for tumor-derived factors

Dmitry I. Gabrilovich*, Pingyan Cheng*, Yuhong Fan{dagger}, Bin Yu*, Ekaterina Nikitina*, Allen Sirotkin{dagger}, Michael Shurin{ddagger}, Tsunehiro Oyama§, Yasushi Adachi§, Sorena Nadaf§, David P. Carbone§ and Arthur I. Skoultchi{dagger}

H. Lee Moffitt Cancer Center, University of South Florida, Tampa;
{dagger} Department of Cell Biology and Cancer Center, Albert Einstein College of Medicine, Bronx, New York;
{ddagger} Department of Surgery, University of Pittsburgh, Pennsylvania; and
§ Department of Medicine and Cancer Center, Vanderbilt University, Nashville, Tennessee

Correspondence: Dmitry Gabrilovich, M.D., Ph.D., H. Lee Moffitt Cancer Center, University of South Florida, MRC-2E, Room 2067, 12902 Magnolia Dr., Tampa, FL 33612. E-mail: dgabril{at}moffitt.usf.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Dendritic cells (DC) play a central role in antitumor immune responses. Abnormal differentiation of DC and their inability to stimulate T cells are important factors in tumor escape from immune-system control. However, the mechanisms of this process remain elusive. Here, we have described one possible molecular mechanism that involves replacement linker histone H1°. A close association between expression of H1° and DC differentiation in vitro has been found. DC production in H1°-deficient mice was decreased significantly, whereas generation and function of macrophages, granulocytes, and lymphocytes appear to be normal. However, these mice had a significantly reduced response to vaccination with antigens. Tumor-derived factors considerably reduced h1° expression in hematopoietic progenitor cells. We have demonstrated that transcription factor NF-{kappa}B is involved actively in regulation of h1°. Thus, H1° histone may be an important factor in normal DC differentiation. Tumor-derived factors may inhibit DC differentiation by affecting H1° expression.

Key Words: monocytes/macrophages • cellular differentiation • tumor immunity


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Professional antigen-presenting cells, dendritic cells (DC), play a central role in induction of antitumor immunity. Defective function of these cells in cancer is considered one of the important factors responsible for tumor escape from immune system control. We, and others, have demonstrated previously that tumor-derived factors dramatically affect DC differentiation. This manifests in decreased production of mature DC and generation of immature myeloid cells incapable of effective antigen presentation [1 2 3 4 5 6 7 ]. Several tumor-derived factors including vascular endothelial growth factor (VEGF) have been implicated in abnormal DC differentiation. However, the molecular mechanisms of this process remain unknown. Using VEGF as a natural inhibitor of DC development, we sought to identify genes essential for differentiation of these cells and, in particular, the genes affected by tumor-derived factors. Identification of such genes is not only important for our understanding of the biology of these cells but also may suggest new strategies to restore DC function in cancer. Using a differential display assay (DDA), we have found that H1° histone is one of the genes inhibited specifically by VEGF in hematopoietic progenitor cells (HPC).

The H1° histone is a lysine-rich member of the H1 family of linker histones. These proteins bind to the linker DNA between nucleosome cores and facilitate the formation of higher-order chromatin structures [8 ]. Considerable evidence connects histone H1 to gene regulation [9 10 11 ]. Recent studies demonstrated that at any given time, most of H1 histone molecules are bound to chromatin, but histone H1 is rapidly exchanged between different chromatin regions in the cell nucleus [12 ]. To date, seven subtypes of H1 have been identified in mammals. Five of these proteins (H1a, H1b, H1c, H1d, and H1e) are collectively termed the somatic subtypes [13 ]. Replacement linker histone H1° differs substantially from the somatic subtypes [14 ]. Somatic H1 histones are linked tightly to DNA replication [15 ]. In contrast, histone H1° expression is rather low throughout the cell cycle. As a result, proliferating cells in culture contain somatic histones almost exclusively. After cells stop proliferation and begin to differentiate, H1° accumulates. A similar pattern is observed in tissues [16 ]. However, the role of this histone in cell differentiation remains unknown. H1°-deficient mice develop normally, suggesting that other H1 subtypes may have overlapping or redundant functions [17 ].

In this study, we have investigated a possible role of H1° in differentiation of myeloid and lymphoid cells. We have found that H1° is involved in differentiation of DC but not macrophages, granulocytes, or lymphocytes. Tumor-derived factors inhibit h1° expression in HPC that may contribute to defective DC differentiation in cancer.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals
Female BALB/c and C57BL/6 mice (6–8 weeks old) were purchased from Harlan Inc. (Indianapolis, IN) and were housed in specific pathogen-free units of the Division of Comparative Medicine at H. Lee Moffitt Cancer Center, University of South Florida (Tampa). h1° Knockout mice were generated by targeted disruption of the h1° gene, as described previously [17 ].

Antibodies and reagents
The following antibody-producing hybridomas were obtained from American Type Culture Collection (ATCC, Manassas, VA) and were used as culture supernatants: anti-CD4 (L3T4, TIB-207), anti-CD8 (Lyt-2.2, TIB-210), and anti-major histocompatibility complex (MHC) class II (I-Ad, TIB-120). Mouse granulocyte macrophage-colony stimulating factor (GM-CSF), G-CSF, M-CSF, interleukin (IL)-4, and tumor necrosis factor {alpha} (TNF-{alpha}) were obtained from RDI (Flanders, NJ); FLT3 ligand (FL), from R&D Systems (Minneapolis, MN); and polyclonal anti-mouse immunoglobulin (Ig), from Sigma Chemical Co. (St. Louis, MO). Purified or fluorescein isothiocyanate (FITC)- or phycoerythrin (PE)-conjugated anti-Gr-1, TER-119, anti-CD11c, CD11b, CD86 (B7-2), I-Ab, CD3, CD4, CD8, CD28, B220, and natural killer (NK)-1.1 antibodies were purchased from Pharmingen (San Diego, CA). Isotype-matched FITC- and PE-conjugated IgG were used as a control of nonspecific binding. FL was injected subcutaneously (s.c.) at dose of 10 µg per mouse daily for 10 days. FITC-conjugated anti-F4/80 and NK-1.1 antibodies were purchased from Serotec (Washington, DC).

