Gain-of-function CCR4 mutations in adult T cell leukemia/lymphoma

We performed RNA-seq of peripheral blood leukemia samples from two ATLL patients, TW36R and TW51R, which allowed us to identify 85 and 127 genes with potential coding region mutations in these two samples, respectively. These candidates included two genes that were mutated in both samples: CCR4 and MICALL1. High expression of CCR4 is a well-known hallmark of ATLL, whereas MICALL1 has not been implicated in this disease. Importantly, both ATLL samples had the same nonsense CCR4 mutation affecting the Y331 codon (Y331*), suggesting that CCR4 mutations might play a critical role in ATLL pathogenesis. The percentage of mutant CCR4 sequencing reads was 39% in TW36R and 55% in TW51R. Because both blood samples had a high proportion of malignant cells (TW36R: 90%, TW51R: 84%), the CCR4 mutations are likely to be heterozygous and potentially might exert a dominant functional effect.

By Sanger sequencing of genomic DNA, we confirmed the heterozygous nature of the CCR4 nonsense mutations in TW36R and TW51R. We extended this analysis to an additional cohort of ATLL primary patient samples (n = 41) and ATLL cell lines (n = 12). CCR4 mutations were detected in 26.4% (14/53) of ATLL samples (Fig. 1, A and B; and Table S1). In five cases for which paired normal DNA was available, three different CCR4 mutations were detected only in the ATLL cells (Q330*, Q330 frameshift, and Y331*), demonstrating that they were acquired somatically during malignant transformation or progression (Fig. 1 C). CCR4 mutations were identified in both primary ATLL patient samples (24%, 10/41) and in ATLL cell lines (33.3%, 4/12). Mutations were heterozygous in all samples except one cell line (ATL42T+). We also confirmed that mutant CCR4 mRNAs were heterozygously expressed in six primary ATLL samples (Table S1). These findings indicated that CCR4 mutations in ATLL might be either gain-of-function or potentially dominant negative.

All CCR4 mutations were either nonsense or frameshift in nature and affected the codons for four nearby amino acids (F326, C329, Q330, and Y331) that are located in an evolutionarily conserved region in the carboxy terminus of the protein (Fig. 1, A and B). The most frequent nonsense mutation affected codon Y331 (Y331*; 7/53, 13%). Four cell lines (ED40515(+), ED40515(−), ED41214(+), and ED41214 C(−)) had Q330* nonsense mutations, which is understandable given that these four lines were established from the same patient at different clinical time points.

In cancer genome studies cataloged in the COSMIC database (Catalogue of somatic mutations in cancer) or conducted by The Cancer Genome Atlas (TCGA) initiative, we found only three primary human cancer samples with CCR4 nonsense mutations, and these mutations did not target the carboxy-terminal cytoplasmic domain. Thus, truncation of the CCR4 cytoplasmic domain by somatic mutation appears to be a specific and frequent genetic event to ATLL.

Chemokine receptors, including CCR4, belong to the seven-transmembrane G protein–coupled receptor (GPCR) family. All CCR4 mutations in ATLL encode truncated receptors that lack most of the carboxy-terminal cytoplasmic domain, which typically serves a regulatory role in GPCRs (Fig. 1, A and B; Luttrell and Lefkowitz, 2002). The loss of this region of CCR4 might deregulate its activity, potentially altering the migration of ATLL cells in response to CCR4 ligands. The CCR4 mutations in ATLL are reminiscent of mutations affecting the chemokine receptor CXCR4 in WHIM syndrome, a human immunodeficiency disease (Diaz, 2005). Most CXCR4 mutations in WHIM syndrome are nonsense or frameshift mutations that truncate the carboxy-terminal cytoplasmic domain of the protein, conferring a gain-of-function phenotype with respect to chemotaxis toward the CXCR4 ligand SDF1 (Balabanian et al., 2005; Diaz, 2005; Kawai et al., 2005).

