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Journal of Leukocyte Biology logoLink to Journal of Leukocyte Biology
. 2008 Jun 27;84(3):824–834. doi: 10.1189/jlb.0807583

Epigenetic mechanisms of age-dependent KIR2DL4 expression in T cells

Guangjin Li 1, Cornelia M Weyand 1, Jörg J Goronzy 1,1
PMCID: PMC2516893  PMID: 18586981

Abstract

Killer Ig-like receptor (KIR) expression is mostly restricted to NK cells controlling their activation. With increasing age, KIRs are expressed on T cells and contribute to age-related diseases. We examined epigenetic mechanisms that determine the competency of T cells to transcribe KIR2DL4. Compared with Jurkat cells and CD4+CD28+ T cells from young individuals, DNA methyltransferase (DNMT) inhibition was strikingly more effective in T cells from elderly adults and the CD4+CD28− T cell line HUT78 to induce KIR2DL4 transcription. In these susceptible cells, the KIR2DL4 promoter was partially demethylated, and dimethylated H3-Lys 4 was increased, and all other histone modifications were characteristic for an inactive promoter. In comparison, NK cells had a fully demethylated KIR2DL4 promoter and the full spectrum of histone modifications indicative of active transcription with H3 and H4 acetylation, di- and trimethylated H3-Lys 4, and reduced, dimethylated H3-Lys 9. These results suggest that an increased competency of T cells to express KIR2DL4 with aging is conferred by a selective increase in H3-Lys 4 dimethylation and limited DNA demethylation. The partially accessible promoter is sensitive to DNMT inhibition, which is sufficient to induce full transcription without further histone acetylation and methylation.

Keywords: molecular biology of aging, deacetylase, cellular immunology, cellular senescence, gene expression

INTRODUCTION

Killer Ig-like receptors (KIRs) comprise a diverse family of regulatory cell-surface receptors that are preferentially expressed on NK cells [1,2,3]. KIRs are glycoproteins characterized by the presence of a long or short cytoplasmic domain. KIRs with long cytoplasmic tails are inhibitory receptors containing one or two ITIMs; KIRs with short tails bind adaptor molecule DAP12 with stimulatory kinase activity [4,5,6,7]. Upon binding with their HLA class I ligands, KIRs transduce a negative signal that prevents the cytotoxicity of NK cells or an activating signal that leads to the target cell lysis. The negative regulatory receptors are the molecular basis for the missing self-hypothesis, explaining why NK cells are tolerized to self, but recognize tumor cells that have lost MHC class I expression. The role of stimulatory KIRs is less clear, but they may influence resistance to infections and susceptibility to autoimmune diseases [1, 8, 9].

KIR expression on NK cells is stochastic. Each NK cell expresses and maintains an individual set of KIRs that represents the major mechanism of NK-cell repertoire diversification [3, 10]. KIR expression patterns on NK cells are mainly controlled at the transcriptional level [11]. All clonally distributed KIR genes share a remarkably high homology in their 5′-untranslated region with >91% nucleotide identity, suggesting that KIRs are regulated by similar transcriptional mechanisms [12]. How expression patterns are established and maintained has therefore been of particular interest. Current evidence points to epigenetic mechanisms; in particular, DNA methylation is of central importance. Hypermethylated CpG in KIR promoters correlated with transcriptional suppression, whereas hypomethylation corresponded to active KIR transcription. Although histone acetylation and methylation are also involved in KIR transcriptional regulation, DNA methylation primarily controls the clonally restricted and allele-specific KIR transcription in NK cells [13]. The two promoters best studied in NK cells are KIR3DL1 and KIR2DL4. The KIR2DL4 promoter has 61% sequence identity with other KIR members, in contrast to the >91% sharing, typical for the KIR family [14]. KIR2DL4 is the only member that is expressed in most or possibly all NK cells, in contrast to the clonal distribution seen in all other KIRs [15].

KIRs are also expressed on T cells, including γδ T cells and small subsets of CD4+ and CD8+ T cells [16,17,18]. Unlike in NK cells, where KIR expression occurs during NK-cell development, KIRs are absent in T cells of neonates. KIR expression in T cells increases with age and is mostly found on senescent memory T cells that have lost CD28 molecule expression [19, 20]. Acquisition of KIRs in T cells is also stochastic and appears to occur successively during clonal expansion [17, 21, 22]. Why T cells start to express KIRs is unclear. It has been suggested that inhibitory KIRs impair immune responses to viruses and tumor antigens, and their increased expression with age contributes to the immune defects in the elderly, generally termed immunosenescence [23,24,25,26,27,28]. In contrast to NK cells, KIRs do not completely inhibit T cell activation but modify the spectrum of effector functions and preferentially inhibit cytokine production while leaving cytotoxic function intact [29, 30]. Stimulatory KIRs have been implicated in a number of autoimmune diseases such as rheumatoid vasculitis, type I diabetes, and psoriatic arthritis [2, 31, 32] and also in tissue-injurious, inflammatory responses such as atherosclerosis [33]. The disease-promoting activities of KIRs may be related directly to their expression on T cells and may explain the increasing incidence of these inflammatory diseases with age [34].