Four different tumor cell lines have been used for generation of conditioned media. MethA cells, methylcholantrene-induced sarcoma developed in BALB/c mice and passaged as an ascitic tumor, was obtained from Dr. L. J. Old. C3 tumor cells (gift from Dr. W. M. Kast, Loyola University, Chicago, Maywood, IL) were constructed by transfecting B6 MEC cells of C57BL/6 mice with EJ-ras and plasmid containing human papillomavirus type 16 [18 , 19 ]. P815 mastocytoma was obtained from ATCC and L929 fibrosarcoma, from Dr. M. Rita Young, Loyola University, Chicago, Maywood, IL. NIH 3T3 fibroblasts obtained from ATCC were used as a control. Conditioned media were obtained from these cell lines by 48 h incubation of confluent cells in RPMI-1640 medium with a reduced concentration of serum (3%). This concentration of serum, which did not affect viability of tumor cells over a short (48 h) period of time, was found to be minimal.

Cell separation and analysis of cell surface receptors
A single cell suspension was prepared from inguinal, axillary, and brachial lymph nodes, spleens, and thymuses, and red cells were removed by hypotonic shock. For analysis of cell surface receptors, cells were washed in phosphate-buffered saline (PBS) supplemented with 0.1% fetal calf serum (FCS) and labeled with appropriate antibodies for 30 min at 4°C. Cells were then washed and analyzed on a FACSCalibur flow cytometer (Becton Dickinson, Mountain View, CA). Bone marrow was obtained from the femurs and tibias of mice and then enriched for HPC by depletion of lineage-specific cells by incubation with antibody cocktail (TIB-207, TIB-210, TIB-120, anti-mouse Ig, anti-B220, anti-Gr-1, and anti-TER119) and complement (Low-Tox® guinea pig complement, Cedarlane Laboratories, Ontario, Canada).

Generation of DC
DC were generated from HPC. Briefly, HPC were seeded into 24-well plates (0.5x106 cells per well) and cultured in RPMI 1640 supplemented with 10% FCS, 20 ng/ml murine GM-CSF, 10 ng/ml IL-4 (RDI), and 50 µm 2-mercaptoethanol. Half of the medium was replaced on day 3. After 5–6 days in culture, cells were collected, and the purity of the DC fraction was determined by fluorescein-activated cell sorter analysis of surface molecules (CD11c, CD11b, B7-2, IAd).

T and B cell isolation and proliferation
Single cell suspensions were obtained from spleens using a cell strainer with 70 µm pores. Red cells were removed by osmotic lysis with ACK buffer, and T cells were purified from spleens using T cell-enriched columns (R&D Systems). B cells were purified by positive selection using anti-B220 antibody and magnetic cell sorting. Briefly, cells were labeled with biotinylated anti-CD45R (B220) antibody on ice for 30 min, washed, and incubated with streptavidin microbeads, followed by separation on MiniMACS column (Miltenyi Biotec, Auburn, CA). B cells were cultured in triplicates in U-bottom 96-well plates (105 cells per well) with 5, 10, or 20 µg/ml lipopolysaccharides (LPS) for 48 h. [3H]-Thymidine (1 µCi; Amersham, Arlington Heights, IL) was added to each well 18 h prior to cell harvest. Incorporation of [3H]-thymidine was counted using a liquid scintillation counter and expressed as counts per minute (cpm).

T cells were cultured in triplicates in U-bottom 96-well plates (105 cells per well) with 1 and 5 µg/ml concanavalin A (Con A) for 72 h. In another experiment, U-bottom wells were coated with 0.2 µg/ml anti-CD3 antibody and 5 µg/ml anti-CD28 antibody. T cells (105 cells per well) were incubated for 72 h, and proliferation was measured as described above. To measure allogeneic mixed leukocyte reaction (MLR), lymph node and spleen cells obtained from control and h1° knockout mice were irradiated (20 Gy), plated in triplicates, and incubated with enriched T cells obtained from BALB/c mice for a 4-day allogeneic MLR. T cell proliferation was measured as described above.

Immunohistochemical analysis
Tissues from animals were snap-frozen in O.C.T. compound and stored at -80°C. Tissue sections (6µm) were air-dried and fixed in acetone, rehydrated in PBS, and blocked for nonspecific staining with 2% normal goat serum in PBS. Primary anti-mouse NLDC-145 (DEC 205; Serotec Ltd., Oxford, UK) antibodies were applied for 60 min at room temperature. Subsequently, all slides were washed in PBS and incubated 30 min with biotinylated secondary antibodies (Jackson Lab., West Grove, PA). Bound antibodies were detected with ABC reagent and Vectastain substrate kit (Vector Lab., Burlingame, CA).

Functional activity of granulocytes and macrophages
Mice were injected intraperitoneally (i.p.) with 1 ml 3.9% sodium thioglycolate (Sigma Chemical Co.). Granulocytes were collected 5–6 h and macrophages, 3–4 days later. Granulocytes were purified on Histopaque/Lymphoprep gradients with density 1.119 and 1.077.

Nitroblue tetrazolium (NBT) test
Macrophages were cultured for 24 h on coverslips in six-well plates in complete medium. Medium was changed, and cells were cultured for additional 24 h. In part of the wells, cells were activated by 24-h incubation with 10 ng/ml phorbol 12-myristate 13 acetate (PMA; Sigma Chemical Co.). After that time, cells were washed in PBS followed by 20 min incubation with 1.2 mg/ml NBT (Sigma Chemical Co.) in PBS containing 36% FCS. Cells were then washed in PBS, fixed in 100% methanol, counterstained for 10 min with 1% safranin O, and counted by microscopy (total magnification, x1000). The results of the NBT test were expressed in arbitrary units (AU) calculated per 100 cells using a formula: 1a + 2b + 3c (where a=cells with less than 25% of cytoplasm covered with formazan, b=cells with 25–75% of cytoplasm covered with formazan, and c=cells with more than 75% of cytoplasm covered with formazan). A NBT test in granulocytes was performed immediately after isolation of cells. For activation of granulocytes, 1-h incubation with 10 ng/ml PMA was used. The results were analyzed exactly as described for macrophages.