We therefore hypothesized that mutant CCR4 isoforms might enhance chemotaxis of the affected cells to CCR4 ligands. To study this, we infected a mouse myeloid cell line, 32Dβ, with retroviral vectors expressing WT CCR4 (CCR4-WT) or CCR4-Q330* coding regions and tested the migration of the transduced cells toward CCL22, the most potent CCR4 ligand (Fig. 2 A; D’Ambrosio et al., 2002). Surface CCR4 levels were similar between CCR4-WT– and CCR4-Q330*–transduced 32Dβ cells, whereas mock vector–transduced 32Dβ cells did not have detectable CCR4 expression (Fig. 2 B). Mock vector–infected 32Dβ cells did not migrate toward CCL22 (Fig. 2, A and B). CCR4-WT–transduced 32Dβ cells migrated with a typical bell-shaped dose–response curve (Fig. 2, A and B). Compared with CCR4-WT, CCR4-Q330*–transduced 32Dβ cells migrated to a significantly greater extent (Fig. 2, A and B). These results support the view that the CCR4 mutants in ATLL are gain-of-function with respect to ligand-directed chemotaxis.

Next we evaluated the ability of CCR4 isoforms to mediate chemotaxis in ATLL cell lines (Fig. 2 C). Most ATLL cell lines express CCR4 on their surface (not depicted), reflecting the high frequency (∼90%) of CCR4 expression in primary ATLL cases (Ishida et al., 2003). To evaluate CCR4-WT and CCR4-Q330* in ATLL cells, we first knocked down the endogenous expression of CCR4 in ED40515(+) cells using an shRNA targeting the 3′ untranslated region (UTR) of the CCR4 mRNA. This resulted in a >90% reduction of surface CCR4 expression (Fig. 2 D) and substantially decreased chemotaxis of the cells (Fig. 2 C). Cells were then transduced with retroviruses expressing CCR4-WT or CCR4-Q330* coding regions, resulting in cell populations with equivalent expression of CCR4 on the cell surface (Fig. 2 D). In chemotaxis assays with CCL17, CCR4-WT–reconstituted cells exhibited greater dose-dependent migration than mock vector–reconstituted cells, but migration of CCR4-Q330*–reconstituted cells was significantly greater than either of the other populations (Fig. 2 C, left). In response to CCL22, the most potent CCR4 ligand (D’Ambrosio et al., 2002), CCR4-Q330*–reconstituted ED40515(+) cells again displayed significantly greater chemotaxis than CCR4-WT–reconstituted cells and responded better to lower concentrations of the ligand (Fig. 2 C, right). Because CCR4 mutations in ATLL samples were heterozygous, we also analyzed whether these mutations could enhance chemotaxis in the presence of WT CCR4. To this end, we transduced the shCCR4-ED40515(+) line with one retroviral vector that coexpresses CCR4-WT and the human Kusabira-Orange (huKO) fluorescent protein and another vector that coexpresses CCR4-Q330 and GFP. This strategy allowed us to compare the phenotype of GFP/huKO double-positive cells that expressed CCR4-WT and Q330* with GFP or huKO single-positive cells that only ectopically expressed one CCR4 isoform (Fig. 2, E and F). Cells expressing both CCR4-WT and CCR4-Q330* showed greater CCL22-directed migration than cells expressing CCR4-WT alone, which was similar to the phenotype of cells expressing CCR4-Q330* alone (Fig. 2 F). These results suggest that ATLL cells may acquire CCR4 mutations to migrate more effectively toward their ligands.

The carboxy-terminal region of CCR4 that is truncated by mutations in ATLL contains a serine- and threonine-rich motif that is shared by many GPCRs (Fig. 1 A, amino acid positions 342–351). These serine and threonine residues become rapidly phosphorylated in response to ligand, which results in receptor internalization and contributes to “desensitization” of the cells to ligand (Luttrell and Lefkowitz, 2002). Carboxy-terminal truncation of CXCR4 in WHIM syndrome impairs receptor internalization, contributing to the enhanced migration of these cells in response to ligand (Balabanian et al., 2005; Kawai et al., 2005).

We therefore studied the change in surface CCR4 levels after CCL22 exposure in CCR4-WT– and CCR4-Q330*–reconstituted ED40515(+) cells. In CCR4-WT–reconstituted cells, surface CCR4 levels declined rapidly, with a 58% reduction at 5 min after CCL22 exposure and a maximum reduction of 75% at 20 min (Fig. 2 G). In comparison, CCR4 internalization in CCR4-Q330*–reconstituted cells was significantly impaired, with a 27% of reduction at 5 min and reaching only a 54% reduction at 20 min (Fig. 2 G). These results suggest that the ATLL CCR4 mutants impair desensitization by ligand, which likely contributes to the enhanced chemotaxis of cells bearing these mutants.