The transcriptional mechanisms that lead to increasing KIR expression with age are unknown. Studies in naïve T cells and in KIR+ and KIR− memory T cells have shown that all T cell subtypes are able to support the activity of the minimal KIR promoter [35]. However, although inhibition of DNA methyltransferase (DNMT) in NK cells is sufficient to activate KIR genes and convert a clonally distributed expression pattern to full expression of all KIR variants, treatment of Jurkat T cells does not induce KIR expression [36]. We hypothesized that KIR activation in T cells with aging is a step-wise process with histone modifications as an early epigenetic step, which then renders the promoter sensitive to DNA demethylation. To examine this hypothesis and to identify the mechanisms of KIR expression in T cells with age, we examined the epigenetics of a KIR promoter in T cells and compared the results with NK cells.

MATERIALS AND METHODS

Cells

Peripheral blood was obtained from individuals aged 20–30 and 70–80 years. Exclusion criteria included the presence or a history of cancer, uncontrolled hypertension, diabetes mellitus, or any chronic inflammatory or autoimmune disease. All elderly individuals were independently living and in good health. Appropriate written informed consent was obtained, and the Emory Institutional Review Board (Atlanta, GA, USA) approved the study.

Mononuclear cells were isolated by Ficoll (Mediatech, Inc., Manassas, VA, USA) density gradient centrifugation. CD4+CD28+ and CD4+CD28− T cells were purified by FACSVantage (BD Biosciences, San Jose, CA, USA) and stimulated with anti-CD3 (OKT3, Ortho Diagnostic Systems, Raritan, NJ, USA), immobilized at 1 μg/ml to generate short-term T cell lines. CD4+ and CD8+ cells were positively selected using magnetic microbeads (Miltenyi Biotec, Auburn, CA, USA). Jurkat and HUT78 T cell lines were maintained as described previously [35]. The NK3.3 NK cell line was kindly provided by Charles T. Lutz (University of Kentucky, Lexington, KY, USA). For the flow cytometric analysis of KIR2DL4 expression, cells were stained with PE-conjugated anti-KIR2DL4 mAb (R&D Systems, Minneapolis, MN, USA) and analyzed on a LSRII cytometer. HUT78 cells were separated into cells expressing intermediate (upper 30%, KIR2DL4int) and not detectable levels of KIR2DL4 (lower 30%, KIR2DL4low) by cell sorting using a FACSVantage. For the DNMT inhibition experiments, cells were cultured with recombinant IL-2 (50 U/ml) and Dynabeads CD3/CD28 T cell expander (Dynal Biotech, Norway). On Day 3, 5-aza-2′-deoxycytidine (5-Aza-dC; Sigma-Aldrich, St. Louis, MO, USA) was added at a final concentration of 1 μM. After 72 h treatment, the cells were harvested to assess KIR2DL4 transcription.

Real-time quantitative (q)RT-PCR

Real-time qRT-PCR was conducted to detect KIR2DL4 expression. For quantitative assays, KIR2DL4 transcripts were quantified using the Mx3000 PCR instrument (Stratagene, La Jolla, CA, USA). The number of KIR2DL4 transcripts was normalized to 1 × 107 β-actin transcripts, and results are given as relative transcript numbers. Primers used were 5′-CTTCGGCTCTTTCCATGGA-3′ and 5′-CACTGAGTACCTAATCACAG-3′ for KIR2DL4 and 5′-ATGGCCACGGCTGCTTCCAGC-3′ and 5′-CATGGTGGTGCCGCCAGACAG-3′ for β-actin.

Bisulfite sequencing and methylation-specific PCR

Genomic DNA was isolated with a QIAamp blood kit (Qiagen, Valencia, CA, USA), and 1 μg purified DNA was treated with sodium metabisulfite. Bisulfite-modified DNA (50 ng) was amplified by PCR using primers 5′-TGTAGGGGTAAGTGAGTTTGAGAT-3′ and 5′-TATACTACCTCCCTCCCATTTC-3′. The PCR conditions were 95°C for 4 min; followed by 35 cycles of 95°C for 1 min, 50°C for 1 min, 72°C for 2 min; and 72°C for 30 min. The resulting amplification products were cloned into the pCR2.1-topoisomerase (TOPO) vector using a TOPO-TA cloning kit (Invitrogen Life Technologies, Carlsbad, CA, USA). Individual subclones were isolated and sequenced.

For quantitative assessment of the methylation at specific CpG sites, PCR products were analyzed using qPCR with 3′-locked nucleic acid primers. For CpG –24/–34, the forward primer for unmethylated DNA (U) and methylated DNA (M) was 5′-GTAGAAGAAGTTTATTTATGTTT-3′; the reverse primer for U was 5′-CTACTACCAAAACACAATAACTCA-3′ and for M was 5′-CTACTACCAAAACGCAATAACTCG-3′. For CpG –47, the forward primers were 5′-GGGTTTTTTATTATATTTTTTGTATT-3′ (U) and 5′-GGGTTTTTTATTATATTTTTTGTATC-3 (M), respectively; the reverse primer for U and M was 5′-CACCAAAACATACCAAAATAATAACC-3′. For CpG –223 and –228, the forward primers were 5′-TTTTGAGTTTGGTTGTTGT-3′ (U) and 5′-TTTTGAGTTTGGTCGTTGC-3′ (M), and the reverse primer for U and M was 5′-CACATTAACCACAACATA-3′. Amplification efficiencies with the primer sets were similar. The reactions were performed in a final volume of 20 μl containing 12.5 pM each primer, 4 mM MgCl2, and 2 μl 1,000,000-fold, diluted PCR products. To correlate the comparative threshold values to copy numbers, standard curves were generated using serial dilutions of plasmids containing U or M PCR products.