Assay of phagocytosis
Macrophages and granulocytes were prepared as described above. Macrophages were activated for 24 h with 5 ng/ml interferon-{gamma} (IFN-{gamma}; RDI) and granulocytes, for 1 h with 10 ng/ml LPS (Sigma Chemical Co.), washed, and incubated for 1 h at 37°C with an aqueous suspension of polystyrene latex beads (0.807 µm in diameter; Sigma Chemical Co.) at a ratio of 100 particles per cell. Cells were then washed in PBS, fixed in 100% methanol for 10 min, and stained with Giemsa. Phagocytosis was analyzed by microscopy (total magnification, x1000) and expressed as index = (axb)/100 (where a=percentage of cells engulfed latex particles, calculated per 100 cells, and b=number of latex particles engulfed by 100 cells). Granulocytes were incubated with latex in suspension, fixed in methanol, put on slides, dried, stained with Giemsa, and analyzed exactly as described for macrophages.

TNF-{alpha} production
Peritoneal macrophages were cultured for 24 h in 3 ml medium, alone or supplemented with 5 ng/ml IFN-{gamma} or 10 µg/ml LPS. Supernatants were collected, and TNF-{alpha} activity was measured in biotest as described before [20 ]. Briefly, 4 x 103/well L929 cells were seeded in flat-bottom 96-well plates. Twenty-four hours later, medium was replaced with the fresh one containing 1% FCS and 5 µg/ml actinomycin D (Sigma Chemical Co.). Different dilutions of cell supernatants were added in triplicates. Plates were incubated for 18 h at 37°C, stained for 15 min with 0.2% crystal violet (Sigma Chemical Co.), washed, and dried. Cells were then lysed with 100 µl 50% acetic acid, and optical density was measured on a spectrophotometer at a wavelength of 540 nm. Calibration of TNF-{alpha} activity was performed using recombinant murine TNF-{alpha} (RDI).

RNA preparation and analysis of gene expression
RNA was extracted from HPC using the GlassMAX® RNA microisolation spin cartridge system (Gibco-BRL, Life Technologies, Grand Island, NY). Reverse transcriptase-polymerase chain reaction (RT-PCR) was performed on RNA samples following a DNA digestion. Reverse transcription was performed using SUPERSCRIPT preamplification system for first-strand cDNA synthesis (Gibco-BRL). The following oligonucleotide PCR primer pairs for the h1° histone were used: forward primer, 5'-CCACGGACCACCCCAAGTATTCAG-3'; reverse primer, 5'-CTTGGCTTTGGGCTTCACGGGTTTT-3'.

HPRT-specific primers were used as a control and have been described elsewhere [21 ]. For analysis of h1°, histone samples were denatured for 30 s at 94°C, annealed for 30 s at 58°C, and extended for 45 s at 72°C for 28 cycles of amplification. This number of cycles was selected to avoid saturation of PCR products and was determined after preliminary experiments. For analysis of HPRT, samples were annealed at 52°C for 25 cycles of amplification. Products were visualized by staining with ethidium bromide after electrophoresis in 1% agarose gel. The PCR product sizes for h1° and hprt were 488 bp and 164 bp, respectively. PCR products were transferred overnight in an alkaline transfer buffer (0.4 N NaOH, 1 M NaCl) onto Hybond N+ nylon transfer membranes (Amersham), hybridized for 2 h in a rapid hybridization buffer (Amersham), and probed with [32P]-labeled oligonucleotide probes: H1°, 5'-CTCCAAAGCCCCAAGCAAGAAACC-3'; HPRT, 5'-GTTGTTGGATATGCCTTGAC-3'.

Gene expression was quantified using UN-Scan-IT software (Silk Scientific, Orem, UT). H1° expression in each sample was normalized for hprt and was expressed as AU (h1°/hprtx100).

For inhibition of nuclear factor (NF)-{kappa}B, we used an adenoviral construct encoding a dominant-negative NF-{kappa}B inhibitor, I{kappa}B{alpha}, with deleted serine phosphorylation sites (I{kappa}B{Delta}N). This construct has been described elsewhere [22 ].

Assessment of colony formation
Colony formation by HPC was measured using semi-solid 1% methylcellulose medium supplemented with recombinant cytokines (Epo, SCF, IL-6, IL-3) supporting the optimal growth of burst-forming units-erythroid (BFU-E), colony-forming unit (CFU)-GM, CFU-M, CFU-G, and CFU-GEMM colonies (Methocult M3434, Stemcell Technologies, Vancouver, Canada). Bone marrow cells were seeded at 15,000 cells per plate. BFU-E colonies were scored on days 8–9, and all other colonies were enumerated on days 12–13.