We explored the influence of the ATLL CCR4 mutants on PI3K–dependent activation of AKT because it has been reported that binding of CCL22 to CCR4 activates AKT in CEM leukemic T cells and in human Th2 cells (Cronshaw et al., 2004). We first studied two ATLL cell lines: ED40515(+), which bears a CCR4-Q330* mutant allele, and KOB, in which CCR4 is WT. Immunoblot analysis revealed that baseline levels of phospho-AKT (P-AKT), a measure of AKT activation, were much lower in ED40515(+) than in KOB (Fig. 3 A, lane 2 vs. lane 6). However, ED40515(+) showed stronger induction of AKT phosphorylation at 10 min after CCL22 exposure than KOB (Fig. 3 A, lane 3 vs. lane 7). The activation of AKT was transient in both cell lines, decreasing by 30 min after stimulation (Fig. 3 A, lanes 4 and 8). These findings indicated that AKT activation is one of the signaling pathways downstream of CCR4 in ATLL cells and suggested that CCR4 mutations might affect the magnitude of AKT activation. To accurately evaluate the relative ability of CCR4 isoforms to activate AKT, we studied ED40515(+) cells that were transduced with a CCR4 shRNA to knock down endogenous CCR4 expression and were reconstituted with CCR4-WT or CCR4-Q330*. Cells were treated with CCL22, and P-AKT levels were measured by FACS at various time points after exposure (Fig. 3 B). Knockdown of endogenous CCR4 reduced AKT activation by CCL22 compared with mock-infected cells, confirming that AKT activation was dependent on CCR4. Ectopic provision of CCR4-WT did not restore AKT activation despite expression of CCR4 at a higher level than in mock-transduced ED40515(+) cells (Fig. 2 D), presumably because the endogenous CCR4 locus in ED40515(+) encodes the CCR4-Q330* isoform. In contrast, cells reconstituted with CCR4-Q330* restored robust AKT activation in response to CCL22 (Fig. 3 B). In KOB cells, reconstitution with CCR4-WT modestly increased P-AKT activation in response to CCL22, but cells reconstituted with CCR4-Q330* again responded with a significantly greater rise in P-AKT levels (Fig. 3, C and D). The effect of heterozygous mutation of CCR4 on AKT activation was analyzed using the dual fluorescence strategy described above for experiments depicted in Fig. 2 (E and F). After CCL22 treatment, ED40515(+) cells expressing both CCR4-WT and CCR4-Q330* or CCR4-Q330* alone had elevated p-AKT levels compared with cells expressing only CCR4-WT (Fig. 3 E). Together, these data demonstrate that CCR4 mutations in ATLL enhance AKT activation in response to ligand engagement.

We next investigated whether AKT activation is involved in CCR4-mediated chemotaxis in ATLL. The pan-PI3K inhibitor BKM120 abrogated CCR4-mediated AKT activation in ED40515(+) cells in a dose-dependent manner, indicating that PI3K signaling contributes to AKT activation (Fig. 3 F). However, BKM120 treatment only partially inhibited CCR4-mediated chemotaxis by ED40515(+) cells (Fig. 3 G). In contrast, the G_αi inhibitor pertussis toxin (PTX) abrogated AKT activation and chemotaxis in these cells (Fig. 3, F and G). Thus, CCR4-mediated chemotaxis of ATLL cells is not primarily caused by PI3K-dependent AKT signaling, but instead depends on G_αi, consistent with previous work (Cronshaw et al., 2004).