In vitro methylation, transfection, and reporter gene assays

The pGL3 luciferase construct, containing the wild-type (WT) KIR2DL4 promoter, spanning from base –6 to base –262 relative to the translation initiation codon, was kindly provided by Dr. John Trowsdale (University of Cambridge, Cambridge, UK). The plasmids were methylated in vitro with SssI methylase (New England Biolabs, Beverly, MA, USA), as described by Bruniquel and Schwartz [37]. Complete methylation was verified by digestion of the plasmids with restriction enzyme HpaII. HUT78 cells were transfected with 2 μg each WT KIR2DL4 construct and the methylated KIR2DL4 construct. pGL3-basic vector or pGL3-promoter vector served as negative and positive control. pRL-TK (Promega, Madison, WI, USA; 0.5 μg) was cotransfected as an internal control. Transfection was performed using Cell Line Nucleofector Kit R (Amaxa, Germany). Forty-eight hours after transfection, the cells were harvested, lysed, and assayed using the Dual Luciferase Reporter Assay System (Promega). Luciferase activity was normalized to the internal control.

Chromatin accessibility assay

Chromatin remodeling was performed using chromatin accessibility real-time (CHART)-PCR [38]. Cell nuclei were prepared as described [39]. Nuclei were washed in restriction enzyme buffer [10 mM Tris (pH 7.4), 50 mM NaCl, 10 mM MgCl, 0.2 mM EDTA, 0.2 mM EGTA, 1 mM β-ME, 0.15 mM spermine, and 0.5 mM spermidine], resuspended in digestion buffer (New England Biolabs), and digested with 40 units restriction enzymes at 37°C for 20 min. Genomic DNA was isolated using a QIAamp blood kit (Qiagen). DNA (100 ng) from restriction enzyme-digested samples and undigested control was amplified by SYBR real-time PCR with the primer set 5′-CGCATGATGTGAAGTGACAAGTCT-3′ and 5′-TCGACTGACCGGTGCAGAGGATGT-3′. The chromatin accessibility was calculated as (1–DNAdigested/DNAundigested).

Chromatin immunoprecipitation (ChIP)

Cells (1×106 per ChIP assay) were cross-linked with 1% formaldehyde for 20 min at room temperature, resuspended in lysis buffer [1% SDS, 10 mM EDTA, 50 mM Tris-HCl, (pH 8.1)], incubated on ice for 10 min, and then sonicated. The samples were precleared and then incubated with specific antibodies or normal rabbit IgG overnight at 4°C. The antibodies used were: antiacetyl-H3, antiacetyl-H4, antidimethyl-H3-Lys 4, antitrimethyl-H3-Lys 4 (all from Upstate Biotechnology, Lake Placid, NY, USA), antidimethyl-H3-Lys 9 (Abcam, Cambridge, MA, USA), and anti-DNMT1 (C-term S1602, Abgent, San Diego, CA, USA). Immunocomplexes were purified using salmon sperm DNA/protein A agarose. Eluted immunocomplexes were treated with proteinase K. DNA was extracted and quantified by qPCR using primers 5′-CCGTTGCGCATGATGTGAAGTGAC-3′ and 5′-TGACCGGTGCAGAGGATGTGGT-3′. The region analyzed covers –42 to –229 bp. Specifically, IP DNA was calculated with the following equation: (DNAspecific IP–DNAcontrol IP)/DNAinput; DNAspecific IP = amount of DNA IP using a specific antibody; DNAcontrol IP = amount of DNA IP with nonspecific IgG; DNAinput = sheared chromatin prior to IP.

RESULTS

Correlation between KIR2DL4 promoter demethylation and KIR2DL4 transcription in T cells

CD28 loss is the most consistent phenotypic change in T cells associated with aging [40, 41]. KIR transcription is largely restricted to CD28− cells. KIR2DL4 transcripts are virtually absent in CD4+CD28+ T cells, and CD4+CD28− T cells have ∼2000 copy numbers relative to 1 × 107 β-actin transcripts (Fig. 1A). To identify a model system to examine the mechanisms underlying KIR expression in T cells, we screened T cell lines for KIR2DL4 expression. In previous studies, we showed that the CD4 T cell line HUT78 resembles senescent T cells in several aspects including CD28 transcription loss, and Jurkat T cell lines continue to express CD28 [42]. Screening of these two T cell lines by real-time PCR demonstrated that the transcriptional activity for KIR2DL4 in Jurkat cells did not exceed background levels, and HUT78 T cells produced ∼1500 copies per 107 β-actin transcripts (Fig. 1B). Of interest, the KIR2DL4 transcript levels in HUT78 were comparable with CD4+CD28− T cells but reached only ∼6% of the transcription seen in the NK cell line NK3.3. Flow cytometric studies did not detect any cell-surface KIR2DL4 on Jurkat cells, and KIR2DL4 was highly expressed on NK3.3 cells (Fig. 1C). The staining of HUT78 was unimodal and shifted compared with background levels, without a population with expression levels similar to NK cells. The staining pattern is suggestive for a universally low KIR2DL4 transcription in HUT78 cells and not a clonally distributed expression pattern with a small percentage of KIR2DL4-expressing cells. This interpretation was confirmed when HUT78 cells with KIR2DL4 expression levels in the upper and lower 30th percentiles were compared. Both sorted subpopulations had equal numbers of transcripts (P=0.8; Fig. 1D).

Fig. 1.

Fig. 1.