Stimulation of T cell responses
For evaluation of DC ability to stimulate primary T cell response, DC generated from h1°+/+ and h1°-/- mice were loaded overnight with 1 mg/ml ovalbumin (OVA; Sigma Chemical Co.) in complete culture medium. In control, cells were incubated only in complete medium. Cells were then washed, irradiated, and incubated at different ratios with T cells from h1+/+ or h1-/- mice. T cell proliferation was measured 4 days later as described above. To evaluate the response of H1°-deficient mice to immunization, two different experimental systems have been used. To measure CD4-mediated response, mice were immunized twice with a 10-day interval with 3 x 105 DC loaded with OVA. Mice were sacrificed after 4–5 days, and splenocytes (105 cells/well) were stimulated for 72 h in triplicates with OVA-derived MHC class II-matching peptide ISQAVHAAHAEINEAGR or irrelevant peptide KYMCNSSCM in U-bottom 96-well plates. Cell proliferation was determined using [3H]-thymidine uptake as described above. To measure CD8-mediated response, mice were immunized twice with a 10-day interval with 50 µg OVA-derived MHC class I-matched peptide SIINFEKL in incomplete Freund’s adjuvant. Mice were sacrificed 4–5 days later, and splenocytes (2x105 well) were stimulated for 24 h with specific and irrelevant peptide. The number of IFN-{gamma}-producing cells was evaluated in quadruplicates using ELISPOT assay as described [7 ].

Statistical analysis
Two-paired statistical analysis was performed using JMP statistical software (SAS Institute, Cary, NC).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
We, and others, have demonstrated previously that tumor-derived factors inhibited DC differentiation from HPC [4 , 21 , 23 , 24 ]. VEGF was found to be one of the factors responsible for this defective differentiation [5 , 25 26 27 ]. To identify genes affected by VEGF, we used a DDA. This method is based on systematic PCR amplification of the 3' terminal portions of mRNAs using anchored primers designed to bind to the 5' boundary of the poly-A tails and upstream primers of arbitrary sequences [28 ]. HPC-enriched bone marrow cells isolated from BALB/c mice were incubated with GM-CSF (20 ng/ml) and TNF-{alpha} (5 ng/ml) with or without VEGF (100 ng/ml). RNA was extracted 1, 2, and 3 h later. cDNA was synthesized using three different anchor primers, and PCR reactions were performed with these anchor primers and 16 different arbitrary primers (Gene Hunter, Nashville, TN). PCR fragments, consistently different between samples, were excised, reamplified, and used as probes in Northern blot hybridization. After confirmation of the specificity, the DNA products were sequenced and compared with sequences in Gene Bank. One differentially expressed fragment, the intensity of which was reduced dramatically in the presence of VEGF, matched perfectly over at least 300 bp with the sequence of the mouse H1° histone gene. This suggests that H1° may play a role in DC differentiation.

Expression of h1° is significantly down-regulated by tumor-derived factors
DDA was performed with recombinant VEGF. Several other factors (IL-10, M-CSF, IL-6, gangliosides) were also implicated in defective DC differentiation in cancer. It is apparent that DC deficiency in cancer is caused by a combination of different known and unknown factors. Therefore, we asked whether the combination of tumor-derived factors contained in tumor cell-conditioned media had the same effect on h1° expression as VEGF. Supernatants were obtained from different tumor cell lines as described in the Materials and Methods. To confirm the negative effect of tumor-derived factors on DC differentiation, HPC-enriched bone marrow cells were cultured in the presence of 25% (v/v) tumor cell supernatants for 5 days with a combination of cytokines known to promote DC differentiation (20 ng/ml GM-CSF and 10 ng/ml IL-4). This protocol, described in detail elsewhere, promotes generation of relatively immature, nonactivated DC [21 ]. To obtain activated DC, 5 ng/ml TNF-{alpha} was added to the culture on day 5. Under normal conditions, within 48 h this treatment results in significant up-regulation of MHC class II, costimulatory molecule B7-2, and the ability of DC to stimulate allogeneic T cells. In this study, the presence of conditioned media from tumor cells affected DC differentiation significantly. The proportion of CD11c+ DC and CD11c+B7-2+ mature DC was decreased almost twofold (Fig. 1A ). An ability of cells generated in the presence of conditioned medium from tumor cells to stimulate allogeneic T cells was more than twofold lower than that of cells generated in the presence of conditioned medium from control 3T3 cells (Fig. 1B) . These data indicate that tumor-derived factors inhibit DC differentiation, consistent with previously published observations [4 , 21 , 25 ].



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  		Figure 1. Conditioned media from tumor cells inhibit DC differentiation. Bone marrow cells enriched for HPC as described in Materials and Methods were cultured for 5 days with GM-CSF and IL-4 alone (control) or in the presence of 25% v/v supernatants from 3T3 fibroblasts (3T3) and two tumor cell lines, MethA sarcoma (MethA) and C3 tumor (C3). On day 5, TNF-{alpha} was added to the cultures, and cells were incubated for an additional 48 h. After that time, cells were collected, washed, and analyzed. (A) Phenotype of cells generated in the presence of tumor cell supernatants. Cells were labeled with FITC-conjugated anti-CD11c antibody and PE-conjugated anti-CD86 (B7-2) antibody. (B) Stimulation of allogeneic T cell proliferation. Cells generated from HPC with GM-CSF, IL-4, and TNF-{alpha} in the presence of tumor cell-conditioned media were used as stimulators of control, allogeneic T cells. [3H]-Thymidine uptake was measured in triplicates as described in Materials and Methods. Cell ratios, T cells:cells generated from bone marrow HPC.

 
To investigate possible effects of tumor-derived factors on h1° expression, HPC were cultured with GM-CSF, IL-4, and conditioned media from different tumor cell lines. After 6 h incubation with conditioned media, RNA was extracted, and expression of the h1° gene was analyzed using RT-PCR with subsequent Southern blot hybridization. To confirm the specificity of the primers used for detection of h1° mRNA, we used RNAs from livers of wild-type h1°+/+ and knockout h1°-/- mice. H1°-specific mRNA was detected in h1°+/+ but not in h1°-/- mice (Fig. 2A , lanes 1 and 2). The level of h1° mRNA was elevated following 6 h of incubation of HPC with GM-CSF and IL-4. All tested tumor cell supernatants decreased expression of the h1° gene significantly (4.5-, 3.2-, 40.6-, and 13.5-fold decrease for tumor supernatants in lanes 5–8, respectively; Fig. 2A ). Similar results were obtained after 24 h incubation of the cells with tumor cell supernatants (unpublished results). These data demonstrate that tumor-derived factors inhibit h1° expression in HPC.