Lastly, we tested whether the acquisition of CCR4 mutations by ATLL cells imparts a selective growth advantage relative to cells with WT CCR4. After shRNA-mediated knockdown of endogenous CCR4 expression, ED40515(+) cells were transduced with a retrovirus expressing CCR4-WT together with GFP or with a retrovirus expressing only CCR4-Q330* (Fig. 4 A, Exp. 1). After puromycin selection of infected cells, these two cell populations were mixed in equal numbers and cultured with or without CCL22 for 12 d. The ratio of GFP⁻/CCR4-Q330*–expressing cells versus GFP⁺/CCR4-WT–expressing cells was monitored every 4 d by FACS (Fig. 4 B, Exp. 1). Without CCL22, the ratio of the two populations did not change. In contrast, in the presence of CCL22, CCR4-Q330*–reconstituted cells preferentially expanded in a time-dependent manner. To confirm this finding, we performed a GFP-swapping experiment in which the GFP⁺ cells expressed CCR4-Q330* and the GFP⁻ cells expressed CCR4-WT (Fig. 4 A, Exp. 2). Again, CCR4-Q330*–reconstituted cells had a selective growth advantage in the presence of CCL22 (Fig. 4 B, Exp. 2). As a negative control, we used empty vectors to create GFP⁺ and GFP⁻ populations and observed no change in the GFP⁺/GFP⁻ ratio in the presence or absence of CCL22 (Fig. 4, A and B, Exp. 3).

In conclusion, the present study demonstrates that mutations in CCR4 are a frequent genetic event in ATLL and provides a mechanistic rationale for their selection in this cancer. Mutant CCR4 isoforms enhanced migration of ATLL cells toward CCL17 and CCL22 and additionally promoted PI3K/AKT activation in response to ligand engagement, leading us to propose two nonmutually exclusive hypotheses regarding the role of CCR4 mutations in ATLL pathogenesis. First, enhanced migration of ATLL cells along a CCL17/CCL22 gradient might allow them to access a favorable microenvironmental niche that could support their proliferation and/or survival. Lymph nodes and skin are plausible ATLL niches because CCL17 and CCL22 are known to be produced by dendritic cells and M2-phenotype macrophages in lymph nodes and also by Langerhans cells and venules in skin (Campbell et al., 1999; Vissers et al., 2001; Vulcano et al., 2001; Chong et al., 2004). Previous studies showed that monocyte and M2-phenotype macrophages can promote ATLL cell growth through cell–cell interaction and by paracrine mechanisms (Chen et al., 2010; Komohara et al., 2013). Thus, the chemotaxis promoted by CCR4 mutations in ATLL might set up favorable interactions between ATLL cells and immune bystander cells and/or disrupt homeostatic mechanisms that keep the growth of these cells in check. Recently, WHIM syndrome-like CXCR4 somatic mutations have been detected in the malignant cells of patients with Waldenström’s macroglobulinemia, and these mutations correlated with bone marrow involvement as well as chemotherapy drug resistance (Roccaro et al., 2014; Treon et al., 2014). Given that CCR4 mutations and WHIM syndrome–like CXCR4 mutations share similar biological properties in terms of chemotaxis and ATK activation (Cao et al., 2014; Treon et al., 2014), it will be interesting to determine whether CCR4 mutations affect the frequency of lymph node or skin involvement in ATLL or the clinical course.

A second hypothesis raised by our findings is that CCR4 mutations may be selected in ATLL for their ability to enhance signaling downstream of this chemokine receptor, including activation of the PI3K/AKT signaling pathway. During physiological CCR4 signaling, ligand desensitization occurs in part because CCR4 is down-modulated from the cell surface, a process which is disrupted by the CCR4 mutations in ATLL. Perhaps as a result, ATLL cells bearing CCR4 mutant isoforms displayed prolonged PI3K/AKT activation in response to ligand. Given the importance of PI3K/AKT signaling to both cellular metabolism and survival, this enhanced PI3K/AKT response might provide a selective advantage for ATLL cells. Indeed, in our long-term competitive growth assay, ATLL cells expressing mutant CCR4 outgrew cells with WT CCR4 in the presence of CCR4 ligand. Finally, it is conceivable that both hypotheses detailed above may pertain. Specifically, the ability of CCR4 mutants to increase chemotaxis toward CCR4 ligands would expose them to higher ligand concentrations, which might contribute to their growth and/or survival.