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KIR2DL4 expression in T cells and NK3.3 cells. (A) KIR2DL4 transcription was quantified by real-time PCR in CD4+CD28+ and CD4+CD28− T cells. The results are shown as KIR2DL4 transcript numbers after normalization for 1 × 107 β-actin transcripts and represent mean ± sd from three independent experiments. Expression was higher in CD4+CD28− than in CD28+ T cells (P=0.02). (B) KIR2DL4 transcription in T cell lines and a NK3.3 cell line; results are presented as mean ± sd from four independent experiments. Expression in HUT78 was higher than in Jurkat cells but lower than in NK3.3 cells (P<0.001). (C) Cells were stained with PE-labeled anti-KIR2DL4 mAb (solid lines) and analyzed by flow cytometry. Staining with PE-IgG served as control (dashed lines). Histograms representative of three independent experiments show unimodal distributions of low (HUT78) and high (NK3.3) KIR2DL4. (D) KIR2DL4 transcription was not different in KIR2DL4low (lower 30% in mean fluorescence intensity) and KIR2DL4int (upper 30%) HUT78 cells, suggesting that KIR2DL4 in HUT78 expression was global and not clonally distributed. Results are shown as transcripts relative to 1 × 107 β-actin transcripts and presented as mean ± sd from two independent experiments (P=0.8).

We selected the T cell lines Jurkat and HUT78 and the NK cell line NK3.3 as model systems to examine the epigenetic changes of the KIR2DL4 promoter that induce transcriptional activity. Previous studies of KIR family members in NK cells have assigned DNA demethylation a central role [13, 36]. The proximal KIR2DL4 promoter has 10 CpG dinucleotides between +18 and –228 bp. Bisulfite sequencing results are shown in Figure 2A. Almost all CpG dinucleotides within the inspected region were demethylated in NK3.3 cells (upper right panel). CpG sites in Jurkat and HUT78 cells were partially methylated but exhibited differences in methylation patterns, in particular, at the four CpG sites between the –47 and –24 bp region. Similar differences in methylation patterns at CpG –24, –34, and –47 were seen for CD4+CD28+ and CD4+CD28− T cells (Fig. 2A). To quantify the methylation pattern, real-time methylation-specific PCR to analyze the CpG dinucleotides at –24/–34, –47, and –223/–228 bp was established. The assays were validated using promoter plasmids containing C or T at these positions. As shown in Figure 2B, the methylation level in Jurkat cells at all CpG sites tested was higher than 75%. In comparison, 40–50% of HUT78 chromosomes were demethylated at –24/–34 and –47, and CpG at –223/–228 remained methylated. The methylation patterns were not different in HUT78 sorted for higher or lower cell-surface expression, again documenting that the results reflect the overall HUT78 population and not a subset of cells transcribing KIR2DL4. Studies in normal CD4+CD28+ and CD4+CD28− T cells yielded a similar promoter methylation pattern as in the tumor lines. KIR2DL4 methylation of the CpG dinucleotides at –24/–34 and –47 bp was much lower in CD4+CD28− than in CD4+CD28+ T cells (∼60% vs. 90%, respectively), as assayed by real-time PCR (P<0.001; Fig. 2B).

Fig. 2.

Fig. 2.

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CpG methylation of the KIR2DL4 core promoter. (A) The methylation patterns of CpG sites in the KIR2DL4 core promoter in Jurkat, HUT78, NK3.3, CD4+CD28+, and CD4+CD28− cells were analyzed by bisulfite sequencing. Genomic DNA was treated with sodium bisulfite and amplified with primer pairs encompassing the +18 to –228 region. The amplification product was subcloned and sequenced. Each row represents an individual subclone. Closed symbols indicate methylated CpG sites; open symbols indicate unmethylated CpG sites. The nucleotide position of CpG sites is indicated relative to the first nucleotide of the translation initiation codon. (B) DNA methylation statuses at CpG –24/–34, –47, and –223/–228 were quantified for Jurkat cells and HUT78 cells with low and intermediate KIR2DL4 cell surface expression and CD4+CD28+ and CD4+CD28− T cells by methylation-specific real-time PCR using 3′-locked nucleic acid primers specific for the methylated and unmethylated sequences after bisulfite treatment. Results are shown as mean ± sd of triplicate copy numbers of methylated DNA/copy numbers of methylated + unmethylated DNA and are representative of three experiments. KIR2DL4 promoter methylation of the CpG dinucleotides at –24/–34 and –47 bp was lower in HUT78 and in CD4+CD28− than in Jurkat cells and CD4+CD28+ T cells (P<0.001). (C) Recruitment of the DNMT1 to KIR2DL4 promoter was assayed by ChIP with antibody to DNMT1. The recovered DNA was quantified by real-time PCR using primers specific for the KIR2DL4 promoter. Results are calculated as the ratio of DNAspecific IP – DNAcontrol IP:DNA input and are shown as mean ± sd of triplicate precipitations.

Chromatin-associated DNMT1 in T cells

The studies showed regional but not full demethylation of the KIR2DL4 promoter in HUT78 cells. DNA methylation in mammalian cells is mainly carried out by three DNMT enzymes: DNMT1, DNMT3a, and DNMT3b, of which DNMT1 is responsible for methylation maintenance. To examine whether DNMT1 is recruited to the KIR2DL4 promoter and plays a role in KIR2DL4 methylation, ChIP assays were performed with Jurkat, HUT78, and NK3.3 cells. As shown in Figure 2C, DNMT1 recruitment to the KIR2DL4 promoter was detected in both T cell lines, but not in NK3.3 cells, consistent with the observation that the promoter is not methylated in NK cells. Recruitment was not different between Jurkat and HUT78 cells (P=0.9), although they differ in methylation. In contrast to the NK cell line, the selective demethylation in HUT78 appears to be unrelated to the lack of global DNMT1 recruitment.