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  		Figure 2. Expression of h1° mRNA in HPC during their differentiation into DC. In all experiments, mRNA was analyzed using RT-PCR and Southern blot as described in Materials and Methods. (A) Effect of tumor cell-conditioned media on expression of the h1° gene. RNA was extracted from livers of control h1°+/+ (lane 1), H1°-deficient h1°-/- mice (lane 2), or HPC (land 3–8). HPC were incubated with 20 ng/ml GM-CSF and 10 ng/ml IL-4 for 6 h in the presence of 30% (v/v) tumor cell supernatants. Top lane, h1° histone mRNA; bottom lane, hprt mRNA. Lane 3, Freshly isolated HPC; lanes 4–8, HPC cultured for 6 h with supernatants from control 3T3 fibroblasts (lane 4), MethA sarcoma cells (lane 5), P815 cells (lane 6), L929 cells (lane 7), and C3 tumor cells (lane 8). Three experiments with the same results have been performed. (B) Expression of the h1° gene during DC differentiation. RNA was extracted from HPC immediately after isolation from bone marrow (0) or after incubation for 3, 5, or 7 days. Lanes 1, Incubation with GM-CSF alone; lanes 2, incubation with GM-CSF and IL-4; lane 3, 5-day incubation with GM-CSF and IL-4 followed by a 48-h incubation with GM-CSF, IL-4, and TNF-{alpha}. Three experiments with similar results were performed. (C) Expression of the h1° gene during differentiation of macrophages. RNA was extracted from HPC immediately after isolation from bone marrow (0) or after incubation for 1, 3, 5, or 7 days with 20 ng/ml M-CSF. Two experiments with similar results were performed. (D) Expression of the h1° gene during differentiation of granulocytes. RNA was extracted from HPC immediately after isolation from bone marrow (0) or after incubation for 1, 3, 5, or 7 days with 20 ng/ml G-CSF. Two experiments with similar results were performed. (E) Effect of inhibition of transcription factor NF-{kappa}B on the expression of h1°. HPC were isolated from bone marrow, infected with adenoviruses, and incubated with GM-CSF and IL-4 for 36 h (lanes 1–3) or 72 h (lanes 4–6). Lanes 1 and 4, control, untreated cells; lanes 2 and 5, HPC infected with 100 multiplicity of infection (MOI) of control replication-defective adenovirus; lanes 3 and 6, HPC infected with 100 MOI of I{kappa}B{Delta}N adenovirus.

 
Expression of h1° is associated with DC differentiation
Expression of h1° mRNA was analyzed in HPC during their differentiation into DC in the presence of GM-CSF and IL-4. In parallel, HPC were cultured with GM-CSF alone. Three-day culture of HPC with GM-CSF alone or GM-CSF and IL-4 resulted in a dramatic 9.5-fold up-regulation of h1° mRNA. Activation of DC with TNF-{alpha} did not affect the expression of the h1° gene (Fig. 2B , lane 3). These data indicate that there is an association between expression of the h1° gene and DC differentiation. We also investigated an association between h1° expression and differentiation of other myeloid cells using two growth factors that support differentiation of macrophages (20 ng/ml M-CSF) and granulocytes (20 ng/ml G-CSF). The presence of M-CSF in culture increased h1° expression by day 3 only threefold (Fig. 2C) . This was significantly less than the up-regulation induced by GM-CSF. G-CSF did not affect h1° expression (Fig. 2D) .

Transcription factor NF-{kappa}B plays a critical role in DC differentiation [22 , 29 , 30 ]. Previously, we showed that VEGF affects DC differentiation by inhibiting activation of this transcription factor in HPC [22 ]. The promoter region of h1° contains a sequence (GGGGACCCCG) matching a described possible consensus sequence for NF-{kappa}B [14 , 31 ]. To test the hypothesis that NF-{kappa}B might regulate expression of h1°, we used an adenoviral construct encoding a dominant-negative NF-{kappa}B inhibitor, I{kappa}B{alpha}, with deleted serine phosphorylation sites (I{kappa}B{Delta}N) [22 ]. These mutations prevent dissociation of I{kappa}B{alpha} from NF-{kappa}B and subsequent activation of transcription. HPC were infected with control or I{kappa}B{Delta}N adenoviruses. Cells were cultured with GM-CSF and IL-4 for 36 h or 72 h. After that time, RNA was extracted, and h1° expression was evaluated. The presence of the NF-{kappa}B inhibitor decreased h1° expression almost threefold at both analyzed time points (Fig. 2E) . This observation suggests that NF-{kappa}B is indeed involved in regulating h1° transcription in HPC.

Decreased presence of the DC in H1°-deficient mice
To investigate possible involvement of H1° in DC differentiation, the presence of DC was analyzed in various organs in six pairs of control and H1°-deficient mice. h1° Knockout mice have been described previously [17 ]. H1°-deficient mice were found to have significantly lower numbers of DC in spleens, lymph nodes, and thymuses than control h1°+/+ mice (Fig. 3A 3B 3C ). In spleens, the decrease in DC was confined predominantly to the population of mature CD11c+B7-2+ DC. In thymuses and lymph nodes, the presence of relatively immature CD11c+B7-2- DC also was reduced. The decreased presence of DC in h1°-/- knockout mice was confirmed using allogeneic MLR (an ability to stimulate allogeneic T cells is a unique characteristic of DC). Spleen and lymph node cells obtained from knockout mice had significantly lower stimulatory activity than those obtained from control animals (Fig. 3D) .