Our findings provide a rationale to test whether inhibition of CCR4 signaling might have therapeutic potential for patients with ATLL. Either a CCR4 inhibitor or a PI3K inhibitor might be considered (Pease and Horuk, 2014). The anti-CCR4 monoclonal antibody KW-0761, which is showing promising results in clinical trials (Yamamoto et al., 2010; Ishida et al., 2012), was designed to promote antibody-dependent cellular cytotoxicity of ATLL cells. This antibody only inhibits chemotaxis weakly (Ishida et al., 2006), and its effect on PI3K/AKT signaling has not been evaluated. To improve the therapy of ATLL, our findings would support the development of a therapeutic anti-CCR4 antibody that both inhibits CCR4 signaling and mediates antibody-dependent cellular cytotoxicity.

All experiments presented have been repeated at least two times, and consistent results were obtained. Data are depicted as means ± SEM. Statistical comparisons were made using the Student’s t test. P < 0.05 was considered statistically significant.

Written informed consent was obtained in accordance with the Declaration of Helsinki and was approved by the Investigational Review Board of the National Cancer Institute (NCI). Some samples were obtained before cytotoxic chemotherapy, whereas others were taken after treatment (Table S1). PBMCs were isolated from ATLL patients by Ficoll-Hypaque. In cases where the ATLL cells were <70% of the mononuclear cells, the ATLL cells were purified by an initial negative selection magnetic column method (Miltenyi Biotec) to enrich for CD4⁺ cells followed by a positive selection on a CD25 column. The resultant population consisted of 70–95% CD4⁺CD25⁺ ATLL cells. The ATLL cell lines were provided by the following researchers: M. Maeda (Kyoto University, Kyoto, Japan; ED40515(+), ED40515(−), ED41214(+), ED41214C(−), ATL43T(+), ATL43Tb(−), ATL55T(+), and ATL42T(+)), Y. Yamada (Nagasaki University, Nagasaki, Japan; ST1, KOB, KK1, and LMY1), T. Hata (Nagasaki University; ST1), T. Naoe (Nagoya University, Nagoya, Japan; ATN1), and N. Arima (Kagoshima University, Kagoshima, Japan; Su9T01). 32Dβ (a variant of the mouse 32D myeloid cell line engineered to express the human IL-2 receptor β subunit) was also used. IL2-dependent cell lines (ED40515(+), ED41214(+), ATL43T(+), ATL55(+), ATL42T(+), KOB, KK, LMY1, and 32Dβ) were cultured with RPMI/10% FCS/penicillin-streptomycin with 100 IU/ml human recombinant IL2. Other lines were cultured with RPMI/10% FCS/penicillin-streptomycin. ED40515(+) and KOB were engineered to express ecotropic retroviral receptors and TET repressor for transduction of retroviral shRNA vectors and expression vectors as previously described (Schmitz et al., 2012).

Doxycycline, puromycin, ethidium bromide, and Lipofectamine 2000 were purchased from Invitrogen. Recombinant human CCL17/TARC and recombinant human CCL22/MDC were purchased from R&D Systems. Human recombinant IL2 was obtained from Roche. Anti–human CCR4 antibodies (clone 1G1) conjugated to PE and Alexa Fluor 647 fluors were purchased from BD. The following antibodies were purchased from Cell Signaling Technology: Akt, P-Akt (Ser473; D9E), P-Akt (S473)–Alexa Fluor 647 (D9E), and Alexa Fluor 647–conjugated isotype control antibody (DA1E). Anti–rabbit IgG-HRP antibody was purchased from GE Healthcare. CD8a(Lyt 2) microbeads were purchased from Miltenyi Biotec. BKM120 was purchased from Selleck Chemicals. PTX was purchased from Sigma-Aldrich. Big Dye Terminator v1.1 was purchased from Applied Biosystems. Human gamma globulin fraction II was purchased from ICN Biomedicals Inc. Paraformaldehyde was purchased from Electron Microscopy Sciences.