Differential effects of DNMT inhibition on KIR2DL4 transcription in HUT78 and Jurkat cells

Jurkat, HUT78, and NK3.3 cells were treated with the DNMT inhibitor 5-Aza-dC for 12, 24, 48, and 72 h, and KIR2DL4 transcription was quantified by real-time PCR. KIR2DL4 expression was already maximal and could not be increased further in the NK cell line (Fig. 3A). DNA demethylation clearly induced KIR2DL4 transcription in HUT78 T cells. In contrast, only minute transcriptional activity was induced in Jurkat T cells at 48 h and 72 h after 5-Aza-dC treatment. In HUT78, transcripts after DNMT inhibition reached levels equaling transcription in the NK cell line. In comparison, Jurkat cells had <4% of the activity in NK3.3 cells. Assessment of the CpG methylation status after treatment showed that HUT78 was more sensitive to the action of 5-Aza-dC. DNMT inhibition resulted in a complete demethylation in positions –24/–34 of the KIR2DL4 promoter in HUT78 cells, and the effect in Jurkat cells was minimal (Fig. 3B). Interestingly, CpG methylation at positions –47 and –223/–228 did not significantly change in both tumor lines. These data suggest that demethylation at positions –24/–34 is sufficient and necessary for KIR2DL4 expression. Methylation at these positions appears to be protected in Jurkat cells, rendering these cells relatively resistant to 5-Aza-dC treatment and unable to express KIR2DL4. Given the limited effect of DNMT inhibition on the methylation pattern in HUT78, although these cells transcribed KIR2DL4, it is also possible that DNMT1 suppressed KIR2DL4 promoter activity through a direct transcriptional suppression as recently reported [43], and the increased transcription with 5-Aza-dC treatment reflected the release of this suppression rather than promoter demethylation.

Fig. 3.

Fig. 3.

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The effects of DNA demethylation on KIR2DL4 transcription. (A) KIR2DL4 transcription in Jurkat, HUT78, and NK3.3 was measured by real-time PCR after DNMT inhibition. Cells were harvested after 12, 24, 48, and 72 h culture in the presence or absence of 1 μM 5-Aza-dC. The results are shown as KIR2DL4 transcript numbers after normalization for 1 × 107 β-actin transcripts and represent mean ± sd from three independent experiments. (B) DNA was purified from the cells after 72 h DNMT inhibition. KIR2DL4 methylation was measured and calculated as described in Figure 2B. (C) Promoter activity of a methylated and unmethylated KIR2DL4 promoter construct (–6 bp to –262 bp) was assessed after transfection into HUT78 cells. pGL3-basic vector and pGL3-promoter vector were used as negative and positive controls. Firefly luciferase activities were normalized to pRL-TK Renilla luciferase activity, which served as an internal control. Methylation decreased KIR2DL4 promoter activity significantly (P=0.0004). The results represent mean ± sd from three independent experiments.

To directly examine the role of methylation in KIR2DL4 transcription, we performed luciferase reporter gene assays. We used a fragment spanning –6 bp to –262 bp of the KIR2DL4 promoter that has been identified as the minimal promoter in a previous study [14]. The pGL3 luciferase construct containing the KIR2DL4 promoter fragment was methylated in vitro. HUT78 cells were transected with methylated or unmethylated plasmid. The luciferase activities, assayed 48 h after transfection, normalized to the internal control pRL-TK Renilla luciferase activity, are shown in Figure 3C. Methylation treatment reduced KIR2DL4 promoter activity to background levels, significantly lower than the activity observed with the unmethylated promoter (P=0.0004). Although a contribution of vector backbone methylation to the transcriptional repression cannot be excluded completely, these data at least support the notion that methylation of the KIR2DL4 promoter is directly involved in activity control.

Effect of age on KIR2DL4 promoter control

To examine whether these observations in T cell tumor lines are relevant for the age-dependent expression of KIR2DL4 in T cells, CD4+ and CD8+ T cells and NK cells were isolated from two different age groups (20–30 and 70–80 years) and examined for KIR2DL4 expression and promoter CpG methylation (Fig. 4, A and B). In young CD4 T cells, the KIR2DL4 promoter was largely methylated at positions –24/–34 and –47 (Fig. 4B). Age only had a minor effect on position –24/–34, consistent with the finding that CD4 T cells are resistant to undergoing phenotypic changes with age [41]. In contrast, methylation patterns changed with age in CD8 T cells. Already in young adults, the KIR2DL4 promoter was slightly more demethylated at positions –24/–34 (P=0.04) and –47 (P=0.01) in CD8 than in CD4 T cells. With age, both positions were demethylated significantly (P=0.01 and 0.004, respectively) to an extent seen in NK cells. Transcriptional studies supported this finding (Fig. 4A). KIR2DL4 transcription was quantified by real-time PCR. The number of KIR2DL4 transcripts in CD8 T cells increased with age (P=0.04). However, it did not reach the transcript number seen in NK cells (P=0.004) but was in the same range as in HUT78 cells. Accordingly, cell-surface expression was low (data not shown). The transcriptional activities increased markedly when CD4 and CD8 T cells were activated in vitro and cultured in the presence of 5-Aza-dC for 3 days (Fig. 4C). The effect of DNMT inhibition on KIR2DL4 transcription was significantly more pronounced in the elderly for CD4 T cells (P=0.01) and CD8 T cells (P=0.001). These data demonstrate that KIR2DL4 transcription can be induced in T cells by DNMT inhibition and that the KIR2DL4 promoter in T cells from the elderly is more susceptible.