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  		Figure 3. Presence of DC in H1°-deficient mice. (A) Population of DC in spleen. Spleens were extracted from five mice per group, and nucleated cells were stained with monoclonal antibodies as described in Materials and Methods. Here and in all other figures, average ± SEM of all experiments is shown. Here and in all other figures, * = statistically significant differences from control h1°+/+ mice (P<0.05). (B) Population of DC in thymuses. Each group included four mice. (C) The number of DC in lymph nodes (LN). Each group included six mice. (D) Stimulation of allogeneic MLR by splenocytes and lymph node cells from H1°-deficient mice. Splenocytes and lymph node cells were isolated from control h1°+/+ and h1°-/- mice, irradiated, and used as stimulators of lymphocytes isolated from control, allogeneic BALB/c mice. Average ± SE of four experiments at stimulator:responder ratio 1:4 is shown.

 
To test the possibility that lack of H1° might affect total production of myeloid cells, we evaluated the number of bone marrow HPC using a colony formation assay. Bone marrow cells isolated from wild-type and knockout mice were cultured in semi-solid methylcellulose medium in the presence of a combination of cytokines supporting growth of erythroid and myeloid colonies. No decrease of hematopoietic progenitor cells was detected in H1°-deficient mice. Moreover, these mice had higher levels of macrophage and GM colonies (CFU-M, CFU-GM) and slightly higher numbers of mixed colonies (Fig. 4 ).



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  		Figure 4. Presence of HPC and effect of FL on DC production in H1°-deficient mice. (A) Bone marrow cells were incubated in replicates in semi-solid methylcellulose medium supplemented with growth factors as described in Materials and Methods. The number of colonies was counted on day 13. Individual results from two control and two h1°-/- mice are shown. (B–D) Control and H1°-deficient mice were treated with FL for 10 days. The number of DC was evaluated in spleens (B) and lymph nodes (C) using flow cytometry. Average ± SE from three independent experiments with spleens and four experiments with lymph nodes is shown. (D) Stimulation of allogeneic MLR by lymph node cells from H1°-deficient mice. Lymph node cells were isolated from control h1°+/+ and h1°-/- mice, irradiated, and used as stimulators of lymphocytes isolated from control, allogeneic BALB/c mice. Average ± SE of two experiments at different stimulator/responder ratios is shown.

 
Decreased FL-inducible DC generation in H1°-deficient mice
The data described above demonstrate that h1° knockout mice have a significantly lower presence of DC than their wild-type counterparts. In control animals, however, DC represent only a minor population of cells. To confirm our findings, we stimulated DC production by administration of FL. Ten days of injections of this growth factor resulted in a dramatic increase in DC production [32 33 34 ]. We hypothesized that if DC differentiation is affected in these knockout mice, it should also manifest in lower responsiveness to FL. Four pairs of control and knockout mice were treated for 10 days with daily s.c. injections of FL (10 µg/mouse). The presence of DC was analyzed in spleens and lymph nodes. h1°-/- Mice had significantly less CD11c+ DC in lymph nodes than control animals (Fig. 4B) . The presence of populations of myeloid CD11c+CD11b+ DC and IAb+B7-2+ mature DC was lowered more than threefold in h1°-/- mice (Fig. 4B) . Significantly lower levels of all populations of DC were also detected in spleens from these animals (Fig. 4C) . Lymph node cells from control mice treated with FL had a high potential to stimulate allogeneic MLR, which reflects an increased proportion of DC in this organ. However, lymph node cells isolated from FL-treated knockout mice had three times lower ability to stimulate allogeneic T cells (Fig. 4D) . Lower numbers of DC in FL-treated mice was confirmed further by staining with anti-NLDC-145 antibody (Fig. 5 ).



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  		Figure 5. DC in tissues of H1°-deficient mice. Samples of spleens (a–d) and liver (e–h) obtained from control and H1°-deficient animals were snap-frozen. Tissue sections (6 µm) were air-dried, fixed in acetone, and stained with NLDC-145 antibodies as described in Materials and Methods. (a, e) Control mice; (b, f) H1°-deficient mice; (c, g) control mice treated with FL; (d, h) H1°-deficient mice treated with FL. Two experiments with the same results were performed. Original magnification, x400.

 
Macrophages and granulocytes develop normally in H1°-deficient mice
We asked whether lack of H1° might affect the development of other myeloid cells. Splenocytes and lymph node cells were stained with antibodies specific for macrophages (F4/80) and granulocytes (Gr-1). In three independently performed experiments, no differences were found between control and h1°-knockout mice in the presence of F4/80+ or Gr-1+ cells (data not shown). Together with the observation that H1°-deficient mice had normal levels of CD11c-CD11b+ macrophages (Fig. 3C) , this result indicates that H1°-deficient mice have normal levels of macrophages and granulocytes. It is possible that defective development of myeloid cells may manifest in abnormal function of these cells. To test this possibility, macrophages and granulocytes were isolated from control and H1°-deficient mice. Several tests were performed to assess function of these cells: the NBT test (spontaneous and stimulated with PMA) to measure production of oxygen radicals and assay of phagocytosis of latex particles (spontaneous and stimulated with LPS or IFN-{gamma}) and TNF-{alpha} production by macrophages (spontaneous and stimulated with LPS or IFN-{gamma}). Granulocytes and macrophages isolated from control and H1°-deficient mice demonstrated equal levels of functional activity (Fig. 6 ). Taken together, these data indicate that lack of H1° did not alter the development of other myeloid cells.