RNA was extracted using the AllPrep kit (QIAGEN) and sequencing libraries were prepared using the TruSeq RNA sample Prep kit v2 (Illumina) according to the manufacturer’s instructions. Paired-end 108-bp read sequencing was performed on a HiSeq 2000 system (Illumina). The mapping of paired-end reads and the extraction of putative single nucleotide variants (SNVs) were performed as previously described (Schmitz et al., 2012). In brief, paired-end reads were mapped to the RNA sequences in the RefSeq database (NCBI build 37) using the Burrows-Wheeler Aligner (BWA) software with default parameters. Reads that failed to map to RefSeq were mapped to RNA sequences in the Ensembl database. The remaining unmapped reads were mapped to the human genome assembly (NCBI build 37). Mutant SNV calls were declared if more than two reads were mutated and the ratio of mutant reads versus total coverage was >20%. SNVs that corresponded to single nucleotide polymorphisms in the dbSNP database (build #132), the 1000 Genomes database (May 2011 release), and the NHLBI GO Exome Sequencing Project (ESP) database (ESP5400 December 2011 release) were excluded. RNA-seq data were submitted to the NCBI Short Read Archive (SRA; accession no. SRP042199).

Genomic DNA of CCR4 exon 2 region was amplified by PCR using the primers CCR4-E2F, 5′-CTTCCCCTCATTAGCTGCTTCTGGTTG-3′; and CCR4-E2R, 5′-CCTGACACTGGCTCAGGAATCTCTTAC-3′. The purified PCR products were sequenced using a Big Dye Terminator v1.1 cycle sequencing kit and analyzed on an ABI 3730 Genetic Analyzer (Applied Biosystems). The following primers were used for sequencing: CCR4-E2.1F, 5′-CCTTCCTGGCTTTCTGTTCAGCACTTG-3′; and CCR4-E2.1R, 5′-TGATTTCCAGGGAGCTGAGAACCTTCC-3′.

RNA was DNase-digested and reverse transcribed using the Omniscript RT kit (QIAGEN). The CCR4 coding region was amplified by PCR using primers CCR4-E2F and CCR4-E2R. The purified PCR products were cloned using the TOPO XL PCR Cloning kit (Invitrogen). Sequencing was performed as described in the Sanger DNA resequencing section.

cBioPortal and COSMIC were used.

CCR4-WT and CCR4-Q330* cDNA was PCR amplified from ED40515(+) cDNA using the primers HindIII-CCR4-S, 5′-GGAAGCTTTTGAAAGGCACCGGGTC-3′; and CCR4-TAG-BamHI-AS, 5′-CGGGATCCCTACAGAGCATCATGGAGAT-3′. The amplified fragments were cloned into HindII–BamHI sites of doxycycline-inducible pRetroCMV/TO/puro and pRetroCMV/TO/PG vectors. MSCV-CCR4-WT-ires-huKO or MSCV-CCR4-Q330*-ires-GFP was made by inserting HindIII (blunted)–XhoI fragments from pRetroCMV/TO/CCR4-WT-puro or pRetroCMV/TO/CCR4-Q330*-puro vectors into EcoRI(blunted)–XhoI sites of MSCV-ires-huKO or MSCV-ires-GFP (provided by S. Tsuzuki, Aichi Cancer Center Research Institute, Nagoya, Japan). shRNA targeting the 3′ UTR of CCR4 was cloned into doxycycline-inducible pRSMX-puro vector with these two annealed oligos (shRNA target sequences are underlined): shCCR4-4A11_F, 5′-GATCCCGAATGAAGTTGTAGGTAATTTCAAGAGAATTACCTACAACTTCATTCTTTTT-3′; and shCCR4-4A11_R, 5′-AGCTAAAAAGAATGAAGTTGTAGGTAATTCTCTTGAAATTACCTACAACTTCATTCGG-3′. pBMN-shCCR4-ires-Lyt2 was made by insertion of XbaI–BsmI fragment from pRSMX-shCCR4-puro vector into XbaI–BsmI sites of pBMN-ires-Lyt2 (provided by G. Nolan, Stanford University, Stanford, CA). Retroviruses for ED40515(+) and KOB were produced by cotransfection of 293T cells with helper plasmids expressing gag-pol genes and an ecotropic pseudotyping env gene and a retroviral vector using Lipofectamine 2000 as described previously (Schmitz et al., 2012). For 32Dβ, retroviruses were produced by transfection of PlatE with retroviral vectors. Target cells were infected with the retroviruses with 8 µg/ml polybrene with spin infection at 1,290 g for 1.5 h. pRetroCMV/TO-puro–, pRetroCMV/TO-PG–, and pRSMX-infected cells was purified by using 2 µg/ml puromycin. pBMN-ires-Lyt2–infected cells were purified by MACS mouse CD8a (Lyt2) microbeads (Miltenyi Biotech). Doxycycline was used at 40 ng/ml for induction of shRNA and cDNA.