Fig. 4.

Fig. 4.

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Influence of age on KIR2DL4 transcription and methylation status in CD4+ and CD8+ T cells. CD4 and CD8 T cells and NK cells were isolated from 20- to 30 (light gray boxes)- and 70- to 80 (dark gray boxes)-year-old individuals, and KIR2DL4 transcription levels (A) and methylation levels (B) were analyzed by real-time PCR. Results are shown as box plots of 13 young and 12 old individuals. CD8 T cells from young individuals had lower transcription and higher methylation levels than those from the elderly. (C) CD4+ and CD8+ cells from eight young and eight elderly, healthy individuals were cultured in the absence or presence of 1 μM 5-Aza-dC for 72 h before KIR2DL4 transcription was quantified. Transcription levels were expressed as transcripts relative to 1 × 107 β-actin transcripts. Before DNMT inhibition, relative transcripts were below 100 in CD4 and in CD8 T cells from young individuals. CD8 T cells from elderly individuals had increased relative copy numbers with a median of 550. Upon DNMT inhibition, KIR2DL4 transcription increased in all individuals but significantly more so in CD4 and CD8 T cells from the elderly. Results are shown as box plots with medians, 25th and 75th percentiles as boxes, and 10th and 90th percentiles as whiskers.

Nucleosome remodeling of the KIR2DL4 promoter

To identify the mechanisms of this increased sensitivity of elderly T cells and the tumor line HUT78 to DNMT inhibition, we compared chromatin remodeling in the two T cell lines HUT78 and Jurkat. Accessibility was examined with CHART-PCR [38]. Stewart et al. [14] have recently mapped a fully functional KIR2DL4 promoter to the first 260 bp. MNase Southern blot of Jurkat cells demonstrated a nucleosomal structure with N1 covering the minimal promoter (Fig. 5A). A PstI cleavage site is located within N1; the amount of product generated by CHART-PCR after PstI cleavage is therefore inversely proportional to chromatin accessibility. Nuclei were digested with PstI; genomic DNA were purified and subjected to real-time PCR amplification using primers encompassing the PstI site. As shown in Figure 5B, N1 nucleosome is 80% accessible in NK3.3 cells, ∼40% in HUT78 cells, and inaccessible in Jurkat cells. As a negative control, nuclei were digested with restriction enzyme EcoRI, which has a cleavage site at –285 outside of the amplified fragment. The ratio of amplified DNA generated from EcoRI-digested samples to those of undigested controls was similar in three cell lines (data not shown). These results suggest that N1 is in an open configuration in NK3.3 cells, partially relaxed in HUT78 cells, and tightly assembled in Jurkat cells.

Fig. 5.

Fig. 5.

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Nucleosome remodeling of the KIR2DL4 promoter. (A) Schematic representation of the nucleosome map. The translation initiation codon, the position of primer used for the CHART-PCR, and the XmnI and PstI cleavage sites relative to the first nucleotide of the translation initiation codon are indicated. (B) Nuclei from Jurkat, HUT78, and NK3.3 were digested with PstI; genomic DNA were purified and subjected to real-time PCR with primers indicated in A. Results were normalized to uncut DNA and are expressed as a percent accessibility. Data shown are the mean ± sd from three independent experiments.

Histone acetylation is uncorrelated with differential KIR2DL4 transcription in T cells

Chromatin structures are controlled by post-translational modification of the N-terminal histone tail regions, including acetylation, phosphorylation, ubiquitination, and methylation. Specific modifications have been correlated with gene activation or silencing. Acetylation of histones H3 and H4, which has been correlated with active transcription, was analyzed using a ChIP assay for the proximal KIR2DL4 promoter. Acetyl H3 and H4 were present in Jurkat cells at a low level. Surprisingly, histone acetylation was not higher in HUT78 cells, although they have a partially open nucleosome N1 and transcribe KIR2DL4 (P=0.9 compared with Jurkat cells). In contrast, H3 and H4 were acetylated in NK3.3 cells, consistent with their more active gene transcription when compared with Jurkat and HUT78 cells (Fig. 6A). KIR2DL4 transcription was not induced in HUT78 or Jurkat T cells when the cells were cultured in the presence of the histone deacetylase inhibitor trichostatin A (data not shown).

Fig. 6.

Fig. 6.

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Histone acetylation and methylation of the KIR2DL4 promoter in T cells. (A) Chromatin was precipitated with antiacetyl-H3 and antiacetyl-H4. (B) H3 dimethyl-Lys 4, H3 trimethyl-Lys 4, and H3 dimethyl-Lys 9 were precipitated with specific antibodies. The recovered DNA was subjected to real-time PCR for KIR2DL4 sequences. Results shown are mean ± sd of three independent experiments.