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  		Figure 6. Functional activity of macrophages and granulocytes in H1°-deficient mice. Macrophages and granulocytes were isolated from peritoneal cavities of control and h1° knockout mice as described in Materials and Methods. Peritoneal macrophages were cultured for 24 h on coverslips in six-well plates in complete medium. Medium was changed, and cells were cultured for additional 24 h. Granulocytes were used immediately after isolation. (A) NBT test. Spontaneous (medium) and PMA-stimulated NBT tests were performed. Macrophages were simulated with 10 ng/ml PMA for 24 h and granulocytes, for 1 h. An NBT test was performed as described in Materials and Methods. The results are expressed as AU calculated per 100 cells using a formula NBT test = 1a + 2b + 3b (where a=cells with less than 25% of cytoplasm covered with formazan, b=cells with 25–75% of cytoplasm covered with formazan, and c=cells with more than 75% of cytoplasm covered with formazan). Average ± SE of three performed experiments is shown. (B) Phagocytosis assay. Macrophages and granulocytes were prepared exactly as described above. Macrophages were activated for 24 h with 5 ng/ml IFN-{gamma}, and granulocytes, for 1 h with 10 ng/ml LPS. After washing with PBS, cells were incubated for 1 h at 37°C with polystyrene latex beads (0.807 µm in diameter) at a ratio of 100 particles per cell. Cells were stained as described in Materials and Methods. Phagocytosis was analyzed by microscopy (total magnification, x1000) and expressed as index = (axb)/100 (where a=percentage of cells engulfed latex particles, calculated per 100 cells, and b=number of latex particles engulfed by 100 cells). Average ± SE of three performed experiments is shown. (C) TNF-{alpha} production by peritoneal macrophages. Spontaneous (Medium) or stimulated with 5 ng/ml IFN-{gamma} (IFN) or 10 µg/ml LPS (PMA) TNF-{alpha} production by peritoneal macrophages was measured as described in Materials and Methods. Average ± SE of three performed experiments is shown.

 
The presence and function of T and B lymphocytes and NK cells in H1°-deficient mice
To evaluate T cell development in H1°-deficient mice, single cell suspensions were prepared from thymuses, lymph nodes, and spleens. Cells were labeled with anti-CD3, -CD4, and -CD8 antibodies and were analyzed by flow cytometry. In three separate experiments, H1°-deficient mice had normal levels of T cells in all tested organs (unpublished results). To analyze T cell function, three stimuli that use different pathways of T cell activation were used: Con A, allogeneic DC, and immobilized anti-CD3 and -CD28 antibodies. T cells from h1° knockout mice demonstrated normal levels of response to all these stimuli (Fig. 7 ). The presence of B cells and NK cells was evaluated in lymph nodes and spleens using anti-CD19 and anti-NK 1.1 antibody, respectively. No differences between control and h1° knockout mice were found (data not shown). B cells isolated from spleens of H1°-deficient mice demonstrated normal levels of proliferation in response to LPS (Fig. 7) .



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  		Figure 7. T- and B-cell function in H1°-deficient mice. (A) B cells were isolated from spleens using anti-B220 antibody and magnetic beads. B cells (105/well) were cultured in triplicates in U-bottom 96-well plates for 48 h with and without LPS. Eighteen hours prior to harvest, cells were labeled with 1 µCi [3H]-thymidine, and thymidine uptake was measured on a liquid scintillation counter. T cells were isolated from spleens using purification columns (R&D Systems). T cells (105/well) were stimulated for 72 h with Con A, and cell proliferation was measured as described above. (B) T cells isolated from spleens as described above were stimulated with control, allogeneic DC (from BALB/c mice) of immobilized anti-CD3 and anti-CD28 antibody as described in Materials and Methods.

 
Functional defects in the population of DC from H1°-deficient mice
Stimulation of primary immune responses is the main feature of DC. We asked whether the decrease in DC production in H1°-deficient mice had any functional consequences. DC were generated from HPC from control and h1° knockout mice and then were incubated overnight in complete medium alone or with 1 mg/ml OVA. After that time, cells were washed and used for stimulation of naïve T cells. The population of DC from control mice stimulated a significant, primary proliferative response in control T cells. DC fraction from knockout mice was unable to stimulate such a response in T cells from knockout mice (Fig. 8A ). Although our previous experiments demonstrated normal function of T cells from H1°-deficient mice, it was possible that these T cells were less effective in responding to primary stimulation. To address this concern, we repeated the same experiments using T cells from control mice as responders. Only DC from control mice were capable of inducing a primary T cell response (Fig. 8B) . T cells from h1° knockout mice responded well to stimulation with control DC (Fig. 8C) . These data indicate that the defect in primary T cell stimulation was because of defective DC generation from H1°-deficient HPC.



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  		Figure 8. Function of DC generated from H1°-deficient mice. DC were generated from bone marrow of control and H1°-deficient mice using GM-CSF and IL-4 as described in Materials and Methods. Cells were collected and incubated overnight with 1 mg/ml OVA. After that time, cells were washed and incubated in triplicates with 105 T cells isolated from the same type of mice (A), from control syngeneic mice (B), or from H1°-deficient mice (C). Different DC:T cell ratios have been used. T cell proliferation was measured 4 days later as described above. (D) DC were generated from control and H1°-deficient mice using GM-CSF and IL-4. Cells were loaded with 1 mg/ml OVA overnight, washed, and injected (3x105 cells per mouse) i.p. into control and H1°-deficient mice. Immunization was repeated once 10 days later. Mice were sacrificed after 4–5 days, and single cell suspension of splenocytes was prepared. Splenocytes (105 cells/well) were stimulated for 72 h in triplicates with MHC class II (H2b)-matching peptide ISQAVHAAHAEINEAGR in U-bottom 96-well plates. Cell proliferation was measured using [3H]-thymidine uptake as described above. Spontaneous proliferation of nonstimulated splenocytes was subtracted. Cumulative results of three performed experiments are shown. (E) Mice were immunized twice with 50 µg OVA-derived H2b-specific peptide (SIINFEKL) in incomplete Freund’s adjuvant. Mice were sacrificed after 4–5 days, and splenocytes (2x105 cells/well) were stimulated in quadruplicates with control or specific peptides. The number of IFN-{gamma}-producing cells was evaluated in ELISPOT as described [7 ]. Each group included four mice. Average ± SD is shown. *, Statistically significant differences from control group.