HTS Transwell 96-well permeable supports with an 8.0-µm pore polyester membrane (Corning) were used for chemotaxis assays of ED40515(+) and KOB. 235 µl of culture medium containing the indicated dose of chemokine was added to the lower chamber, followed by the addition of the Transwell inserts and the addition of cells in growth medium (75 µl) at a concentration of 5 × 10⁵ cells/ml into the upper chamber. ChemoTx (5 µM pore; Neuro Probe) was used for chemotaxis assays of 32Dβ. 300 µl of chemokine-containing medium was added to the lower chamber, and 55 µl of cells in growth media (2 × 10⁶ cells/ml) were added to the upper chamber. After incubation for 2 h at 37°C in 5% CO₂, the upper chamber was removed and 50 µl of 1× PBS with a 1:100 dilution of SPHERO particles (SpheroTECH) and 1:2,500 dilution of 10 mg/ml ethidium bromide was added into each lower chamber. The numbers of cells and SPHERO particles in lower chamber were quantified with a FACSCalibur (BD), and the numbers of cells that had migrated in each condition were normalized based on the SPHERO particle count.

Cells were washed with PBS containing 200 µg/ml of human gamma globulin (Human Gamma Globulin Fraction II; ICN Biomedicals Inc.) twice and incubated for 30 min on ice with anti–human CCR4-PE or CCR4–Alexa Fluor 647 (1G1; BD) or control isotype IgG. After washing with FACS buffer (PBS/1% FCS) twice, the cells were analyzed on a FACSCalibur (BD). For the CCR4 internalization assay, cells were exposed to CCL22 for varying lengths of time and were then treated for 1 min at 37°C to an acid buffer (pH 3.0, consisting of 50 mM glycine and 100 mM NaCl) to remove cell-bound CCL22. The cells were washed twice and then stained with anti–human CCR4–Alexa Fluor 647 on ice for 30 min, fixed with 2% paraformaldehyde (Electron Microscopy Sciences) on ice for 10 min, and analyzed on a FACSCalibur.

Cells were washed and resuspended in 2× SDS sample buffer at 10⁶ cells/100 µl and boiled for 5 min. Samples were separated on Novex 4–12% Tris-Glycine gel (Invitrogen) and transferred to a PVDF membrane (Immobilon-P; EMD Millipore). Proteins were detected with the following primary antibodies: p-AKT(S473) (D9E) and AKT (5G3; Cell Signaling Technology). An anti–rabbit IgG-HRP antibody (GE Healthcare) was used as the secondary antibody.

After incubation with 50 ng/ml CCL22 for the indicated time, cells were fixed in 2% paraformaldehyde for 10 min at room temperature and then permeabilized with cold methanol at −20°C overnight. Cells were washed twice in FACS buffer and stained with p-Akt (S473)–Alexa Fluor 647 (D9E) or isotype control (DA1E; Cell Signaling Technology) for 20 min at room temperature. Cells were washed and resuspended in FACS buffer for analysis on a FACSCalibur.

Table S1, included as a separate Excel file, shows Sanger sequencing analysis of exon 2 of CCR4 (NM_005508.4) in 53 cases of ATLL samples.

We thank the following researchers for providing ATLL cell lines: Michiyuki Maeda, Yasuaki Yamada, Tomoko Hata, Tomoki Naoe, and Naomichi Arima. We also thank Richard N. Bamford, Liyanage P. Perera, Ryan Young, Art Shaffer, and Michele Ceribelli for helpful discussions.

This research was supported by the Intramural Research Program of the National Institutes of Health, National Cancer Institute, Center for Cancer Research. Roland Schmitz was supported by the Dr. Mildred Scheel Stiftung für Krebsforschung (Deutsche Krebshilfe).

The authors declare no competing financial interests.