KIR2DL4 chromatin remodeling in T cells includes selective shifts in histone 3-Lys 4 methylation

As histone acetylation appears not to be involved in the initial chromatin remodeling of the KIR2DL4 promoter in T cells, we were interested in the contribution of histone methylation. Lys 4 methylation of the H3 tail is generally correlated with an active and Lys 9 methylation with a repressed transcriptional state. ChIP assay using a specific antibody to H3 dimethyl-Lys 4 revealed a sixfold higher level in HUT78 cells than in Jurkat cells (P=0.007; Fig. 6B). In contrast, H3 trimethyl-Lys 4 was hardly detectable in T cells and in particular, not different between Jurkat and HUT78 cells (P=0.96; Fig. 6B). H3 dimethyl-Lys 4 and H3 trimethyl-Lys 4 were abundantly detected to be associated with the KIR2DL4 promoter in NK3.3 cells. H3 dimethyl-Lys 4 was 14-fold and 2.5-fold higher in NK3.3 cells than in Jurkat and HUT78, respectively; H3 trimethyl-Lys 4 was tenfold higher than that in Jurkat or HUT78 cells. In contrast to H3-Lys 4, H3 dimethyl-Lys 9 was highly associated with the KIR2DL4 promoter in Jurkat cells, essentially unchanged in HUT78 cells, and very low in NK3.3 cells (Fig. 6B). These results show that the histone modification of the active KIR2DL4 promoter in NK cells is characterized by acetylated H3 and H4, increased H3 dimethyl-Lys 4 and H3 trimethyl-Lys 4, and decreased H3 dimethyl-Lys 9, consistent with the current paradigm of gene activation. Of these histone modifications, only increased H3 dimethyl-Lys 4 is seen in HUT78 T cells that have a relaxed nucleosome and partial activation of KIR2DL4 transcription.

DNA demethylation induces full KIR2DL4 promoter expression without further changes in histone acetylation and methylation

Results shown in Figure 3 demonstrated that inhibition of DNMT was sufficient to dramatically augment KIR2DL4 transcription in HUT78 cells and to bring it to the same level that is seen in NK cells. We hypothesized that the fully active promoter in HUT78 cells after DNA demethylation would also assume the histone modifications that are characteristic for the promoter in NK cells. HUT78 were treated with 5-Aza-dC for 72 h, at which time, ChIP assays for acetylated and methylated histones were performed, which covered the –42- to –229-bp region. Results are shown in Figure 7. The active KIR2DL4 promoter in HUT78 did not show any more histone modifications than the minimally active promoter. Acetylated H3 and H4 remained low to undetectable, H3 dimethyl-Lys 9 remained high, and H3 dimethyl-Lys 4 did not increase further, suggesting that full transcription can be achieved without these histone modifications.

Fig. 7.

Fig. 7.

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Induction of KIR2DL4 transcription by DNA demethylation does not involve histone acetylation or methylation. Cells were cultured in the absence (shaded bars) or presence of 1 μM 5-Aza-dC (solid bars) for 72 h, at which time, KIR2DL4 was fully expressed in treated HUT78 T cells. ChIP assays were performed with antiacetyl-H3 antibody (A), antiacetyl-H4 antibody (B), antidimethyl-Lys 4 antibody (C), or antidimethyl-Lys 9 antibody (D); and KIR-specific sequences were amplified. DNA demethylation in HUT78 induced transcription without changes in histone acetylation and methylation patterns.

DISCUSSION

The current model of KIR expression suggests that NK cells have permissive histone conformations allowing KIR transcription in all NK cells including those that do not express a particular KIR gene. The clonal diversity of KIR expression is maintained exclusively by CpG island methylation [36, 44]. KIRs are also expressed on end-differentiated or senescent T cell subsets. Data presented here suggest that CpG motifs in the KIR2DL4 promoter are progressively demethylated with age in CD8 and in CD4+CD28− T cells. The promoter in Jurkat T cells is representative of that in young KIR− CD28+ T cells, is fully methylated, and is organized in a positioned nucleosome with the core promoter wrapped in the first nucleosome adjacent to the translation start site and inaccessible. Treatment with 5-Aza-dC induces only minimal demethylation and KIR2DL4 transcription. Inhibition of DNMT alone up-regulates KIR2DL4 transcription in the HUT78 cells to the maximal level seen in NK cells. T cells in elderly individuals behave much like HUT78, and the promoter in T cells from young adults is less responsive to DNMT inhibition. We identified two hallmarks that indicated whether a promoter is sensitive to DNMT inhibition. One characteristic finding is demethylation of CpG –24/–34 and –47 in HUT78 and CD4+CD28− T cells or CD8 T cells from the elderly. Dimethylation of H3-Lys4, as seen in HUT78 cells, appears to be a second early marker of promoter activation in T cells.

Post-translational histone modification is generally accepted as the major mechanism for regulating nucleosome stability [45, 46]. In particular, histone acetylation is thought to promote the disruption of nucleosomes prior to transcription initiation [47]. It was therefore unexpected that with the exception of H3-Lys 4, HUT78 exhibits the histone acetylation and methylation pattern typical of an inactive promoter, although the nucleosome was partially relaxed. Even when HUT78 fully transcribes KIR2DL4 after DNMT inhibition, it does so in the absence of and without the need for H3 and H4 histone acetylation or H3-Lys 9 demethylation. This finding is in contrast to the KIR2DL4 expression in the NK cell line NK3.3, which has histone modification typical for an active promoter.

Histone modification and DNA methylation are inter-related and dynamic events [48, 49]. Proteins that bind to methyl-CpG such as methyl CpG-binding proteins 1–4 not only recruit transcriptional repressor molecules but also modify the surrounding chromatin, providing a direct link between DNA methylation and chromatin remodeling [50, 51]. The sequence of events may vary; most examples indicate repressive histone modifications to be followed by DNA methylation [52]. However, in some cases, DNA methylation appears to be a primary event [52,53,54]. Recent studies suggested that histone signatures correlate with the competency to transcribe KIR genes. Chan et al. [13] showed that the inactive KIR promoter in NK cells has a relatively high level of acetylated H3 and H4 and trimethylated H3-Lys 4. Despite this, DNA methylation is maintained. Transcription initiation required DNA demethylation but no further histone modification. Santourlidis et al. [55] identified H4-Lys 8 acetylation in NK cells and CD8 T cells that have the potential to transcribe KIRs, and CD4 T cells and B cells were characterized by H3-Lys 9 dimethylation and therefore proposed that histone patterns confer the competence to transcribe KIRs.