 
Next, we asked whether the same effect could be observed in vivo. DC fractions were prepared and loaded with OVA as described above. Control and H1°-deficient mice were immunized twice by i.p. injections of 3 x 105 DC at a 10-day interval. Mice were sacrificed 5–6 days later, and splenocytes were stimulated with MHC class II-matching OVA-derived peptide or control peptide. As shown in Figure 8D , control mice demonstrated a significant level of peptide-specific response, whereas response of H1°-deficient mice was much lower. Similar experiments were performed to assess CD8-mediated response. Mice were immunized with OVA-derived MHC class I-matching peptide, and the levels of peptide-specific IFN-{gamma}-producing cells were evaluated in ELISPOT assay. H1°-deficient mice demonstrated a more than twofold lower number of IFN-{gamma}-producing cells than control mice (Fig. 8E) . These data indicate that decreased DC production in H1°-deficient mice results in a defective response to vaccination.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we have demonstrated for the first time that the absence of H1° results in defective differentiation of at least one group of hematopoietic cells, DC. H1° deficiency did not lead to a general suppression of bone marrow progenitor cells. On the contrary, the number of myeloid progenitor cells in bone marrow was slightly higher in h1°-/- mice. The finding that lack of H1° histone resulted in only a partial block of DC maturation may suggest that it is involved in transcriptional regulation of genes responsible for DC differentiation. This is consistent with the data demonstrating an important role of H1 histones in gene regulation [9 10 11 ]. It appears that this function is not completely compensated by the other H1 histones present in H1°-deficient mice.

Recently, two major populations of the DC have been identified. These two populations, myeloid-related and lymphoid-related DC, have different origins, phenotypes, and possibly different functions [32 , 35 ]. Myeloid-related DC have the phenotype of CD11c+CD11b+ cells, whereas lymphoid-related DC have the phenotype of CD11c+CD11b-. In our study, lack of H1° had a much more profound effect on differentiation of myeloid than lymphoid DC (Fig. 5) . This suggests that H1° may play an especially important role in differentiation of one particular group of DC, the myeloid-related DC. The effect of decreased production of DC in H1°-deficient mice after treatment with FL is similar to that observed in mice treated with a combination of VEGF and FL [34 ]. This indirectly supports the hypothesis that H1° could be involved in the molecular mechanism of tumor-mediated suppression of DC differentiation. Because DC differentiation requires activation of transcription factor NF-{kappa}B [22 , 29 , 30 ], and tumor-derived factors are known to inhibit this activation, we hypothesized that NF-{kappa}B might be involved in transcriptional regulation of h1°. To test this hypothesis, we blocked NF-{kappa}B activation with a dominant-negative inhibitor and found a significant reduction in h1° mRNA level. These data suggest that h1° expression can be regulated by NF-{kappa}B. Thus, inhibition of h1° via inhibition of NF-{kappa}B in HPC might be one of the molecular mechanisms of defective DC differentiation in cancer.

Our experiments have shown that h1° expression was associated closely with DC differentiation induced by GM-CSF and IL-4. In contrast, M-CSF and G-CSF had little or no effect on h1° expression. The analysis of phenotype and function of granulocytes and macrophages in H1°-deficient mice demonstrated normal function of these cells. It appears that lack of H1° does not affect differentiation and function of macrophages and granulocytes, but rather targets differentiation of DC specifically. The mechanism of association between H1° and DC development is unclear. As it is most likely that H1° works via regulation of expression of other genes, we speculate that H1° may control one or several DC-specific genes. Our data showed that NF-{kappa}B is involved in control of h1° expression. As NF-{kappa}B activation is critically important for differentiation of DC, it is possible that increased h1° expression down-stream of NF-{kappa}B is required for suppression of genes inhibiting DC maturation. Our experiments with DDA showed up-regulation of several bands after treatment with VEGF, consistent with this hypothesis. The nature of these bands is under investigation.

Our findings are consistent with the H1° role described in other experimental systems. DC are terminally differentiated cells. Many studies have shown that H1° accumulates during the process of terminal differentiation [36 37 38 ]. In human diploid fibroblasts, the H1° synthesis rate is increased during the Go phase of cell cycle [39 ]. It was suggested that H1° might prevent binding of the rRNA transcription factor UBF to regulatory sequences of the rRNA and thus reduce ribosomal RNA synthesis and inhibiting cell proliferation [40 ]. This may provide an opportunity for other mechanisms to promote cell differentiation.

The results of this study show that lack of H1° results in decreased production of DC from HPC. We asked whether this decrease would affect the ability of DC to stimulate a primary immune response. To address this question, DC were generated from bone marrow HPC and loaded with OVA. As expected, the proportion of mature DC generated from H1°-deficient HPC was significantly lower than that generated from control HPC. This decrease was sufficient to significantly affect the ability of these cells to induce an OVA-specific, primary T cell response, a function specific for DC (Fig. 8) . This defect was not a result of possible T cell abnormalities in H1°-deficient mice, not only because T cells from these mice showed normal levels of responses to various stimuli, but also because DC generated from control mice were effective in stimulating T cells from h1° knockout mice. At the same time, cells generated from H1°-deficient HPC were unable to stimulate an OVA-specific response in control T cells. To assess the response of mice to immunization with OVA, we used specific OVA-derived peptides recognized by MHC class II and class I. Splenocytes from control mice immunized with OVA-loaded DC responded to stimulation with OVA-derived peptides. The defect in DC production in h1°-knockout mice made their immunization with soluble protein ineffective. H1°-deficient mice responded poorly to immunization with OVA-loaded cells as well as to peptide in Freund’s adjuvant. These data are similar to that obtained in tumor-bearing mice [25 ] and suggest that an H1° deficit has significant, functional consequences.

In conclusion, this study demonstrates that H1° is involved in differentiation of DC but not in differentiation of other myeloid cells. It appears that tumor-derived factors inhibit h1° expression, and their inhibition may play an important role in the defective DC differentiation in cancer.


   ACKNOWLEDGEMENTS
 
This study was supported by National Institutes of Health grant CA 84488 (D. I. G.).

Received January 16, 2002; revised March 4, 2002; accepted March 6, 2002.


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 DISCUSSION
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