In contrast, different epigenetic mechanisms were responsible to confer the competency to transcribe KIR2DL4 in the model system that we have used to examine the age-dependent KIR2DL4 expression in T cells. At one end of the spectrum are Jurkat T cells that do not transcribe KIR2DL4 and have a compact nucleosome with virtually no acetylated histones or di- or trimethylated H3-Lys 4. CpG motifs in the Jurkat promoter are largely methylated; DNMT inhibition only initiates marginal transcription. The lack of KIR2DL4 transcription after 5-Aza-dC treatment appeared to be a result of persistent CpG methylation at positions –24/–34. HUT78 is a T cell line mimicking end-differentiated CD4 T cells that have lost CD28 and gained KIR expression. The DNA methylation pattern of the KIR2DL4 promoter in HUT78 is similar to CD28− T cells and different from the biologically younger CD28+ T cells. T cells from elderly and HUT78 behave similarly to NK cells in that DNA methylation inhibition is all that is needed to induce full transcriptional activity. Nucleosome 1 of the core KIR2DL4 promoter is decondensed with increased accessibility and recruitment of transcription factors. However, in contrast to NK cells, histone modification changes were limited to an increase in dimethylated H3-Lys4 without any evidence for acetylated histones (Fig. 6). Moreover, this histone acetylation and methylation pattern did not change when DNMT inhibition induced full KIR2DL4 transcription (Fig. 7).

Promoter activation in the absence of histone acetylation is not unprecedented. Independence of histone acetyltransferases was suggested for the nucleosome remodeling at the IL-12 p40 promoter [56]. In preadipocytes, histone H3-Lys 4 dimethylation appeared to have a gatekeeper function for several genes that are eventually expressed in adipocytes such as apM1, glut4, gpd1, and leptin [57]. Promoter-associated H3-Lys 4 was found in undifferentiated cells with the potential to differentiate into adipocytes and apparently was involved in RNA polymerase II loading. Inhibition of H3-Lys 4 methylation by treatment with methylthioadenosine prevented adipocyte development. Different from HUT78, the beginning of apM1 transcription was associated with H3 hyperacetylation and H3-Lys 4 trimethylation [57], a pattern that we only found in fully developed NK cells.

Our data are consistent with the model that H3-Lys 4 methylation signals KIR promoter competence in T cells. Hazzalin and Mahadevan [58] have recently shown that Lys 4-methylated H3 is subject to continuous dynamic acetylation. Moreover, in their study of the c-fos and c-jun promoters, they have shown that hyperacetylation is associated with gene repression and not induction, suggesting that dynamic acetylation turnover and not high acetylation levels facilitate transcription [58]. Although we did not detect increased acetylated H3 in HUT78 compared with Jurkat cells, it is possible that the function of dimethylated H3-Lys 4 is to increase acetylation turnover.

KIR expression in T cells correlates highly with the loss of CD28 [17], which is a constitutively expressed receptor providing costimulatory signals critical for the productive activation of naïve and central memory T cells. Loss of CD28 expression occurs with age and is generally considered as a marker of end-differentiation or cellular senescence [23, 40, 59]. Jurkat and HUT78 differ in CD28 expression and have been considered as model systems for different stages of T cell differentiation. Also, KIR2DL3 and KIR2DS1 are expressed in HUT78 in addition to KIR2DL4 and not in Jurkat cells. Our data about primary cells give further support to the notion that the results from these two T cell lines allow for conclusion on the in vivo mechanism of T cell aging. Similar to HUT78, CD4+CD28− T cells and CD8 T cells from older individuals exhibited higher CpG demethylation and KIR2DL4 transcription levels than CD4+CD28+ T cells and CD8 T cells from young adults, and CD4+ and CD8+ T cells in the elderly individuals were more sensitive to DNMT inhibition than those in the young individuals.

Based on our findings, we propose the following model of KIR expression on T cells with aging. In unprimed T cells, the proximal promoter is condensed and inaccessible. Early enabling steps of transcription include targeted CpG demethylation and Lys 4 dimethylation of H3. At this stage, KIR expression may be low or absent, but DNMT inhibition is sufficient to induce maximal promoter activity. Further histone modification, including H3 and H4 acetylation, H3-Lys 4 trimethylation, and failure of DNMT recruitment to the promoter, is characteristic for stable transcription in NK cells but is not absolutely required for transcription in T cells, which with a partially competent KIR promoter, are already present in the young adult and increase in frequency with age. In the elderly, epigenetic regulation of KIRs in T cells is similar to NK cells and is highly sensitive to inhibition of DNMT activity. Elderly individuals may therefore be more susceptible to functional KIR expression during memory and effector T cell differentiation [22], where it negatively impacts effector T cell function.

Acknowledgments

This work was funded in part by grants from the National Institutes of Health (RO1 AR 41974, RO1 AR 42567, RO1 AG 15043, and UI9-AI 44142) and by a grant from the General Clinical Research Center (MO1 RR00039). The authors thank Tamela Yeargin for manuscript editing.

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